Dynamics of Bone and Cartilage Metabolism, Second Edition: Principles and Clinical Applications

Dynamics of Bone and Cartilage Metabolism

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Dynamics of Bone and Cartilage Metabolism Edited by MARKUS J. SEIBEL Department of Medicine Division of Endocrinology and Metabolism University of Heidelberg Medical School Heidelberg, Germany

SIMON P. ROBINS Skeletal Research Unit Rowett Research Institute Aberdeen, United Kingdom

JOHN P. BILEZIKIAN Departments of Medicine and Pharmacology Division of Endocrinology College of Physicians and Surgeons Columbia University New York, New York

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Academic Press is an imprint of Elsevier 30 Corporate Drive, Suite 400, Burlington, MA 01803, USA 525 B Street, Suite 1900, San Diego, California 92101-4495, USA 84 Theobald’s Road, London WC1X 8RR, UK This book is printed on acid-free paper. Copyright © 2006, Elsevier Inc. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the publisher. Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone: (+44) 1865 843830, fax: (+44) 1865 853333, E-mail: [email protected]. You may also complete your request on-line via the Elsevier homepage (http://elsevier.com), by selecting “Support & Contact” then “Copyright and Permission” and then “Obtaining Permissions.” Library of Congress Cataloging-in-Publication Data Dynamics of bone and cartilage metabolism / Markus J. Seibel, Simon P. Robins, and John P. Bilezikian, editors.— 2nd ed. p. ; cm. ISBN-13: 978-0-12-088562-6 (hardcover : alk. paper) ISBN-10: 0-12-088562-X (hardcover : alk. paper) 1. Bones—Metabolism. 2. Cartilage—Metabolism. 3. Extracellular matrix. 4. Bones—Metabolism—Disorders. 5. Biochemical markers. [DNLM: 1. Bone and Bones—metabolism. 2. Biological Markers. 3. Cartilage—metabolism. 4. Extracellular Matrix—metabolism. WE 200 D997 2006] I. Seibel, M. J. II. Robins, Simon P. III. Bilezikian, John P. QP88.2.D96 2006 612.7’5—dc22 2006004804 British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library. ISBN 13: 978-0-12-088562-6 ISBN 10: 0-12-088562-X For information on all Academic Press publications visit our Web site at www.books.elsevier.com Printed in the United States of America 06 07 08 09 10 9 8 7 6 5

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Contents Contributors xiii Preface xix

IV. Concluding Remarks References 50

CHAPTER 3 Vitamin K Dependent Proteins of Bone and Cartilage

PART I Components of the Organic Extracellular Matrix of Bone and Cartilage

CAREN M. GUNDBERG AND SATORU K. NISHIMOTO I. Abstract 55 II. Introduction 55 III. Osteocalcin 56 IV. Matrix Gla Protein 59 V. Gas6 63 VI. Vitamin K/Warfarin 64 References 65

CHAPTER 1 Structure, Biosynthesis and Gene Regulation of Collagens in Cartilage and Bone I. II. III. IV. V. VI. VII. VIII.

KLAUS VON DER MARK Introduction 3 The Collagen Families 5 Bone Collagens 7 Cartilage Collagens 9 Collagen Biosynthesis 14 Collagen Genes and Transcriptional Regulation 17 Factors Regulating Collagen Biosynthesis Conclusions 25 References 26

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CHAPTER 4 Noncollagenous Proteins; Glycoproteins and Related Proteins DICK HEINEGÅRD, PILAR LORENZO, AND TORE SAXNE I. Introduction 71 II. Cartilage Extracellular Matrix 72 III. Bone, Extracellular Matrix 77 IV. Concluding Remarks 79 References 80

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CHAPTER 5 Proteoglycans and Glycosaminoglycans

CHAPTER 2 Fibrillogenesis and Maturation of Collagens

I. II. III. IV.

SIMON P. ROBINS I. Introduction 41 II. Fibrillogenesis 42 III. Cross-linking 44 v

TIM HARDINGHAM Introduction 85 Glycosaminoglycans 86 Proteoglycans in Cartilage 87 Aggrecan 88

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V. Leucine-rich Proteoglycans in Cartilage and Bone 93 VI. Perlecan in Cartilage 96 References 96

CHAPTER 6 I. II. III. IV. V. VI. VII.

Growth Factors

PHILIPPA HULLEY, GRAHAM RUSSELL, AND PETER CROUCHER Introduction 99 Insulin-like Growth Factors 99 The Transforming Growth Factor-Beta/Bone Morphogenetic Protein Superfamily 100 Fibroblast Growth Factors 103 Wnts 104 Additional Growth Factors 105 Summary 106 References 107

CHAPTER 10

CHAPTER 11 I. II. III. IV. V.

CHAPTER 7 Prostaglandins and Proinflammatory Cytokines I. II. III. IV.

I. II. III. IV. V. VI. VII.

Acid Phosphatases

HELENA KAIJA, LILA O.T. PATRIKAINEN, SARI L. ALATALO, H. KALERVO VÄÄNÄNEN, AND PIRKKO T. VIHKO I. Acid Phosphatases 165 II. Tartrate-Resistant Acid Phosphatase (TRACP) 166 III. Prostatic Acid Phosphatase (PAP) 173 References 176

Matrix Proteinases

IAN M. CLARK AND GILLIAN MURPHY Introduction 181 Aspartic Proteinases 181 Cysteine Proteinases 182 Serine Proteinases 184 Metalloproteinases 186 References 192

LAWRENCE G. RAISZ AND JOSEPH A. LORENZO Introduction 115 Prostaglandins 115 The Role that Cytokines have in Osteoclast Formation and Function 117 The Role that Proinflammatory Cytokines have in Bone and Cartilage Metabolism 118 References 121

PART II Structure and Metabolism of the Extracellular Matrix of Bone and Cartilage

CHAPTER 8 Integrins and Other Adhesion Molecules

I. II. III. IV.

M. H. HELFRICH AND M. A. HORTON Abstract 129 Introduction 129 Molecular Structure of Adhesion Molecules 130 Adhesion Molecules in Cells of the Osteoblast Lineage 134 Adhesion Molecules in Osteoclasts 139 Adhesion Molecules in Chondrocytes 142 Conclusion 144 References 144

CHAPTER 9

Alkaline Phosphatases

JOSÉ LUIS MILLÁN I. Introduction 153 II. Structure And Regulation of the TNAP Gene 153 III. Protein Structure 155 IV. Function of TNAP 157 V. Clinical Use 160 References 161

CHAPTER 12 Mineralization, Structure and Function of Bone ADELE L. BOSKEY Abstract 201 The Structure and Function of Bone 201 Bone Mineralization 204 Bone Modeling and Remodeling 209 References 209

CHAPTER 13 Bone Structure and Strength EGO SEEMAN I. Introduction 213 II. Gravity and the Need for Stiffness, Flexibility, Lightness and Speed 213 III. The Material Composition and Structural Design of Bone 214 IV. Bone Modeling and Remodeling – The Mechanism of Bone’s Construction during Growth and Decay with Advancing Age 216 V. Strength Maintenance 217 VI. Conclusion 219 References 219

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CHAPTER 14 I. II. III. IV. V.

VI.

JANE B. LIAN AND GARY S. STEIN Abstract 221 Introduction 221 Developmental Signals for Cartilage and Bone Tissue Formation 222 Osteogenic Lineage Cells 227 The Osteoclast: A Functionally Unique Cell for Physiologically Regulated Resorption of Bone Mineral 236 Perspectives 242 References 242

CHAPTER 15 I. II. III. IV. V. VI.

VII. VIII. IX.

X.

XI.

The Cells of Bone

Signaling in Bone

T. JOHN MARTIN AND NATALIE A. SIMS Abstract 259 Introduction 259 The Control of Osteoclasts 260 Signaling in the Control of Osteoclast Activity 261 Signals from the Osteoblast Lineage that Control Osteoclast Formation 261 Hormone and Cytokine Influences on the Contact-dependent Regulation of Osteoclasts 261 Discovery of the Physiological Signaling Mechanisms in Osteoclast Control 262 Rank Signaling 263 Coupling of Bone Formation to Resorption – Release of Growth Factors from bone Matrix 264 Coupling of Bone Formation to Resorption – Autocrine/Paracrine Regulation by Differentiating Osteoblasts 266 Coupling of Bone Formation to Resorption – Are Osteoclasts a Source of Coupling Activity? 266 References 267

IX. Activation of the Cyclic Adenosine Monophosphate Second-Messenger System by Parathyroid Hormone 281 X. Identification of a Second PTH Receptor 282 XI. Physiological Actions of PTH 282 XII. Cell-to-Cell Communication: Osteoblasts and Osteoclasts 284 XIII. Preferential Actions of PTH at Selected Skeletal Sites 285 References 287

CHAPTER 17 Interaction of Parathyroid Hormone-related Peptide with the Skeleton I. II. III. IV. V.

CHAPTER 18 The Vitamin D Hormone and its Nuclear Receptor: Mechanisms Involved in Bone Biology GEERT CARMELIET, ANNEMIEKE VERSTUYF, CHRISTA MAES, GUY EELEN, AND ROGER BOUILLON I. Introduction 307 II. Metabolism of Vitamin D 308 III. Nuclear Vitamin D Receptor 309 IV. Vitamin D and Bone Cells 312 V. Pathology and Therapy Related to Vitamin D Availability, Metabolism, and Function 314 VI. Conclusions 319 References 319

CHAPTER 19 Sex Steroid Effects on Bone Metabolism

CHAPTER 16 Parathyroid Hormone: Structure, Function and Dynamic Actions LORRAINE A. FITZPATRICK AND JOHN P. BILEZIKIAN I. Introduction 273 II. Structure of the PTH Gene 273 III. Chromosome Location 274 IV. Control of Gene Expression 275 V. Biosynthesis of Parathyroid Hormone 278 VI. Metabolism of Parathyroid Hormone 278 VII. Receptor Interactions of Parathyroid Hormone and Parathyroid Hormone-Related Protein 279 VIII. Structure of the PTH/PTHrP (PTH1R) Receptor 279

DAVID GOLTZMAN Abstract 293 Introduction 293 Molecular Biology and Mechanism of Action 294 The Skeletal Actions of PTHrP 298 Summary 301 References 302

I. II. III.

IV. V.

DAVID G. MONROE, THOMAS C. SPELSBERG, AND S. KHOSLA Abstract 327 Introduction 327 Molecular Structures, Synthesis, Mechanism of Action of Major Sex Steroids, and Transcriptional Coregulator Function 328 Effects of Sex Steroids on Bone Cells and Bone Turnover 331 Effects of Estrogens and Androgens on Bone Metabolism in Men Versus Women 335

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VI. Effects of Sex Steroids on Extraskeletal Calcium Homeostasis 336 VII. Summary 337 References 338

CHAPTER 20 Physiology of Calcium and Phosphate Homeostases I. II. III. IV. V.

VI. VII. VIII. IX. X. XI.

RENÉ RIZZOLI AND JEAN-PHILIPPE BONJOUR Abstract 345 Introduction 345 Body Distribution of Calcium 346 Determinants of Extracellular Calcium Concentration 346 Relative Importance of the Various Calcium Fluxes in Controlling Extracellular Calcium Homeostasis 349 Homeostatic Responses to Hypocalcemia 349 Calcium and Bone Growth 352 Body Distribution of Phosphorus 353 Determinants of Extracellular Phosphate Concentration 353 Homeostatic Responses to Changes in Phosphate Supply or Demand 356 Conclusions 357 References 357

CHAPTER 21 The Central Control of Bone Remodeling PAUL A. BALDOCK, SUSAN J. ALLISON, HERBERT HERZOG, AND EDITH M. GARDINER I. Introduction 361 II. Actions of Leptin 362 III. Sympathetic Nervous System 365 IV. Neuropeptide Y and the Y Receptors 369 V. Interaction between Leptin and Y2-Regulated Bone Antiosteogenic Pathways 371 VI. Concluding Remarks 373 References 373

CHAPTER 22 New Concepts in Bone Remodeling I. II. III. IV. V.

DAVID W. DEMPSTER AND HUA ZHOU Introduction 377 An Overview of the Remodeling Cycle 377 Functions of Bone Remodeling 380 The Role of Apoptosis in Regulating Bone Balance 381 Possible Mechanisms whereby a Reduction in Activation Frequency may Protect against Fracture 383 References 386

CHAPTER 23

Products of Bone Collagen Metabolism

JUHA RISTELI AND LEILA RISTELI I. Introduction 391 II. Products of Bone Collagen Synthesis, the Procollagen Propeptides 392 III. Degradation Products of Type I Collagen IV. Closing Remarks 402 References 403

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CHAPTER 24 Supramolecular Structure of Cartilage Matrix I. II. III. IV. V. VI. VII. VIII. IX. X. XI. XII.

PETER BRUCKNER Summary 407 Introduction 407 Light and Electron Micrography 408 Biochemistry of Cartilage 409 Studies of Fibril Structures by X-ray Diffraction 411 Structure of Fibril Fragments Obtained by Mechanical Disruption of Tissue 411 Studies of Collagen Cross-linking in Cartilage Fibrils 412 Reconstitution of Aggregates from Soluble Collagens and other Macromolecules 412 Studies of Transgenic Mice and of Human Genetic Matrix Diseases 414 Correlating Structure With the Biomechanical Role of Articular Cartilage 415 Models of Cartilage Fibril Structure 416 Future Perspectives 417 References 418

CHAPTER 25

Products of Cartilage Metabolism

DANIEL-HENRI MANICOURT, JEAN-PIERRE DEVOGELAER, AND EUGENE J.-M. A. THONAR I. Introduction 421 II. The Chondrocyte and its Extracellular Matrix 422 III. Products of Collagen Metabolism 424 IV. Products of Aggrecan Metabolism 429 V. Products of the Metabolism of other Proteoglycans 434 VI. Products of the Metabolism of Link Protein and Hyaluronan 435 VII. Other Products of Chondrocyte Metabolism 437 VIII. Concluding Statement 438 References 439

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CHAPTER 26 Fluid Dynamics of the Joint Space and Trafficking of Matrix Products PETER A. SIMKIN I. Introduction 451 II. Interpretation of Marker Data and Strategies for Dealing with Them 451 III. Conclusion 456 References 456

CHAPTER 27 Transgenic Models of Bone Disease I. II. III. IV.

BARBARA E. KREAM AND JOHN R. HARRISON Introduction 457 Generation of Mouse Models 457 Transgenic Models in Bone Biology 460 Perspectives and Future Directions 465 References 465

PART III Markers of Bone and Cartilage Metabolism

CHAPTER 30 Measurement of Parathyroid Hormone HARALD JÜPPNER AND GHADA EL-HAJJ FULEIHAN I. Abstract 507 II. Background 507 III. Different Immunometric Assays for the Detection of PTH 509 IV. Primary Hyperparathyroidism 510 V. Hyperparathyroidism in Renal Osteodystrophy 510 VI. Pseudohypoparathyroidism (PHP) 510 VII. Conclusion 511 References 511

CHAPTER 31 New Horizons for Assessment of Vitamin D Status in Man GARY L. LENSMEYER, NEIL BINKLEY, AND MARC K. DREZNER I. Measurement of Vitamin D3 (Cholecalciferol) and Vitamin D2 (Ergocalciferol) 513 II. Measurement of 25-Hydroxyvitamin D 513 III. Measurement of 1,25-Dihydroxyvitamin D 524 References 525

CHAPTER 28 The Role of Genetic Variation in Osteoporosis ANDRÉ G. UITTERLINDEN, JOYCE B.J. VAN MEURS, FERNANDO RIVADENEIRA, JOHANNES P.T.M.VAN LEEUWEN, AND HUIBERT A.P. POLS I. Abstract 471 II. Osteoporosis has Genetic Influences 471 III. Genome-wide Approaches to Find the Genes 473 IV. Association Analysis of Candidate Gene Polymorphisms 476 V. Haplotypes 478 VI. Meta-analyses 480 VII. Osteoporosis Candidate Genes: Collagen Type Iα1 and the Vitamin D Receptor 482 VIII. Summary 483 References 484

CHAPTER 29 Measurement of Calcium, Phosphate and Magnesium HEINRICH SCHMIDT-GAYK I. Measurement of Calcium 487 II. Measurement of Phosphate 495 III. Measurement of Magnesium 499 References 502

CHAPTER 32 Measurement of Biochemical Markers of Bone Formation I. II. III. IV. V. VI. VII.

KIM E. NAYLOR AND RICHARD EASTELL Abstract 529 Introduction 529 Propeptides of Type I Procollagen 530 Total Alkaline Phosphatase 533 Bone Alkaline Phosphatase 533 Osteocalcin 535 Discussion 537 References 537

CHAPTER 33 Measurement of Biochemical Markers of Bone Resorption MARIUS E. KRAENZLIN AND MARKUS J. SEIBEL I. Introduction 541 II. Collagen Related Markers 543 III. Non-Collagenous Proteins of the Bone Matrix 553 IV. Osteclast Enzymes 555 References 557

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CHAPTER 34 Variability in the Measurement of Biochemical Markers of Bone Turnover

CHAPTER 38 Monitoring Anabolic Treatment

TUAN V. NGUYEN, CHRISTIAN MEIER, AND MARKUS J. SEIBEL Introduction 565 Sources of Pre-Analytical Variability in the Measurement of Biochemical Markers of Bone Turnover 566 Statistical Consideration of Variability 571 Summary 577 References 578

JOHN P. BILEZIKIAN AND MISHAELA R. RUBIN I. Introduction 629 II. Cellular and Regulatory Mechanisms of the Anabolic Actions of Parathyroid Hormone 630 III. Pharmacokinetics of Teriparatide in Human Subjects 631 IV. Actions of Parathyroid Hormone to Improve Bone Quality 632 V. Conclusions 642 References 642

CHAPTER 35 Validation of Biochemical Markers of Bone Turnover

CHAPTER 39 Monitoring of Antiresorptive Therapy

I. II.

III. IV.

I. II. III. IV.

KIM BRIXEN AND ERIK FINK ERIKSEN Introduction 583 Validation of Biochemical Markers by Calcium Kinetics 584 Validation of Biochemical Markers by Bone Histomorphometry 588 Conclusions 592 References 592

I. II. III.

IV.

CHAPTER 36 Genetic Markers of Joint Disease MICHEL NEIDHART, RENATE E. GAY, AND STEFFEN GAY I. Introduction 595 II. Ankylosing Spondylitis 595 III. Reactive Arthritis 597 IV. Rheumatoid Arthritis 598 V. Juvenile Rheumatoid Arthritis 605 VI. Conclusion 606 References 606

CHAPTER 37 Laboratory Assessment of Postmenopausal Osteoporosis PATRICK GARNERO AND PIERRE D. DELMAS I. Introduction 611 II. Postmenopausal Bone Loss 612 III. Management of Postmenopausal Osteoporosis 616 IV. Conclusion 624 References 624

CHRISTIAN MEIER, TUAN V. NGUYEN, AND MARKUS J. SEIBEL Introduction 649 Effects of Pretreatment Bone Turnover and Mineral Density on Therapeutic Outcomes The Role of Markers of Bone Turnover in Monitoring Antiresorptive Osteoporosis Therapy 651 Interpretation of Changes in Bone Turnover Markers 661 References 665

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CHAPTER 40 Age-related Osteoporosis and Skeletal Markers of Bone Turnover I. II.

III. IV.

CLIFFORD J. ROSEN Introduction 671 Epidemiology and Pathogenesis of Age-Related Osteoporosis-Relationship to Bone Turnover 672 Markers of Bone Turnover and Age-Related Osteoporosis – Clinical Implications 679 Summary 683 References 683

CHAPTER 41 Steroid-Induced Osteoporosis I. II. III. IV. V.

IAN R. REID Summary 689 Introduction 689 Epidemiology 689 Pathogenesis 690 Effects of Glucocorticoids on Histological Indices of Bone Turnover 691

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VI. Effect of Glucocorticoids on Markers of Bone Turnover 692 VII. Evaluation of Steroid-Treated Patients 694 VIII. Treatment and Follow-up 694 References 695

IV. Summary 762 References 762

CHAPTER 46 Primary Hyperparathyroidism

CHAPTER 42 Transplantation Osteoporosis: Biochemical Correlates of Pathogenesis and Treatment CAROLINA A. MOREIRA KULAK AND ELIZABETH SHANE I. Introduction 701 II. Kidney Transplantation 702 III. Cardiac Transplantation 705 IV. Liver Transplantation 706 V. Lung Transplantation 706 VI. Bone Marrow Transplantation 707 VII. Mechanisms of Bone Loss After Transplantation 707 VIII. Prevention and Management of Transplantation Osteoporosis 709 IX. Conclusions 711 References 711

CHAPTER 43

Secondary Osteoporosis

JEAN E. MULDER, CAROLINA A. MOREIRA KULAK, AND ELIZABETH SHANE I. Introduction 717 II. Hyperthyroidism and Osteoporosis 717 III. Osteoporosis Secondary to Hypogonadism 724 IV. Anticonvulsant Drugs and Osteoporosis 730 References 732

SHONNI J. SILVERBERG AND JOHN P. BILEZIKIAN Introduction 767 Etiology 767 Clinical Presentation 768 Bone Markers in Primary Hyperparathyroidism 770 V. Cytokines in Primary Hyperparathyroidism 772 VI. Treatment of Primary Hyperparathyroidism 772 VII. Summary 775 References 775 I. II. III. IV.

CHAPTER 47

ANDREAS GRAUER, ETHEL SIRIS, AND STUART RALSTON I. Introduction 779 II. Etiology and Pathogenesis 779 III. Treatment 784 References 788

CHAPTER 48 I. II. III. IV. V.

CHAPTER 44 I. II. III. IV. V. VI.

Osteomalacia and Rickets

MARC K. DREZNER Definition 739 Etiology 741 Incidence and Epidemiology 742 Calciopenic Rickets and Osteomalacia 743 Phosphopenic Rickets and Osteomalacia 747 Normal Mineral Rickets and Osteomalacia 750 References 751

CHAPTER 45 Assessment of Bone and Joint Diseases: Renal Osteodystrophy ESTHER A. GONZÁLEZ, ZIYAD AL ALY, AND KEVIN J. MARTIN I. Introduction 755 II. Biochemical Assessment of Renal Osteodystrophy 757 III. Skeletal Imaging in Renal Osteodystrophy

Metastatic Bone Disease

JEAN-JACQUES BODY Abstract 793 Introduction 794 Use of Markers of Bone Turnover for the Diagnosis of Bone Metastases 796 Use of Markers of Bone Turnover for the Monitoring of Tumor Bone Disease 802 Prediction of the Development of Bone Metastases 806 References 806

CHAPTER 49 I. II. III. IV. V.

I. II. III. IV. V. VI.

Rare Bone Diseases

MICHAEL P. WHYTE Introduction 811 Osteopenia 812 Osteosclerosis and Hyperostosis Ectopic Calcification 823 Other Disorders 825 References 826

CHAPTER 50

761

Paget’s Disease of Bone

817

Osteogenesis Imperfecta

FRANCIS H. GLORIEUX AND FRANK RAUCH Introduction 831 Classification 832 Diagnosis 833 Differential Diagnosis 834 Pathogenesis 834 Bisphosphonate Therapy in OI 835

xii VII. Medical Therapies Other than Bisphosphonates 838 VIII. Potential Future Therapies 839 IX. Conclusions 839 References 839

CHAPTER 51 Rheumatoid Arthritis and other Inflammatory Joint Pathologies STEVEN R. GOLDRING AND MARY B. GOLDRING I. Abstract 843 II. Introduction 843 III. Effects of Joint Inflammation on Skeletal Remodeling 844 IV. Effects of Joint Inflammation on Cartilage Remodeling 852 V. Conclusion 858 References 858

Contents

CHAPTER 52 Osteoarthritis and Degenerative Spine Pathologies I. II. III. IV. V. VI. VII. VIII. IX. X. XI.

KRISTINA ÅKESSON Introduction 871 Characteristics of OA 872 Etiology 872 Treatment Options 873 Current Diagnostic Procedures 873 Biochemical Aspects of Osteoarthritis 873 Markers of Bone Turnover 874 Markers of Cartilage Metabolism 878 Spine Degeneration and Markers 882 Summary 883 Conclusions 884 References 884

Contributors

Kristina Åkesson Associate Professor, Department of Orthopedics, Malmö University Hospital, 205 02 Malmö, Sweden

John P. Bilezikian Department of Medicine, College of Physicians and Surgeons, Columbia University, New York, NY, USA Departments of Medicine and Pharmacology, College of Physicians and Surgeons, Columbia University, New York, USA

Sari L. Alatalo Finnish Red Cross Blood Service, Helsinki, Finland Susan J. Allison Postgraduate Scholar, Bone Research Program, Garvan Institute of Medical Research, 384 Victoria Street, Sydney NSW 2010, Australia

Neil Binkley University of Wisconsin, Madison, Wisconsin, USA Jean-Jacques Body Dept of Internal Medicine and Endocrinology/ Bone Diseases Clinic, Institut J. Bordet, Univ. Libre de Bruxelles, Brussels, Belgium

Ziyad Al Aly Division of Nephrology, Saint Louis University School of Medicine, St. Louis, Missouri, USA

Jean-Philippe Bonjour Division of Bone Diseases, WHO Collaborating Center for Osteoporosis Prevention, Department of Rehabilitation and Geriatrics, University Hospitals, CH - 1211 Geneva 14 (Switzerland)

Paul A. Baldock Senior Research Officer, Bone Research Program, Garvan Institute of Medical Research,384 Victoria Street, Sydney NSW 2010, Australia John P. Bilezikian Departments of Medicine and Pharmacology, College of Physicians and Surgeons, Columbia University, New York, NY

Adele L. Boskey Starr Chair in Mineralized Tissue Research, Hospital for Special Surgery, New York, NY 10021 and Weill Medical College and Graduate School of Medical Sciences of Cornell University, New York, NY 10021

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xiv Roger Bouillon Laboratorium for Experimental Medicine and Endocrinology, K. U. Leuven, Gasthuisberg, Herestraat 49, 3000 Leuven, Belgium Kim Brixen Department of Endocrinology, Odense University Hospital, DK-5000 Odense C, Denmark Peter Bruckner Department of Physiological Chemistry and Pathobiochemistry, University of Münster, Münster, Germany Geert Carmeliet Laboratorium for Experimental Medicine and Endocrinology, K. U. Leuven, Gasthuisberg, Herestraat 49, 3000 Leuven, Belgium Ian M Clark School of Biological Sciences, University of East Anglia, Norwich NR4 7TJ, UK Peter Croucher Academic Unit of Bone Biology, University of Sheffield Medical School, Sheffield S10 2RX, United Kingdom Pierre D. Delmas Professor of Medicine, Université Claude Bernard, Lyon, France and Director INSERM Research Unit 403, Lyon, France David W. Dempster Regional Bone Center, Helen Hayes Hospital, West Haverstraw, New York, USA Jean-Pierre Devogelaer Department of Rheumatology, Saint Luc University Hospital, Université Catholique de Louvain, 1200 Brussels, Belgium Marc K. Drezner Professor of Medicine, University of Wisconsin, Madison, Wisconsin, USA Richard Eastell From the Academic Unit of Bone Metabolism, University of Sheffield, Sheffield, UK Guy Eelen Laboratorium for Experimental Medicine and Endocrinology, K. U. Leuven, Gasthuisberg, Herestraat 49, 3000 Leuven, Belgium

Contributors

Erik Fink Eriksen Novartis Pharma, Basel, Switzerland Lorraine A. Fitzpatrick Global Development, Amgen, Thousand Oaks, CA Ghada El-Hajj Fuleihan Calcium Metabolism and Osteoporosis Program, American University of Beirut-Medical Center, Beirut, Lebanon Edith M. Gardiner Associate Professor, School of Medicine, The University of Queensland, Head, Skeletal Biology Unit, Centre for Diabetes & Endocrine Research, Ground Floor, C Wing, Bldg 1, Princess Alexandra Hospital, Ipswich Road, Brisbane QLD 4102, Australia Patrick Garnero Research Scientist, INSERM research unit 403 and Vice-President Synarc Molecular Marker Division, Lyon, France Renate E. Gay Research Scientist, INSERM research unit 403 and Vice-President Synarc Molecular Marker Division, Lyon, France Steffen Gay Research Scientist, INSERM research unit 403 and Vice-President Synarc Molecular Marker Division, Lyon, France Francis H. Glorieux Genetics Unit, Shriners Hospital for Children, 1529 Cedar Avenue, Montréal, Québec, Canada H3G 1A6 Mary B. Goldring Medical Center, Harvard Medical School, Boston, MA; New England Baptist Bone and Joint Institute, Boston, MA 02215 Steven R. Goldring Department of Medicine, Rheumatology Division, Beth Israel Deaconess David Goltzman Calcium Research Laboratory, Department of Medicine, Royal Victoria Hospital, McGill University, Montreal, H3A 1A1, Canada

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Contributors

Esther A. González Division of Nephrology, Saint Louis University School of Medicine, St. Louis, Missouri, USA

Helena Kaija Research Center for Molecular Endocrinology, University of Oulu, Finland

Andreas Grauer Procter & Gamble Pharmaceuticals Mason, OH, USA

S. Khosla Division of Bone Diseases, WHO Collaborating Center for Osteoporosis Prevention, Department of Rehabilitation and Geriatrics, University Hospitals, CH - 1211 Geneva 14 (Switzerland)

Caren M. Gundberg Department of Orthopaedics and Rehabilitation, Yale University School of Medicine, New Haven, Connecticut 06510 Tim Hardingham Wellcome Trust Centre for Cell-Matrix Research, Faculty of Life Sciences, University of Manchester, Manchester M13 9PT, United Kingdom

Marius E. Kraenzlin Division of Endocrinology, Diabetology and Clinical Nutrition, University Hospital Basel, Petersgraben 4, CH-4031 Basel, Switzerland Barbara E. Kream Departments of Medicine and Genetics and Developmental Biology, University of Connecticut Health

John R. Harrison Division of Orthodontics, University of Connecticut Health Center, Farmington, CT 06030

Center, Farmington, Ct 06030

Dick Heinegård Departments of Experimental Medical Science and Clinical Science, BMC plan C12, SE-22184, Lund, Sweden

Carolina A. Moreira Kulak Department of Endocrinology, Federal University of Parana, Hospital de Clinicas, Curitiba, Brazil Division of Endocrinology and Metabology of Hospital de Clinicas, Federal university of Parana (SEMPR), Curitiba-/Brazil

M. H. Helfrich Department of Medicine and Therapeutics, University of Aberdeen, Institute of Medical Sciences, Foresterhill, Aberdeen AB25 2ZD, United Kingdom

Johannes P.T.M. van Leeuwen Department of Epidemiology & Biostatistics, Erasmus Medical Centre, Rotterdam, The Netherlands

Herbert Herzog Adjunct Professor, Faculty of Medicine, The University of New South Wales, Principal Research Fellow, Head Obesity and Energy Homeostasis Research Group, Director Neurobiology Program, Garvan Institute of Medical Research, 384 Victoria Street, Sydney NSW 2010, Australia

Gary L. Lensmeyer University of Wisconsin, Madison, Wisconsin, USA

M. A. Horton Bone and Mineral Centre, Department of Medicine, The Rayne Institute, London WC1E 6JJ, United Kingdom

Joseph A. Lorenzo Director, Bone Biology Research, Professor of Medicine, University of Connecticut Health Center, 263 Farmington Avenue, MC 1317, Farmington, CT 06030–1317.

Philippa Hulley Botnar Research Centre, Institute of Musculoskeletal Sciences, University of Oxford, Oxford OX3 7LD, United Kingdom

Pilar Lorenzo Departments of Experimental Medical Science and Clinical Science, BMC plan C12, SE-22184, Lund, Sweden

Harald Jüppner Endocrine and Pediatric Nephrology Units, Massachusetts General Hospital and Harvard Medical School, Boston, MA, USA

Christa Maes Laboratorium for Experimental Medicine and Endocrinology, K. U. Leuven, Gasthuisberg, Herestraat 49, 3000 Leuven, Belgium

Jane B. Lian University of Massachusetts Medical School, Department of Cell Biology, 55 Lake Avenue North, Worcester, MA 01655, USA

xvi Daniel-Henri Manicourt Laboratoire de Chimie Physiologique (Metabolic Research Group, Connective Tissue Section), Christian de Duve Institute of Cellular Pathology and Department of Rheumatology, Saint Luc University Hospital, Université Catholique de Louvain, 1200 Brussels, Belgium Klaus von der Mark Dept. of Experimental Medicine and Connective Tissue Research, Friedrich-Alexander, University of Erlangen Nuremberg, Germany Kevin J. Martin Division of Nephrology, Saint Louis University School of Medicine, St. Louis, Missouri, USA T. John Martin St. Vincent’s Institute of Medical Research, 9 Princes Street, Fitzroy 3065, Australia

Contributors

Tuan V. Nguyen Bone and Mineral Research Program, Garvan Institute of Medical Research, St. Vincent’s Hospital and University of NSW Research Program University of New South Wales, Sydney, Australia Satoru K. Nishimoto Department of Molecular Sciences, The University of Tennessee College of Medicine, Memphis, Tennesse 38163 Lila O.T. Patrikainen Research Center for Molecular Endocrinology, University of Oulu, Finland Huibert A.P. Pols Department of Endocrinology, Laboratory Group, Im Breitspiel 15, 69126 Heidelberg, Germany

Christian Meier Endokrinologische Praxis & Labor, University Hospital Basel, Switzerland

Lawrence G. Raisz Interim Director, Musculoskeletal Institute, Board of Trustees Distinguished Professor of Medicine, University of Connecticut Health Center, 263 Farmington Avenue, MC-3805, Farmington, CT 06030

Joyce B.J. van Meurs Department of Epidemiology & Biostatistics, Erasmus Medical Centre, Rotterdam, The Netherlands

Stuart Ralston Rheumatic Diseases Unit, University of Edinburgh, Edinburgh, UK

José Luis Millán The Burnham Institute, 10901 North Torrey Pines Road, La Jolla, CA 92037-1005, USA

Frank Rauch Genetics Unit, Shriners Hospital for Children, 1529 Cedar Avenue, Montréal, Québec, Canada H3G 1A6

David G. Monroe Biochemistry and Molecular Biology, Mayo Clinic College of Medicine, Rochester, MN

Ian R. Reid Department of Medicine, University of Auckland, Private Bag 92019, Auckland, New Zealand

Jean E. Mulder Department of Medicine, Brigham and Women’s Hospital, Harvard University, Boston, MA 02115

Juha Risteli Department of Clinical Chemistry, FI-90014 University of Oulu, Oulu, Finland

Gillian Murphy Dept of Oncology, Cambridge University, Cambridge CB2 2XY, UK

Leila Risteli Department of Clinical Chemistry, FI-90014 University of Oulu, Oulu, Finland

Kim E. Naylor From the Academic Unit of Bone Metabolism, University of Sheffield, Sheffield, UK

Fernando Rivadeneira Department of Epidemiology & Biostatistics, Erasmus Medical Centre, Rotterdam, The Netherlands

Michel Neidhart Research Scientist, INSERM research unit 403 and Vice-President Synarc Molecular Marker Division, Lyon, France

xvii

Contributors

René Rizzoli Division of Bone Diseases, WHO Collaborating Center for Osteoporosis Prevention, Department of Rehabilitation and Geriatrics, University Hospitals, CH - 1211 Geneva 14 (Switzerland) Simon P. Robins Matrix Biochemistry Group, Rowett Research Institute, Bucksburn, Aberdeen AB21 9SB Clifford J. Rosen Maine Center for Osteoporosis Research and Education, St. Joseph Hospital, 900 Broadway, Bldg #2, Bangor, Maine 04401 USA Mishaela R. Rubin Department of Medicine, College of Physicians and Surgeons, Columbia University, New York, NY, USA Graham Russell Botnar Research Centre, Institute of Musculoskeletal Sciences, University of Oxford, Oxford OX3 7LD, United Kingdom Tore Saxne Departments of Experimental Medical Science and Clinical Science, BMC plan C12, SE-22184, Lund, Sweden Heinrich Schmidt-Gayk Department of Endocrinology, Laboratory Group, Im Breitspiel 15, 69126 Heidelberg, Germany

Natalie A. Sims University of Melbourne, Department of Melbourne, St. Vincent’s Health, 41 Victoria Pde, Fitzroy 3065, Australia Ethel Siris Department of Medicine, Columbia University College of P&S, New York, NY, USA Thomas C. Spelsberg Professor of Biochemistry, Mayo Clinic College of Medicine, Rochester, MN Gary S. Stein University of Massachusetts Medical School, Department of Cell Biology, 55 Lake Avenue North, Worcester, MA 01655, USA Eugene J.-M. A. Thonar Departments of Biochemistry, Internal Medicine and Orthopedic Surgery, Rush Medical College, RushPresbyterian-St. Luke’s Medical Center, Chicago, Illinois, 60612 USA André G. Uitterlinden Department of Internal Medicine, H. Kalervo Väänänen Institute of Biomedicine, Department of Anatomy, University of Turku, Turku, Finland

Ego Seeman Dept of Endocrinology and Medicine, Austin Hospital, University of Melbourne, Melbourne, Australia.

Annemieke Verstuyf Laboratorium for Experimental Medicine and Endocrinology, K. U. Leuven, Gasthuisberg, Herestraat 49, 3000 Leuven, Belgium

Markus J. Seibel Bone Research Program, ANZAC Research Institute, University of Sydney and Concord Hospital, Sydney, Australia

Pirkko T. Vihko Department of Biological and Environmental Sciences, Division of Biochemistry, University of Helsinki, Finland

Elizabeth Shane Department of Medicine, College of Physicians and Surgeons, Columbia University, New York, New York 10032

Michael P. Whyte Center for Metabolic Bone Disease and Molecular Research, Shriners Hospitals for Children; and Division of Bone and Mineral Diseases, Washington University School of Medicine At Barnes-Jewish Hospital; St. Louis, Missouri

Shonni J. Silverberg Department of Medicine, College of Physicians and Surgeons, Columbia University, New York, USA Peter A. Simkin Professor of Medicine, Adjunct Professor of Orthopaedics, University of Washington

Hua Zhou Regional Bone Center, Helen Hayes Hospital, West Haverstraw, New York, USA

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Preface to the Second Edition

The first edition of “Dynamics of Bone & Cartilage Metabolism”, published at the very end of the last century, in 1999, was successfully received by biomedical scientists in the bone field and by clinicians throughout the world. Since the first edition was published, research in bone and cartilage metabolism has progressed at a rapid pace leading to new insights into basic science as well as new ways in which markers of bone and cartilage metabolism can be used clinically. This second edition of “Dynamics of Bone & Cartilage Metabolism” incorporates these advances while maintaining the general structure of the first edition. It is a thorough update with all chapters either extensively revised or completely rewritten. To reflect the changing climate of knowledge, twelve new chapters have been added that, we believe, greatly enhance the substance and completeness of the book. The topics of these new chapters include: “Acid Phosphatases”, “Bone Structure, Architecture and Strength”, “Signalling Mechanisms in Bone”, “The Central Control of Bone Remodelling”, “Transgenic Models of Bone Disease”, “Models of Cartilage Metabolism and Disease”, “Measurement of

Parathyroid Hormone”, “Measurement of Vitamin D”, “Variability of Bone Markers”, “Monitoring of Anabolic Treatment”, “Monitoring of Anti-resorptive Treatment” and “Osteogenesis Imperfecta”. These new chapters add greatly to updated chapters and together provide a complete repository of information on this subject. We are grateful to all authors for their efforts to deliver their new or revised chapters within the time constraints, always a challenge. We are also grateful to the excellent staff at Elsevier-Academic Press, Tari Broderick, Karen Dempsey and Renske van Dijk, who helped us so enthusiastically throughout the preparation of this new edition of Dynamics of Bone and Cartilage Metabolism. MARKUS SEIBEL, Sydney SIMON ROBINS, Aberdeen JOHN BILEZIKIAN, New York June 2006

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Part I

Components of the Organic Extracellular Matrix of Bone and Cartilage

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Chapter 1

Structure, Biosynthesis and Gene Regulation of Collagens in Cartilage and Bone Klaus von der Mark

Department of Experimental Medicine and Connective Tissue Research, Friedrich-Alexander, University of Erlangen Nuremberg, Germany

VI. Collagen Genes and Transcriptional Regulation VII. Factors Regulating Collagen Biosynthesis VIII. Conclusions References

I. Introduction II. The Collagen Families III. Bone Collagens IV. Cartilage Collagens V. Collagen Biosynthesis

I. INTRODUCTION

collagen which may provide additional elasticity [8–11]. In addition, cartilage contains minor amounts of other collagen types depending on the cartilage type and location (see below). The basic function of collagens in cartilage and bone is to provide the structural scaffold to tissues into which minerals, proteoglycans, and glycoproteins can be firmly incorporated, thus being responsible for the unique physiological and mechanical properties of these tissues. But on top of biomechanical functions, collagens play an important role in all tissues, including cartilage and bone as biological substrates for cell adherence. Essential cell biological functions such as proliferation, cytoskeletal organization, migration, differentiation, and apoptosis are regulated by collagens, mediated by transmembrane receptors of the integrin and syndecan families [12]. Since collagens provide the major organic component with 90% of the dry mass in bone, or 60% in cartilage, respectively, it is obvious that defects in structure,

Collagens provide the structural framework of bones and cartilages and hold responsibility for shape and most of the biomechanical properties such as resistance to pressure, torsion, and tension [1]. In vertebrates, 27 genetically distinct collagen types with rather diverse structural and biochemical features have been identified, but only about half of them are represented in cartilage and bone [2–4] (Table I). Their specific functions in the tissues are only partially known. In cartilage and bone, fibril-forming collagens are dominant: the bone matrix consists basically of two collagen types, about 95% type I and 5% type V collagen which are assembled into heterofibrils [5]. Similarly, the backbone of all cartilages is made of types II/XI collagen heterofibrils which are decorated with so-called FACIT collagen types IX, XII, or XIV (see below) [3, 4, 6, 7]. These fibrils are interwoven with a microfibrillar mesh made of type VI Dynamics of Bone and Cartilage Metabolism

3

Copyright © 2006 by Academic Press. All rights of reproduction in any form reserved.

4

KLAUS VON DER MARK Table I.

Collagens in Cartilage and Bone

Type

Subunits

I

α1(I) α2(I)

α1(I)2 α2

I II

α1(I) α1(II)

[α1(I)]3 [α1(II)]3

III

α1(III)

[α1(III)] 3

V

[α1(V)]3 [α1(V)]2 α3(V)

XXIV

α1(V) α2(V) α3(V) α1(XI) α2(XI) α3(XI) α1(XXIV)

[α1(XI) α2(XI) α3(XI)] [α1(XI)]2 α2(V) n.d.

Bone, eye

XXVII

α1(XXVII)

n.d.

Cartilage; eye and ear

Forms fibers of high tensile strength; most abundant collagen Rare form Major cartilage collagen; forms heterofibrils with Col IX + XI Often in mixed fibrils with type I collagen; present in reticular fibers and elastic tissues; cystine bridges in triple helix Propeptide partially retained in the fibrils; forms hetero-fibrils with type I collagen controls fibril diameter Homologous to Col V; nucleates and controls cartilage coll. fibril formation; α3(XI) same gene as for α1(II) Similar to type V collagen, contains TSP motif in N-propeptide Col27a1 gene 156 kb, 61 exons

VI

α1(VI) α2(VI) α3(VI)

[α1(VI) α2(VI) α3(VI)]

Microfibrillar collagen Widespread, in cartilage (pericellular), intervert, disk dermis, placenta, lung vessel wall

Contains vWF and Kunitz type protein inhibitor domains; forms beaded filaments; highly disulfide crosslinked

X

α1(X)

[α1(X)]3

IX

[α1(IX) α2(IX) α3(IX)]

XII

α1(IX) α2(IX) α3(IX) α1(XII)

[α1(XII)]3

XIV

α1(XIV)

[α1(XIV)]3

XVI

α1(XVI)

[α1(XVI)]3

XX XXI XXII

α1(XX) α1(XXI) α1(XXII)

XI

Molecular forms

Tissue distribution Fibril-forming collagens Bone, dermis, tendon, ligaments cornea, most other tissues Dermis, dentin Cartilage, notochord, vitreous body embryonic epithelia, retina Reticular fibers of most tissues (lung, liver, dermis, spleen, vessel wall, etc.) In vitro: hamster lung cell cultures lung, cornea, bone, fetal membranes; together with Col I Cartilage, vitreous body Bone

Network forming, short chain collagens Hypertrophic cartilage

FACIT collagens Cartilage, vitreous humor Splice variant without NC-4 domain in cornea Perichondrium, ligaments, tendon Cartilage, dermis, tendon, vessel wall, placenta, lung, liver Cartilage (territorial matrix) papillary dermis Cornea; sternal cartilage Blood vessel wall Myotendinous junction, articular cartilage surface

biosynthesis, assembly, or turnover of collagens generally will lead to severe diseases such as osteoporosis, osteoarthritis, chondrodysplasias, or osteogenesis imperfecta. In order to understand the specific functions of the various collagens in cartilage and bone and the reasons for their failure in connective tissue diseases, it is necessary to have a close look at the specific structural and biochemical features of the collagens. An overview on the structural and functional features of cartilage and bone collagens, and their interactions with other matrix proteins and cell receptors, will therefore be given in the first part of this chapter. In order to understand the dynamics of collagen metabolism in bone and cartilage, it is helpful to gain

Characteristic features

Strong inter- and intramolecular interactions between NC1-domains Mutations in Col X-NC-1 →SMCD

Covalently linked to type II collagen fibrils; NC4 domain projects into cartilage matrix; contains chondroitin sulfate Large cruciform shaped NC3 domain; associated with type I collagen fibrils Associated with type I collagen Integrates into discrete Col II/XI fibrils Less FN III repeats than Col XIV TSP and vWFA domain, but no FNIII rep. Present only in tissue junctions

insight into the various levels of transcriptional and translational regulation of collagen biosynthesis and turnover. Therefore, in the second part of this chapter, the various steps of collagen biosynthesis and post-translational modifications will be summarized, and an overview is given on the structure of the collagen genes and their cis-acting regulatory element. How these are regulated by growth factors, cytokines, hormones, and transcription factors is one of the current challenges in understanding dynamics of connective tissue turnover and homeostasis. The scope of this chapter does not permit a detailed and complete presentation of our current knowledge on structure, biosynthesis, and regulation of collagens in bone

5

CHAPTER 1 Structure, Biosynthesis and Gene Regulation of Collagens in Cartilage and Bone

and cartilage. Fortunately, a number of excellent review articles and book chapters are available, such as the recent, most comprehensive book chapter on collagens by Kielty and Grant [2] and many other articles dealing with structure and function of collagen types [6, 13–16], with collagen gene [17–19], collagen biosynthesis and regulation [20–23], collagen degradation [24, 25], and collagenrelated diseases [26–29]. No further information can be given here on collagens which do not play a major role in cartilage and bone such as types IV, VII, VIII, and XIII–XIX. Aspects of extracellular assembly to supramolecular structures, in particular fibril formation and cross-link formation, will be dealt with in Chapters 2, 11, and 20. Collagen degradation by proteases will be covered in Chapter 10 on matrix proteases.

II. THE COLLAGEN FAMILIES Up to 2004, at least 27 genetically distinct collagen types had been identified in mammals, encyphered by 42 genes coding for their subunits [2, 6, 15] (Table I). Based on molecular structure and sequence homology, collagens have been grouped into seven or eight different families, including the fibril-forming collagens, FACIT collagens, microfibrillar collagens, network-forming collagens, transmembrane collagens, multiplexins, and others (see Table I). Since the first edition of this book seven new collagen types have been discovered, partially by screening sequence data banks for homology with collagen and procollagen peptide sequences [30–37]. Four of these new collagens (types XX, XXI, XXII, and XXIV) were found also in cartilage, but their functions are not yet known. All collagens consist of one or several collagenous, triple-helical domains, flanked or interrupted by noncollagenous domains which are largely removed by proteolytic processing in fibrillar collagen type I, II, and III procollagen. They are retained, however, in the mature molecule in most non-fibril-forming collagens.

A. The Collagen Triple Helix The key feature of all collagens is the triple helix, a coiled-coil structure in the form of a right-handed helix of 1.5 nm diameter, composed of three polypeptide chains (α-chains) [38–40] (Fig. 1). A structural requirement for the assembly of polypeptide chains into a collagen triple helix is the occupation of every third position by a glycine residue, resulting in the (Gly-X-Y)n repeat structure characterizing all collagens. The α-chains form a stretched, left-handed helix with a pitch of 18 amino acid residues per turn [41] and assemble around a central axis in a manner allowing all glycine residues to be positioned in the center of the triple helix. The more bulky side chains of the other amino acids in the X and Y position occupy the outer positions, where they are available for lateral interactions with adjacent collagen molecules to form fibrils. Another typical feature of the collagen triple helix is the high content of proline and hydroxyproline (ca. 20%). The hydroxyl groups of 4-hydroxy-proline are essential for the formation of intramolecular hydrogen bonds and thus critically determine the thermal stability of each collagen triple helix [40, 42]. The melting temperature of the type I collagen triple helix is 39⬚C at neutral pH, but can be considerably higher, e.g. 46⬚C in chicken type X collagen [43], or 65⬚C in the aminoterminal 7S domain of type IV collagen [44]. Triple helical domains vary considerably in their length: in fibril-forming collagens they span 300 nm, corresponding to 1000 amino acid residues per processed α-chain, while in other collagens, such as the multiplexins, they may include only nine triplets. Interruptions of the Gly-X-Y-Gly-X-Y- structure by one amino acid residue causes flexibility in the triple helix and renders the helix susceptible to proteolytic attack, e.g. in collagen type IV [45]. The native or denatured state of a triple helix may be measured by optical rotation, circular dichroism or resistance to proteases like pepsin, trypsin, or chymotrypsin [46]. For example, native type I collagen molecules have an optical rotation of ε = –1000⬚ at 405 nm, which drops

p t

p

Figure 1 Type I procollagen as a prototype of fibril-forming collagens. In types I, II, and III procollagens N- and C-terminal propeptides are removed after secretion by specific proteases, in types XI and V the N-propeptides are larger (see Fig. 3) and only partially cleaved. The C-propeptides contain AsN-linked mannose-rich oligosaccharides, while the triple helical part contains only hydroxylysine-linked glucosyl-galactosyl disaccharides or monosaccharides.

6

KLAUS VON DER MARK

to –336⬚ after denaturation. The resistance of the native triple helix to most proteases has been the basis for almost all biochemical isolation procedures of collagens in the past, using pepsin to destroy non-collagenous proteins while leaving the collagen triple helices intact. Triple helical collagenous domains are also found in proteins such as C1q, acetylcholine esterase and MARCO, a macrophage scavenger receptor [47, 48].

NC-domains of IX, XI, XII, and XVI collagens. Many noncollagenous domains, e.g. in collagen VI and XI, contain thrombospondin- and von Willebrand-factor-like domains or fibronectin-type III repeats [59]. In the non-fibril-forming collagens, the noncollagenous domains often show more specific and important structural and functional features than the triple helical domains. Thus, the C-propeptide of type XVIII collagen is retained after cleavage as a protein with antiangiogenic properties (endostatin) [60].

B. Noncollagenous Domains In the various collagens the triple helical domains are flanked or interrupted by noncollagenous domains. While the triple helix is a highly conserved structural protein element like the α-helix or the β-pleated sheet, there is a wide structural and functional diversity among the noncollagenous domains of the different collagen families, often bearing essential functions specific for each collagen. Fibril-forming collagens are synthesized as procollagens with a triple helix of 300 nm length, which is flanked by noncollagenous propeptides (NC-domains) at both ends [2, 49]. The globular C-propeptides, consisting of about 250 amino acid residues per α-chain, are all homologous and serve as a nucleation site for chain assembly and triple helix formation, while the N-propeptides regulate the fibril size when retained in the molecule [50, 51]. These procollagen peptides are largely removed by specific proteases after secretion, a prerequisite for fibril formation [52]. The C-propeptide of type II collagen remains in cartilage after cleavage from the procollagen molecule as a stable, hydroxyapatite-binding molecule (“chondrocalcin”) and may participate in cartilage calcification [53]. The C-terminal NC-1 domain of type X collagen, a highly compact and stable, bell-shaped trimer, also binds calcium and is highly homologous to TNFα [54], but does not bind to the TNFα receptor; its role is rather structural, serving as a nucleation site for triple helix assembly as well as for network assembly [55, 56]. In “FACIT” collagens (fibril-associated collagens with interrupted triple helices) [57], which include collagen types IX, XII, XIV, XVI, XIX, XX, XXI, and XXII, the collagenous domains are interrupted by two or three noncollagenous domains, while the aminoterminal domains form large, cross-shaped entities anchored in the extracellular matrix. In the more recently discovered “multiplexins” (collagens with multiple triple helix interruptions, [58] see Table I) the collagenous domains are short and separated by nine or ten noncollagenous domains. Despite the differences between fibril-forming and FACIT collagens, there is also sequence homology among the N-terminal

C. Structural and Functional Diversity of Collagens There is considerable complexity and diversity in the structure of the different collagen types, their splice variants, their triple helical and nontriple helical domains, and their assembly into extracellular matrix structures: fibrils, flexible meshworks, hexagonal sheets, beaded filaments, anchoring fibrils and perhaps other, yet unknown, structures [15]. The fibril-forming collagens represent with seven members (including the recently discovered collagen XXIV and XVII) and about 90% of the total collagens, the most abundant and widespread family of collagens in vertebrates. Type IV collagens with a more flexible triple helix assemble into supercoiled chicken-wire like meshes that are restricted to basement membranes. Types VIII and X collagens are short-chain collagens which form hexagonal sheets. Type VI collagen is highly disulfide cross-linked and assembles into a meshwork of beaded filaments interwoven with collagen fibrils. Collagens IX, XII, and XIV associate as single molecules with collagen fibrils (FACITcollagens) [3, 7], while others (Col XIII and XVII ) span cell membranes [61–63]. Little is known on the extracellular architecture of the more recently discovered collagens, except for collagens XXIV and XXVII, which have all the features of fibril-forming collagens [37]. Owing to their tensile strength and torsional stability, the major function of fibril-forming collagens is to support tissue architecture and stability. Despite structural similarity, however, the fibrillar collagen types I, II, and III have different, specific functions in different tissues, different immunological properties and show specific interactions with different cell types. For example, replacement of type II collagen by the similar type I collagen in cartilage, e.g. in joint repair by fibrous cartilage, causes severe loss of cartilage-specific features of the tissue. Initial concepts of specific collagen functions were mostly derived from their distribution in tissues and from in vitro experiments, i.e. from cell and organ culture studies. Recently, there is ample evidence for specific cell biological functions of

CHAPTER 1 Structure, Biosynthesis and Gene Regulation of Collagens in Cartilage and Bone

7

fibrillar collagens, such as types I and II collagen, in serving as substrates for cell adhesion, proliferation, migration, and differentiation, mediated by β1 integrins [12, 64–67]. More recently, natural and artificially introduced mutations in human and animal collagen genes, and the inactivation of collagen genes by homologous recombination in mice (“knockout mice”) [68–70], have provided valuable information on the role of some collagens in cartilage and bone. Surprising results of gene knockout experiments in mice or from human mutations have often caused a revision of common opinions on the function of various collagens; thus, after inactivation of the type II collagen gene Col2a1 in mice, rather normal development of long bones and of the eye was observed, although early expression of Col2a1 in embryonic corneal and retinal epithelia [71–73] and several in vitro studies had strongly suggested a critical role of collagen in early epithelial–mesenchymal interactions [71, 74]. The lack of type II collagen in the notochord of the Col2a1−/− mice did not prevent somite differentiation as expected [75], but the resorption of the notochord during development of the spine [69]. More and valid information on the function of individual collagens and their domains have been obtained from targeted mutations in distinct domains and tissue-specific inactivation (conditional knockout) of collagen genes.

a quarter-staggered heterofibrils with diameters between 25 and 400 nm (Fig. 2). Fibrils thicker than 50 nm show a characteristic banding pattern in the electron microscope with a periodicity of 65–67 nm (D-period) [16, 77, 78]. Adachi and Hayashi [79] have shown that inclusion of type V collagen controls the fibril diameter of type I collagen fibrils; in embryonic chick cornea, for example, a content of 20% type V collagen limits the fibril diameter to 25 nm [80]. In contrast, collagen fibrils in bone with a content of ca. 5% type V collagen reach diameters of 400 nm or more [76, 81]. Embedded in hydroxyapatite crystals and various bone-specific phosphoproteins and glycoproteins and SLRPs (small leucine-rich proteoglycans) such as osteoadherin (see Chapter 3), type I/V heterofibrils reveal unmatched biomechanical properties concerning load bearing, tensile strength, and torsional resistance. They serve also as a nucleation site for hydroxyapatite crystals [82] (see Chapter 12). In the osteons of compact bone, collagen fibrils seem to run parallel in two nearly orthogonal directions, forming twisted, nearly rectangular plywood-like layers [83]. As the assembly of collagen molecules into fibrils will be dealt with in detail in Chapter 2, here the structure of the bone collagen molecules, their biosynthesis and regulation, are focused on.

III. BONE COLLAGENS

A. Type I Collagen

The organic mass of the bone matrix comprises about 90% of type I and 5% of type V collagen [76], the remainder being bone-specific phospho- and glycoproteins such as osteopontin, bone sialoprotein, osteocalcin, osteonectin, and others. In bone, type I and V collagen assemble into

1. MOLECULAR STRUCTURE AND TISSUE DISTRIBUTION

Figure 2

Type I collagen is the most abundant, longest-known and best-studied collagen in vertebrates. It forms 90% of the organic mass of bone and tendon and is the major collagen of skin, ligaments, cornea, and many interstitial

Packing of types II, XI, and IX collagen in the cartilage collagen heterofibril, showing the location of the covalent cross-links between type II and IX collagen. The globular NC4 domains of type IX collagen reach out of the fibril. (From: D. Eyre (2001) Collagen of articular cartilage. Arthritis Res. 4, 30–35, with kind permission by BioMed Central, Ltd.)

8 connective tissues. Much of our information on biochemical and biophysical properties, cross-linking, and biosynthesis of collagens is based on research on this collagen, but may be applied to other collagens. The human type I procollagen is made of 2 proα1 chains of 1464 amino acid residues [84] and a somewhat shorter pro α2 subunit (1366 amino acid residues) (Fig. 1) [85]. It is synthesized in large quantities by fibroblasts, osteoblasts [86], and odontoblasts [87], and to a lesser extent by nearly all other tissue cells [12]. Although purified type I collagen can be reconstituted to crossbanded fibrils in vitro, in vivo type I collagen is always incorporated into heterofibrils containing either type III collagen, e.g. in skin and reticular fibers [88], or type V collagen in bone, tendon cornea and other tissues [76, 89] or both. Type I/III collagen heterofibrils are a constituent of reticular fibers of most parenchymal tissues such as lung, kidney, liver, muscle, or spleen, with the exception of hyaline cartilage, brain, and vitreous humor [90]. The key role of type I collagen in bone is most evident from mutations in the human type I collagen genes COL1A1 and COL1A2 as the cause of osteogenesis imperfecta (OI), a group of hereditary disorders characterized by a decrease in bone mass, enhanced bone fragility and multiple fractures (see also Chapter 54; for reviews see [28, 91]. The severity and progress of the disease is rather variable and ranges from mild forms to more severe and lethal forms. The mode of inheritance of the OI is in most cases autosomal dominant. More than 160 different mutations have been reported in the COL1A1 and COL1A2 genes, located on chromosomes 17q21.3 and 7q21.3–q22, respectively [28]. Most mutations affect glycine residues, leading to impaired triple helix formations, even in heterozygotes, but also exon skipping, frameshift mutations, RNA splicing mutations and basepair deletions or insertions have been identified. Generally it is difficult to predict the severity of the phenotype from the type of mutation. The general rule is that apparently mild mutations affecting the stability of the triple helix, owing to a glycine substitution, result in a more severe phenotype than entire exon deletions, allowing the formation of intact, although shortened, triple helices. The reason is that one affected α chain with a triple helical interruption may exert a dominant negative effect and impair seven type I collagen molecules. Furthermore, impaired triple helix formation in the rough endoplasmic reticulum leads to over-hydroxylation of proline and lysine residues [92, 93]. Premature chain terminations or deletions of exons in one COL1A1 allele which do not affect triple helix assembly may cause haplo-insufficiency, but will still allow the formation of intact collagen I molecules from the unaffected allele [28, 91].

KLAUS VON DER MARK 2. PHYSIOLOGICAL FUNCTIONS

Besides its biomechanical properties, type I collagen is important as adhesive substrate for many cells and plays a major role in tissue and organ development, in cell migration, proliferation and differentiation, in wound healing, tissue remodeling, and hemostasis [12, 74]. For example, in vitro studies have shown that many epithelial and endothelial cells acquire a polar cell shape and develop a luminal structure when cultured in a three-dimensional hydrated collagen lattice [94]. Cells recognize native type I collagen via α1β1, α2β1, and α11β1 integrins [65, 95] which are transmembrane receptors and confer signals from the extracellular matrix to intracellular signal cascades and to the actin cytoskeleton [96–98]. Furthermore, α11β1 integrin is able to organize the assembly of extracellular type I collagen molecules into fibrils [99]. Thus, reconstituted hydrated lattices consisting of native type I collagen fibrils are widely used for cell and tissue culture purposes, allowing cells to migrate, proliferate, and differentiate in a native, three-dimensional environment. Similarly, freezedried collagen sponges find wide applications in surgery and tissue engineering as scaffolds for wound and tissue repair by supporting adhesion and invasion of connective tissue cells [100–102]. Valuable and important information on the role of type I collagen has been gained from the MOV13 mouse strain, in which the expression of α1(I) chains is blocked owing to an insertion of the MULV Moloney virus into the first intron [103, 104]. Homozygous embryos die at day 13.5 owing to vessel rupture, but early development of organs is normal in the absence of type I collagen. Organ culture of salivary glands or lung buds showed that branching morphogenesis is also normal in the absence of type I collagen [105], possibly owing to a supplementing effect by type III collagen. Interestingly, in organ culture of tooth and bone anlagen α1(I) collagen is expressed despite the insertion of the Moloney virus in the Col1a1 gene [106, 107]. This observation led to the discovery of a bonespecific control of Col1a1 transcription starting at a site which differs from the transcriptional control in fibroblasts (see Section VI).

B. Type V Collagen Type V and XI collagen are closely related in structural and evolutionary terms and have therefore been grouped into a clade of fibril-forming collagens with similar biochemical properties and similar functions in the organism. Type V collagen usually co-distributes with type I collagen in bone, corneal stroma, interstitial matrix of smooth muscle, skeletal muscle, liver, lung, and placenta [80, 108], while type XI

CHAPTER 1 Structure, Biosynthesis and Gene Regulation of Collagens in Cartilage and Bone

collagen is attributed to cartilage collagen. Both collagens precipitate in their pepsin-treated form between 0.8 and 1.2 M NaCl at acidic pH, a feature which was decisive for their first discovery and separation from the dominant type I or type II collagens, respectively [114, 109]. Five of the pro-α-chains of collagen V and XI are further characterized by large amino terminal noncollagenous domains, which are partially retained in the fibrils [50]. The 400 amino acid residue globular domains of α1, α2, α3 (V), and α1 and α2(XI), located between signal peptide and the short triple helix of the N-propeptide, are about twice as large as the corresponding cysteine-rich region in α1(I), α1(II), and α1(III) (Fig. 2). They contain a proline/arginine rich domain (PARP domain) which is similar in α1(V), α2(V), α1(XI), and α2(XI), as well as in the N-terminal domain of collagen XII [110]. The domains are processed only partially after secretion, leaving stubs of 70–100 kDa in the fibril [50, 111] where they are critical for controlling fibril assembly and growth (see below). Unusual is the high content of tyrosine sulfate in the N-propeptide of α1(V) and α2(V) collagen chains [112]. With 40% of the tyrosine residues being O-sulfated, a strong regulating interaction with the more basic triple-helical part is likely to stabilize the fibrillar complex. In contrast to types I, II, and III collagen, the triple helical parts of types V and XI collagen are resistant to digestion with vertebrate collagenase (MMP1), but not to stromelysin [113]. Depending on the tissue, there is some heterogeneity in the composition of type V and XI molecules. Most tissues contain type V collagen molecules consisting of 2α1(V) and one α2(V) chain, but [α1(V)]3 homotrimers have been isolated from tumor cells [114], and some tissues contain α1(V), α2(V), α3(V) heterotrimers (see Table I). Also type V/XI hybrid molecules containing α1(V) and α2(XI) collagen chains were described in articular cartilage [4, 115], in bone [116], and in vitreous humor [117]. Immunohistochemical identification of type V collagen in tissue sections with antibodies requires demasking of the epitopes with acid or enzymes [118, 119], indicating a dense packing of type V collagen within the type I/V collagen heterofibrils [89].

IV. CARTILAGE COLLAGENS The backbone of all cartilaginous tissues of the vertebrate body is a heterofibril containing type II collagen as the predominant collagen type, into which type XI collagen is incorporated. The cartilage collagen fibril is decorated with FACIT collagens, mostly type IX collagen, which is covalently linked to type II collagen [7, 120]. The large N-terminal noncollagenous domains of FACIT

9

collagens, which are globular in the case of type IX collagen and cross-shaped in the case of types XII or XIV collagen, reach out of the fibril into the adjacent matrix space and may serve to anchor the collagen fibril in the proteoglycan matrix (see Fig. 2). Young growth cartilage contains about 85–90% type II collagen with 5–10% type XI and 5–10% collagen IX, while adult articular cartilage may have as little as 1% collagen IX and 3% collagen XI [4]. Substantial differences exist in the matrix composition of different cartilaginous tissues. Elastic cartilage contains collagens II, IX, and XI like hyaline cartilage, but elastin in addition. Fibrous cartilage contains a substantial amount of type I collagen besides type II collagen; type I collagen is also found in prechondrogenic tissue, e.g. in the perichondrium, and in tendon insertions [121]. In mammalian articular cartilage, type I collagen is restricted to the articular surface, where also the newly discovered collagen XXII was located [32], while chicken articular cartilage contains up to 30% type I collagen in the upper zone [122]. In osteoarthritic joints type I collagen was found in osteochondrophytes and fibrous repair tissue. There is still a debate on the question whether articular chondrocytes turn on type collagen I synthesis in osteoarthritis. While some studies report on the presence of type I collagen in osteoarthritic cartilage [123, 124], other immunohistological and in situ hybridization studies confirm the expression of α2(I) mRNA, but not of type I collagen protein in OA cartilage [125]. In contrast, type III collagen is an integral part of mammalian articular cartilage, where it has been located predominantly in the pericellular environment of chondrocytes [126]. Electron microscopic studies have shown that it is also incorporated in the type II collagen fibril [127]. Recently two new fibril-forming collagens (types XXIV and XXVII) [31, 37], and two new FACIT-like collagens (types XX and XXII) [30, 32] were found to be expressed in bone or cartilage, but their function is not yet known. The cartilage collagen fibrils are interwoven with a microfibrillar mesh consisting of type VI collagen, which is prominent in the chondrons of articular cartilage [9, 11]. Type X collagen, a network-forming collagen, is expressed predominantly in fetal and juvenile hypertrophic growth cartilage. It was also located to the upper zone of articular cartilage in certain joints in human, dog, and mouse cartilage [128–130], and in chondrocyte clusters of osteoarthritic cartilage and osteophytes [131, 132]. It supports endochondral ossification [55, 133, 134] and hematopoietic cell differentiation [135]. As a specific marker for hypertrophic chondrocytes it is widely used to analyze chondrocyte differentiation in skeletal development, but also to describe endochondral ossification in fracture callus and osteophyte formation [132, 136].

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A. Type II Collagen 1. STRUCTURE AND LOCALIZATION

Type II collagen is found predominantly, but not exclusively, in hyaline cartilage [137, 138], where it accounts for approximately 90% of the total collagen. It is a homotrimeric molecule with the composition [α1(II)]3 with similar size and biochemical features as type I collagen [3, 120], but it contains substantially more hydroxylysinelinked galactosyl-glucosyl disaccharides than type I collagen (ten disaccharides per α1(II) vs two per α1(I) chain). In early embryonic cartilage, type II collagen heterofibrils generally appear as 25–50-nm thin, unbanded fibrils, while in calcified cartilage or after reconstitution in vitro type II collagen may form up to 400-nm thick crossbanded fibrils with a 68-nm banding pattern repeat similar to type I collagen. Type II collagen exists in two splice variants; in the IIA splice form which is dominant in mature cartilage, exon 2 coding for a 69 amino acid residue, cysteine-rich domain in the N-terminal propeptide is spliced out. It is retained in the IIA splice variant, a transient embryonic form which was found in prechondrogenic mesenchyme, in perichondrium and vertebrae [139, 140]. Type II collagen is not only the major collagen of hyaline elastic [141] and fibrous cartilage [142, 143], but also represents the major collagen of vitreous humor [137, 144] and the nucleus pulposus of intervertebral disks. Furthermore, type II collagen is synthesized transiently by many embryonic epithelia such as notochord [73], cornea epithelium [72, 73], retina pigment epithelium, cranio facial mesenchyme, and endocardial and mesocardial tissues [71, 145–147]. 2. THE ROLE OF TYPE II COLLAGEN IN CARTILAGE FORMATION AND STABILITY

Much has been learned on the role of type II collagen in cartilage development and function from mutations in the human COL2A1 gene, causing a rather diverse spectrum of skeletal dysplasias such as achondrogenesis, hypochondroplasia, Stickler syndrome, spondyloepiphyseal dysplasia congenita, and Kniest syndrome [28, 148–150]. As in OI, the more severe forms of chondrodysplasias result from dominant negative mutations affecting triple helix stability such as glycine substitutions, overmodification of the proα1(II) and partial or complete intracellular degradation of type II procollagen molecules containing only one mutated α1(II) chain. For example, in the cartilages of achondrogenesis fetuses with a Gly769Ser mutation [151] or a Gly913Cys mutation causing hypochondrogenesis [152], no type II collagen was found, but instead a matrix containing types I and III collagen. Both mutations are lethal, demonstrating that type I and III collagen cannot replace the function and properties of type II collagen.

Inactivation of the type II collagen gene in mice by homologous recombination had severe consequences on skeletal development, in particular on notochord turnover [69] and vertebral development [153]. Interestingly, it did not affect early embryonic development of the eye, somites or craniofacial tissues, as had been predicted from the expression of type collagen in numerous early embryonic epithelia [71, 146]. Type II collagen-deficient mice die as a result of breathing and weaning inability due to thorax malformation and cleft palate formation, respectively. Long bones are shortened due to impaired endochondral ossification, and vertebral bodies and ribs are abnormal. Similar to achondrogenesis or hypochondrogenesis patients, the cartilaginous tissues contain chondrocyte-like cells, but a matrix consisting of types I and III collagen instead of type II collagen. Although the cartilaginous matrix contains aggrecan, the density of the collagen fibrils was lower and their structure abnormal [153]. Compared to other fibrillar collagens, type II collagen has unique antigenic properties: antibodies raised against chicken type II collagen cross-react with type II collagens from all other species including human, mouse, rat, calf, dog, sheep, and shark [90, 154], indicating highly conserved antigenic epitopes in the type II collagen molecule. Like other matrix components of articular cartilage, which has an immune privilege as a nonvascularized tissue, type II collagen is a major target for autoimmune responses in rheumatoid arthritis [155–157]. In animal models, purified native type II collagen has been shown to induce arthritis in certain strains of mice and rats [158, 159]. The major B-cell epitopes of type II collagen have been identified in the mouse, rat, and human system. They are confomationdependent and located near the integrin-binding site [160, 161]. Interestingly, the major T-cell epitope in rat and human type II collagen which are MHC restricted includes a galactosyl-glucosyl carbohydrate residue [162].

B. Type XI Collagen Type XI collagen is a heterotrimer consisting of α1, α2, and α3 (XI) subunits found predominantly in hyaline cartilage and vitreous humor associated with type II collagen. The α3(XI) subunit is identical in its amino acid sequence with α1(II) as it is translated from the Col2a1 gene, but it differs from α1(II) by a higher degree of hydroxylation and glycosylation [109, 163]. In mature cartilage, half of the α1(XI) molecules are replaced by α1(V) chains [164]. As in type V procollagen, the N-terminal domains of α1(XI) and α2(XI) chains are processed only partially after secretion, leaving stubs of 70–100 kDa in the fibril [163, 165]. Complex alternative splicing occurs within

CHAPTER 1 Structure, Biosynthesis and Gene Regulation of Collagens in Cartilage and Bone

the aminoterminal noncollagenous domains of α1(XI) and α2(XI) [166–168]. A proline- and arginine-rich subdomain (PARP-domain) of the aminopropetides of α1(XI) and α2(XI) seems to be rather stable and persists in the cartilage matrix [110, 169]. As an integral part of the cartilage collagen fibril, type XI collagen serves as a nucleation site of type II/XI collagen fibril formation and regulates the lateral growth of the fibrils [170, 171]. Within the fibril, type XI collagens are covalently cross-linked to each other through their N-telopeptide to helix interaction sites [7] and to the end of type II collagen molecules. The N-propeptide may stick out of the gap domains in the heterofibril (see Figs 2, 3) [172]. The pivotal role of type XI collagen in control of cartilage collagen fibril assembly became apparent also from the analysis of the gene defect of the cho-mouse mutant, which is affected with an autosomal recessive chondrodysplasia: a point mutation in the α1(XI) gene leading to chain determination caused absence of type XI collagen, resulting in a cartilage with irregular thick collagen fibrils, disorganized cartilage growth plate and disturbed chondrocyte differentiation [173]. Similarly, an in-frame deletion in the α2(XI) gene caused an autosomal recessive bone dysplasia or autosomal dominant Stickler syndrome [174].

C. Fibril-Associated Collagens with Interrupted Triple Helices (FACIT Collagens) 1. TYPE IX COLLAGEN

First evidence for the presence of additional collagenous proteins in cartilage was obtained in the form of various pepsin-resistant small collagenous fragments [175, 176]. Combined protein chemical and molecular biological efforts led to the elucidation of the complex structure of type IX collagen [177, 178]. It is a heterotrimeric molecule consisting of α1(IX), α2(IX) and α3(IX) chain, with three triple helical segments (COL1–COL3) that are interrupted and flanked by four globular domains NC1-NC4 [179, 180] (Figs 3, 4). Electron microscopical analysis of carefully dissected cartilage collagen fibrils revealed that the highly cationic NC4 domain, the largest domain with 243 amino acid residues and a pI of 9.7, reaches out from the fibril where it presumably interacts with proteoglycans [181]. A “hinge region” in NC3, caused by supernumerary amino acids in the NC3 domain of the α2(IX) chain, allows flexibility in the molecule. It is covalently linked to the surface of cartilage collagen fibrils with the collagenous domains COL1 and COL2 [182, 183]. The α2(IX) chain contains a chondroitin sulfate side chain [184, 185], linked to

-

Figure 3

11

Structural domains of fibril-forming collagens and the microfibrillar collagen type VI.

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Figure 4

Collagenous and noncollagenous domains of the FACIT collagens type IX, XII, and XIV. A = von Willebrand factor A domain.

a serine residue in the NC3 hinge region [186]. This CS chain is considerably longer in vitreous humor than in cartilage [187]. Owing to a second transcription start site between exons 6 and 7 in the α1(IX) collagen gene (see below), a shorter form of the α1(IX) collagen lacking the entire NC4 globule is expressed in the cornea and vitreous humor [188, 189]. The importance of type IX collagen for the integrity of cartilage matrix was underlined by the result of inactivation of the α2(IX) gene in mice. Animals homozygous for the deficiency in α2(XI) showed severe defects in cartilage development and revealed degenerative changes in adult articular cartilage similar to osteoarthritis [190]. 2. TYPES XII, XIV, AND XVI COLLAGEN

Type XII collagen is located predominantly in the perichondrium and articular surface, while type XIV collagen is more uniformly distributed throughout articular and tracheal cartilage [191]. Type XII collagen is a homotrimeric molecule with sequence homologies to type IX collagen in the C-terminal NC1 and COL1 domain, but only two collagenous domains which associate with type I or II collagen fibrils [192–194] (Fig. 4). The large, cross-shaped NC3 domain at the amino end reaches out into the perifibrillar

space [195]. This domain contains vWFA-domains, TSP and FN type III domains. Type XII collagen exists in two splice variants [196, 197]: the smaller IIB form with an α1(XII) chain of MW 220 kDa is found in skin, periosteum, and perichondrium. The larger XII form with an α-chain of MW 310 kDa was found in an epidermal cell line and contains a chondroitin sulfate chain. Collagen type XIV has a similar structure, although the cross-shaped NC3 domain is smaller than in Col XII [193, 198]. Type XIV collagen colocalizes with type I collagen in skin, tendon, lung, liver, placenta, and vessel walls by immunofluorescence [199, 200], and to some extent with type XII collagen in skin [201]. However, it does not bind directly to type I collagen, but to the dermatan sulfate side chain of decorin which associates with type I collagen [202, 203]. Collagen XVI is predominatly synthesized by fibroblasts and myoblasts, but also found in the territorial matrix of chondrocytes [204–206]. By immunohistochemistry and immunogold electron microscopy it was shown that collagen XVI is integrated in a discrete population of thin, D-banded collagen fibrils containing type II and XI that are distinct from type IX collagen-containing fibrils [204]. In the papillary dermis, however, collagen XVI is a component of fibrillin-1-containing microfibrils [204].

CHAPTER 1 Structure, Biosynthesis and Gene Regulation of Collagens in Cartilage and Bone 3. TYPES XIX, XX, XXI, AND XXII COLLAGENS

These recently discovered collagens are phylogenetically and structurally related to FACIT collagens, but whether they are associated with fibrils in vivo remains to be shown. Type XIX collagen is restricted to muscle in the embryo, but not in the adult [207, 208]. Collagen XX is most abundant in the corneal epithelium, but by RTPCR it has also been detected in sternal cartilage and tendon [30]. The collagen XXI gene codes for a short FACIT collagen containing a vWFA and a Tsp (thrombospondin)-domain like the other FACIT collagens, but little is known on the protein [209]. Collagen XXII exhibits a restricted localization at tissue junctions such as myotendinous junctions, the hair follicle basement membrane or the articular cartilage–synovial fluid interface [32].

D. Microfibrillar Collagens: Type VI Collagen Type VI collagen is the major collagenous component of microfibrils in elastic fibers and in a larger variety of tissues including cartilage, skin, blood vessels (intima), cornea, placenta, uterus, ciliary body, iris, and others [2, 210]. The type VI collagen molecule is a highly glycosylated, cysteine-rich heterotrimer consisting of two α-chains of ca. 1000 amino acid residues (α1(VI) and α2(VI)), and the long α3(VI) chain with about 3000 amino acid residues; the short triple helical core accounts only for about 20% of the molecule. It was discovered first in the form of pepsinresistant “short-chain” collagen with three α-subunits in smooth muscle and placenta [211]. The three α-chains share three noncollagenous domains which are homologous to the von Willebrand factor-A domain [212, 213]; the α3(VI) which exists in multiple splice variants [214–216] contains an additional eight VWF-A-domains at the N-terminus [216]. The C-terminal domain of α3(VI) also contains a Kunitz-type inhibitor motif and is essential for collagen VI assembly and secretion [217]. Interestingly, the α3(VI) subunit is down-regulated by γ-interferon, but the other subunits are not [218]. Type VI collagen interacts with other matrix proteins and proteoglycans, e.g. hyaluronan, heparan sulfate, decorin or NG2-proteoglycan [219], but also with type IV collagen in basement membranes [220]. Examination of type VI collagen in tissues or cell cultures by electron microscopy often reveals beaded filaments with 25-nm beads aligned in 100-nm intervals [221, 222]. Such structures can be assembled in vitro from type VI collagen tetramers which connect and overlap at the globular ends when visualized by rotary shadowing [210, 223]. Complexes of matrilin-1 and biglycan or decorin decorate type VI collagen and link it to type II collagen [224].

13

In the presence of lumican, however, type VI collagen can also assemble into hexagonal networks similar to type X collagen [225]. In tissues like skin or cartilage the type VI collagen forms a highly disulfide cross-linked, branched network, interwoven with fibrillar collagens [9]. Type VI collagen expression is up-regulated already in early stages of chondrocyte differentiation [226]. In mature cartilage it is preferentially located in the pericellular space [8, 9, 11]. It is enhanced in osteoarthritic cartilage [227], but has not been identified yet in calcified bone tissues. In some tissues such as nucleus pulposus and in some tumors type VI collagen filaments may assemble in a parallel fashion to give rise to sheets with the characteristic 100-nm periodicity [222, 228]. The pericellular location of type VI collagen is consistent with its highly adhesive properties for many cell types. In contrast to other fibrillar collagens, several RGD-sequences in the α2-and α3(VI)-chain were found to be recognized by integrin receptors [229].

E. Network Forming Collagens: Type X Collagen Hypertrophic cartilage in the growth plate of fetal and juvenile long bones, ribs, and vertebrae contains a shortchain collagen, type X collagen [55, 230–232] which is unique to this tissue in the normal organism and only found elsewhere under pathological conditions, e.g. in osteoarthritic articular cartilage and in chondrosarcomas [131, 132, 233–235]. There is, however, recent evidence that type X collagen is also present in small amounts in the menisci and in the surface layer of articular cartilage of certain human, mouse, and dog joints [128–130]. Type X collagen is a homotrimeric collagen with a 130-nm triple helical core (460 amino acids per chain), a large C-terminal globular NC1 domain and a short amino terminal NC2 globule [55, 232]. It is homologous in both sequence and tertiary structure to type VIII collagen, which is produced by endothelial cells [236]. Type VIII collagen assembles into sheets with a hexagonal arrangement in the Descemet’s membrane [237], and in vitro reconstitution experiments with chicken type X collagen indicate that this collagen is able to form similar structures [238]. The three C-terminal NC1 domains of type X collagen molecule associate with unusually high affinity to a dense bell-shaped trimer [54, 56, 239] which is homologous to the complement factor C1q and adipoQ and TNFα, not only by amino sequence [239] but also by their crystal structure [54]. Mutations in the NC1 domains lead to cartilage growth abnormalities and waddling gait in patients affected with Schmid-type metaphyseal chondrodysplasia (SMCD) [240–245]. In vitro studies on the chain assembly

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of type X collagen demonstrated that most Schmid mutations located at the inner face of NC1 monomer severely affect triple helix assembly [240, 246]. The function of type X collagen is not entirely clear. The severe cartilage and joint defects in SMCD patients clearly indicate a critical role for type X collagen in endochondral ossification; similarly, transgenic expression of a truncated chicken type X collagen gene in mice causes growth plate abnormalities and a hunch back [134, 247]. There is also indirect evidence that type X collagen might be involved in matrix-vesicle-initiated calcification of hypertrophic cartilage [133]. Surprisingly, however, the phenotypic changes in the growth plate of two strains of type X collagen knockout mice were only moderate [133, 248]. A subset of the Col10a1 knockout mice suffered from severe impairment of hematopoiesis, causing prenatal or early postnatal death [135, 249].

F. Multiplexins and Other Collagens Collagen types XV, XVI, XVII, XVIII and XIX are composed of 8–12 collagenous domains interrupted by short noncollagenous domains, allowing a flexibility in the molecular shape which is not possible in fibril-forming collagens [14, 250–254]. The cellular location and tissue distribution of most of these collagens has been described; yet, little is known of the possible function of these collagens. Collagen type XIII is a type II transmembrane collagen with a short, noncollagenous carboxyterminal domain in the cytoplasm and a large extracellular part which contains three collagenous domains [61]. Collagen type XIII may be involved in cell–matrix interactions by binding to fibronectin [255] and α1β1 integrin [256], but it has also antiadhesive properties. Bone is a major physiological site for collagen XIII expression [257], but its role in this tissue is still unclear. Collagen type XVIII, a heparan-sulfate-containing collagen which anchors cells to the basement membrane has also been localized in hyaline and fibrous cartilage [258–260]. The C-terminal NC1 domain called endostatin of Col XVIII has received much attention recently as it persists as a separate entity in tissues and blood and was shown to inhibit tumor angiogenesis by interfering with endothelial cell sprouting [60, 261].

V. COLLAGEN BIOSYNTHESIS Most of our knowledge on the mechanisms of collagen biosynthesis is based on studies on fibril-forming collagens, in particular type I collagen. For this reason, in this

chapter I will focus on biosynthesis events and mechanisms elucidated for the fibrillar collagens, but it is likely that most of it will apply also for other collagens.

A. Steps of Collagen Biosynthesis 1. TRANSCRIPTION AND ALTERNATIVE SPLICING

Most collagen genes are transcribed into different mRNA species owing to alternative splicing, to a different extent of polyadenylation at the 3′ end as in the case of type I, II, and III collagen, or to several transcription start sites in some collagen genes, as in the case of type IX collagen α2(I) (for review see [21, 262]). In the cornea and in the vitreous, a shorter form of the α1(IX) collagen gene (Col9a1) is generated owing to an alternative transcription start site between exons 6 and 7 [189]. Alternative splicing of collagen mRNAs was first described for type II collagen which exists in an embryonic, longer form (IIB) and a shorter, adult form (IIA) from which exon 2 is spliced out [140, 263]. Alternative spliced forms have been reported for many collagens such as collagen VI (see above), XI [168] and many others, for example type XIII with more than 12 splice variants [264]. Further heterogeneity among collagen mRNA species is introduced by heterogeneity in polyadenylation at the 3′ end. Thus, two α1(I) mRNA species of 4.8 and 5.8 kb length can be observed in fibroblast cultures [265, 266], and also for α2(I) and α1(II), and α1(III) two different mRNA species are detected in cell cultures depending on the cell type and culture conditions [267, 268]. 2. Translation

Collagens are synthesized as procollagen molecules in the lumen of the rough endoplasmic reticulum (RER), transported to the Golgi and secreted (with the exception of membrane-spanning collagens) into the extracellular space via secretory vesicles. The nascent pre-proα-chains protrude after translation from the ribosome-bound collagen mRNA into the lumen of the RER with the help of signal-recognition particles and receptors. Immediately after intrusion, the signal peptide is cleaved off, and the nascent procollagen chains are modified by hydroxylation and glycosylation before they align into triple helical procollagen molecules, which are secreted and processed before assembling into complex extracellular aggregates (for reviews see [21, 49]). 3. Hydroxylation and Glycosylation

In the triple helical region of all procollagens about half of the proline residues in the Y-position of the Gly-XY-triplets are converted to 4-hydroxyproline by the enzyme

CHAPTER 1 Structure, Biosynthesis and Gene Regulation of Collagens in Cartilage and Bone

prolyl-4-hydroxylase, an event which occurs only on nascent collagen α-chains before the triple helix is formed [269–271] (Fig. 5). A few proline residues (all in X position) are converted to 3-hydroxyproline by the enzyme prolyl-3-hydroxylase [272, 273]. Two isoforms of 4-prolylhydroxylase have been described: the type I enzyme with the composition [α(I)]2β2 has been cloned from human, rat, and chicken sources [274] and accounts for 60–90% of the total prolylhydroxylase activity in most cell types, while the type II form [α(II)]2β2 is more prevalent in chondrocytes [275]. The four subunits of the prolylhydroxylase encircle and slide along nascent pro α-chains, but are unable to hydroxylate prolyl-residues after the triple helix has formed. Therefore, the time when the α-chains are kept in a single-chain conformation is critical for the extent of hydroxylation of prolyl residues; the same is true for hydroxylation of lysine residues. Prolyl-3- and -4-hydroxylase, as well as lysylhydroxylase, require Fe2+, oxygen and 2-oxoglutarate as cofactors [269]. Ascorbic acid is required nonstochiometrically for uncoupled decarboxylation of 2-oxoglutarate. Thus, lack of oxygen or ascorbic acid, as well as chelation of Fe2+ by chelators such as α,α′-dipyridyl or o-phenanthroline, effectively prevent hydroxylation of

15

proline and lysine during collagen synthesis [276]. Since non- or under-hydroxylated collagens are degraded to a large extent intracellularly [277], the inhibition of prolylhydroxylase activity is considered a potent tool to control excess of collagen synthesis in fibrotic diseases [278]. A number of competitive inhibitors of prolylhydroxylase have been developed and are being tested: pyridine 2,4dicarboxylate, 3,4-dihydroxylbenzoate or L-mimosine [278, 279]. The hydroxylation of lysine to δ-hydroxylysine by the enzyme lysylhydroxylase [2, 280–282] is not only important for the glycosylation of collagen, but also for the formation of the covalent pyrrolic and pyridinium crosslinks formed after oxidation of lysine and hydroxylysine to aldehydes [283] (see Chapter 2). The extent of lysyl hydroxylation varies with the collagen type, species, the cell and tissue type, and may change during development and aging or under pathological conditions. For example, two lysine residues in the α1(I) chain, both in the telopeptide region, but about ten lysine residues in the α1(II), are hydroxylated to 5-hydroxylysine. Subsequently, glucosyl and galactosyl residues are transferred to the hydroxyl groups of hydroxylysine by glycosyl transferases [270, 284]

l

Figure 5 Biosynthesis of procollagen in the lumen of the rough endoplasmic reticulum (RER). After hydroxylation of the nascent α-chains, triple helix assembly and glycosylation the procollagen molecules are transported to the Golgi apparatus, secreted in secretory vesicles to the cell surface and processed. PDI, protein disulfide isomerase; HSP47 and BiP, molecular chaperones; PPI, prolyl-peptidyl-cis-trans-isomerase.

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KLAUS VON DER MARK

to form glucosyl-galactosyl disaccharides, while mannoserich oligosaccharides are coupled to a N-glycosylation receptor sequence in the C-propeptides of fibril-forming collagens [285]. 4. ALIGNMENT OF THE TRIPLE HELIX

Assembly of the procollagen chains to triple helical molecules occurs after hydroxylation of the nascent chains and completion of the C-propeptides. The correct conformation of the C-propeptides and of some N-propeptides is stabilized by the formation of intrachain disulfide bonds [286]. This is catalyzed by the enzyme protein disulfide isomerase (PDI) [287, 288], a reducing enzyme with a catalytic site similar to thioredoxin and present in many plant and animal cells. Interestingly, PDI is identical to the β-subunit of prolyl-4-hydroxylase [269, 271], and thus has a dual role: as a β2 dimer it catalyzes and controls the formation of intrachain and interchain disulfide bonds in the C- and N-propeptides; in combination with two α-subunits of the prolyl hydroxylase it catalyzes the hydroxylation of proline. Procollagen chain assembly involves a two-stage recognition event in the fibril-forming collagens [22]. (1) The first association and chain recognition in type I procollagen is driven by the C-propeptides [286]; essential for assembly are hydrophobic sequence patches with four hydrophobic amino acids that are conserved in their positions in all fibril-forming collagens. In type III procollagen, assembly of the procollagen chains and triple helix formation does not require the formation of interchain disulfide bonds which form later after the assembly of the trimer [289]. (2) Rate-limiting for the folding of the triple helix is a cis–trans-isomerization of prolyl peptide bonds in the α-chains, which is catalyzed by the enzyme peptidyl-prolyl cis–trans-isomerase (PPI) [288, 290]. Critical support for a key role of PPI in collagen chain-folding was the observation that cyclosporin, an inhibitor of PPI and other cyclins, blocks folding of procollagen I and III in cell cultures [290]. After alignment of the C-propeptides, in type I procollagen triple helix formation starts from the C-terminus and proceeds towards the N-terminus; it is driven by a patch of 3–4 Gly-Pro-Hypro triplets located at the C-terminal of the triple helix by providing a core stabilized by hydrogen bonds [22]. Although the role of C-propeptide in the chain assembly of fibril-forming collagens is firmly established, their role in chain selectivity in cells that produce several collagen types is not clear yet. Interestingly, only procollagen chains containing eight cysteines in the C-propeptide are able to form homotrimers (e.g. pro α1(I), α1(II), α1(III)), while procollagen chains lacking two or three of the eight cysteine residues such as pro α2(I) can only form heterotrimers [22].

5. Chaperones Involved in Folding and Processing of Procollagen

Several chaperones such as BiP and HSP47 are involved in the post-translational modification, folding, and processing of the procollagen molecule [291]. HSP47 is a heat shock protein and is expressed at ten-fold enhanced levels in chicken fibroblasts at 42–45⬚C [292]. It binds cotranslationally to the nascent pro-α-chain in the RER and prevents unspecific chain folding and aggregation [292, 293]. Interestingly, it also binds to native, triple helical collagen molecules in the RER and controls bundle formation, transport to the Golgi apparatus, and procollagen processing [291]. Although it has a basic pI of 9.0, the glycoprotein binds to basic collagen chains and to native collagen with a kDa of 10−6 to 10−7 M, but with high on and off rates [294]. In inflammatory situations, such as after treatment of cells with TNFα, HSP47 appears on the cell surface where it may invoke autoimmune reactions, causing rheumatoid arthritis [295]. 6. Procollagen Processing

Following secretion of procollagens in secretory vesicles, the C-propeptides of the fibril-forming collagen, and the N-propeptides of type I and II, partially also of type III collagen are cleaved by specific proteases. Both procollagen-Nproteinase and procollagen-C proteinases belong to a family of Zn2+-dependent metalloproteinases M12 [23, 296]. The N-proteinase exists in two splice variants in a longer form of 140 kDa (pNP1) and a shorter form of 70 kDa (pNP-2) [297, 298]. Both cleave the N-propeptides of types I and II procollagen [299], but only in the native state, leaving the N-telopeptide in the hairpin-loop configuration. The N-proteinase is a structural homolog to atrolysin and adamalysin II and contains an RGD-sequence and properdin-like repeats in the C-terminal part [23]. The procollagen-C-proteinase cleaves the C-propeptides of types I, II, and III collagen, as well as laminin 5 and lysyl oxidase precursor [23]. Curiously, the enzyme is partially identical to Bone Morphogenetic Protein 1 (BMP-1) or the mouse tolloid protease, and related to Drosophila tolloid, Astacin, and BP10 [300, 301]. All these enzymes share the Zn2+-binding proteinase domain, but the procollagen C-proteinases cCP-2 (mTld) (MW 110 kDa) or pCP-1 (BMP1) (70 kDa) differ in their C-terminal parts. The C-terminal propeptides that are cleaved off are retained for some while in the tissue. This has been shown in particular for type II collagen: the C-propeptide has been isolated from cartilage as an intact, disulfide-bonded entity which has been given the name chondrocalcin [53] as it is found in particular in calcified cartilage. Besides the described C-proteinases of the BMP-1 type, other proteinases seem to be able to cleave procollagen

CHAPTER 1 Structure, Biosynthesis and Gene Regulation of Collagens in Cartilage and Bone

propeptides, e.g. cathepsin D [302]. This possibility must be taken into consideration in view of the recent finding that inactivation of both alleles of BMP-1 in mice did not dramatically alter procollagen processing, but caused failure of body wall closure [301]. 7. Cross-linking

A key enzyme required for cross-linking of collagen α-chains and molecules is the enzyme lysyloxidase [282, 303], which catalyzes the oxidation of the ε-amino group of certain lysine and hydroxlysine residues to aldehyde groups. Lysine- or hydroxylysine-derived aldehyde groups initially form covalent Schiff-base-type bonds with ε-amino groups of other lysine and hydroxylysine residues in adjacent α-chains within the same or neighboring molecules. More detailed information about collagen cross-link formation will be provided in Chapter 2.

B. Translational and Post-translational Regulation of Collagen Biosynthesis Regulation of collagen biosynthesis occurs at several levels, involving a complex pattern of transcriptional, translational, and post-translational control mechanisms [304]. The following section will review studies showing the role of translational and post-translational mechanisms controlling the biosynthesis of fibrillar collagens. Little is known on translational or post-translational regulation of the other collagens. 1. Evidence for Translational Regulation

Evidence for translational control of collagen mRNA was provided by the observation that chondrocytes contain untranslated α1(I) mRNA [305]. Also splice variant of α2(I) mRNA was found in chondrocytes; curiously, it is translated into a noncollagenous protein [306, 307] using a different reading frame, but the function of this protein is as yet unclear. Another mechanism regulating collagen mRNA translation may be associated with the 3′-untranslated sequences. Polymorphic mRNA species exist for α1(I), α2(I), α1(II), α1(III), and α1(IV); for example Northern blotting of α1(I) reveals two bands of 4.7 and 5.7 kb in fibroblast monolayer [19, 265], which are expressed in different ratios depending on the cell type. This ratio increases significantly under the influence of TGFβ [308] which also increases the half life of the mRNA that is normally 9–10 h. In fibroblasts cultured in three-dimensional collagen gels, however, only the smaller form of α1(I) mRNA is expressed [267, 268]. As in other mRNA species, the 3′ untranslated segment of collagen mRNA is responsible for the stability and the translation rate of the collagen mRNA [309].

17

2. Regulation by MRNA Folding

There is experimental evidence that translation initiation of α1(I), α2(I), α1(II), and α1(III) mRNA may be controlled by highly conserved nucleotide sequences close to the translation initiation codon. RNA patches of about 50 nucleotides contain inverted repeat sequences which form stem loops or dimers and thus prevent translation initiation [310]. Deletion of these sequences in the α2(I) mRNA in vitro did not, however, always affect mRNA translation [311]. 3. Feedback Inhibition by N- and C-propeptides

Studies on fibroblasts with Ehlers–Danlos syndrome VII as well as cell culture studies indicated that the N-propeptides of type I and III collagen induce feedback inhibition of type I collagen synthesis [312]. The N-propeptide of α1(I) specifically inhibited cell-free translation of α1(I) mRNA [313] but, curiously, with fragmentation of the N-propeptide into smaller fragments, the inhibitory effect became unspecific [314]. Strong support for a specific role of the N-propeptide was the observation that it binds specifically to fibroblast membranes and is taken up by endocytosis [315]. Furthermore, expression of a metallothionine-inducible N-propeptide minigene in fibroblasts inhibited type I collagen synthesis [316]. Also, the type I procollagen C-propeptide has been implicated in the regulation of procollagen synthesis after internalization [317, 318]. It is secreted in significant amounts by osteoblasts and inhibits collagen synthesis to 80% of the control level [319]. In contrast to the N-propeptide, however, it is transported to the nucleus and seems to control collagen gene expression at the transcriptional level, while smaller fragments of the C-propeptide may regulate collagen synthesis both negatively or positively at the posttranscriptional level [318]. Similarly, the C-propeptide of type II collagen appears to inhibit translation of α1(II) mRNA [320].

VI. COLLAGEN GENES AND TRANSCRIPTIONAL REGULATION One of the current challenges in collagen research which needs to be solved for better understanding of the pathogenesis of collagen-related bone and cartilage diseases is the elucidation of mechanisms regulating tissue-specific expression of distinct collagen types. Tissue-specific expression of the various collagen types is regulated mostly, but not exclusively, at the transcriptional level and involves multiple control mechanisms at different levels. At the cell surface hormones, growth factors, cytokines, and their antagonists compete for their receptors; ligand binding

18

KLAUS VON DER MARK

triggers intracellular signaling pathways regulating expression and activity of transcription factors, and in the cytoplasm and nucleus complex interactions of transcription factors with promoter, enhancer, or silencer elements regulate expression of collagen genes [21, 304, 321, 322]. Since the deciphering of the first collagen cDNA sequences, coding for proα2(I) and proα1(I) mRNA [323–325], the gene structure of 42 vertebrate collagen genes has been resolved. Cis-acting regulatory domains and the corresponding trans-acting factors are, however, known only for few collagens genes, predominantly in the fibrillar collagens, collagens IV, VI, and X, which will be briefly summarized in the following section.

A. Genes Coding for Fibrillar Collagens The genes coding for fibril-forming collagens are rather conserved and characterized by numerous short exons interrupted by fairly long introns. The triple helices of the genes of α1(I) (COLlA1 for the human gene, Col1a1 for the murine), of α2(I) (Col1a2), α1(II) (Col2a1), α1(III) (Col3a1), and all three chains of type V and XI collagen are encoded by 44 short exons, flanked at the 5′-end by two longer exons coding for the N-propeptide, and at the 3′-end by four exons coding for the conserved C-propeptide and 3′-untranslated regions [19]. In contrast, most of the nonfibril-forming collagens and invertebrate collagens are composed of fewer and larger exons. The exons coding for triple helical sequences are all multiples of 9 bp corresponding to a Gly-Xaa-Yaa-triplet; most common are 45, 54, and 109 bp [18, 19, 326]. This feature is unique for fibril-forming collagens and suggests that the fibrillar collagen genes have evolved from an ancestral multi-exon gene coding for collagen triplets. In contrast to the highly conserved size and number of exons in fibrillar collagen genes, there is considerable variation in the intron lengths between species and collagen types. For example, the gene COL1A1, coding for human pro α1(I), has a total length of 18 kb [327, 328] while the gene size COL1A2 for α2(I) is 40 kb [328] . All genes coding for human fibrillar collagens, even those belonging to the same type, are located on different chromosomes (for overview see [19, 329]). 1. Regulatory Domains and Transcriptional Regulation

As in most regulated genes, transcription of collagen genes is regulated by multiple cis-acting elements located in the proximal promoter and by the tissue-specific action of enhancer and silencer elements located further upstream of the promoter or in the introns. Identification of these

elements in collagen genes has led in the past to the identification of DNA binding factors responsible for tissuespecific regulation of collagen gene expression, including the identification of transcription factors such as cbfa1/ runx2 or SOX9. At the same time, increasing knowledge has accumulated on growth factors, hormones, and cytokines and their receptors in the regulation of collagen gene expression. One of the current challenges is to bring together the hormone- and growth factor-regulated signaling pathways and the transcriptional mechanisms regulating tissuespecific expression of collagen genes. 2. Type I Collagen Genes

a. Col1a1 Genes Studies by Sokolov et al. [330, 331], and Rossert et al. [332, 333] indicate that for human COL1A1 expression in skin a minimal promoter is required which is located within −476 bp and −900 bp upstream of the transcription start site of the COL1A1 gene. Although in transgenic mice the −476 bp promoter seems sufficient for tissuespecific expression even without intron sequences [331], further positive and negative regulatory sequences were identified in the promoter and in the first intron of the COLA1 genes [322, 334]. Slack et al. [335] reported that a reporter gene construct containing a 440-bp segment of the human COL1A1 promoter lacked tissue specificity without the first intron. Several cis-acting intronic sequences with a highly conserved 29-bp domain involved in transcriptional regulation were located between +292 and +670 bp [336]. Between +820 and +1093 bp a silencer element is active when combined with a 332-bp promoter fragment. The effect of the intronic sequences varies with their size, location, and orientation within the reporter gene constructs, and the cell type used for transfection. First evidence for a key role of the first intron in fibroblast-specific expression of Col1a1 was obtained from the MoV13 mouse mutant which was generated by infection with the Moloney virus [103, 104]. Accidental insertion of the virus into the first intron of the Col1A1 gene caused a complete block of Col1a1 transcription in fibroblasts. Mice homozygous for the insertion of the virus did not contain any type I collagen and died at day 13.5 due to heart rupture [103, 104]. Interestingly, however, the α1(I) was expressed in tooth and bone anlagen when cultured in chorioallantoic membranes in vitro, thus bypassing the termination of bone differentiation in vivo at day 13.5. This indicated a different transcriptional regulation in osteoblasts and odontoblasts as compared to fibroblasts [87, 106]. In fact, Rossert et al. [332, 333] located a bone-specific enhancer element between −1656 and −1540 bp upstream of the transcription

CHAPTER 1 Structure, Biosynthesis and Gene Regulation of Collagens in Cartilage and Bone

start site which promotes expression in osteoblasts. The transcription factor which binds to this element remains to be elucidated; it is apparently not cbfa1 which binds to the OSe2 consensus sequences located in the proximal promoter of Col1a1 [337] (see below). Another bonespecific site may be the Vit.D3-responsive element within the 3.6 kb promoter sequences of rat Col1A1, which is down-regulated by Vit.D3 in ROS osteoblasts, but not in fibroblasts [338]. Similarly, glucocorticoids which have been shown to down-regulate collagen synthesis also decrease expression of Col1a1 promoter-reporter gene constructs [339, 340]. b. Col1a2 Genes Similar to the Col1a1 gene, regulatory sequences have been described in the first intron of Col1a2 between +418 and +524 bp upstream of the transcription start site [341]. Curiously, the intron of human COL1A2 suppressed reporter gene transcription in chicken fibroblast, while the murine intron enhances transcription in mouse NIH 3T3 fibroblasts [341]. A −2000 bp Col1a2 promoter-reporter gene is active in transiently transfected osteoblasts at −300 bp. TGFβ is one of the major factors stimulating type I collagen synthesis in fibroblasts (see below) [342]. A TGFβ-activating element (TAE) with a nuclear factor I-like sequence was identified about 1600 bp upstream of the Col1a1 transcription start site [343]. A NF-1 binding site mediates the inhibitory effect of TGFβ on the α2(I) promoter [344]. There are also data showing that an AP-1 site in the proximal promoter of COL1A2 is responsible for the regulation of α2(I) expression by TGFβ [345], while other data indicate that the transcription factor Sp1 but not Ap1 is required for the early response of COL1A2 expression to TGFβ [346]. A proximal element within the human α2(I) collagen promoter located between −161 to −125 bp relative to the transcription start site is responsible for down-regulation of Col1a2 promoter activity by interferon γ (IFNγ) [347]. This IFNγ-response element (TaRE) is located between −265 and −241 bp. A potent far upstream enhancer element between 15 and 17 kb 5′ of the Col1a2 gene strongly increased the expression of a reporter gene in transgenic mice in dermis, fascia, and other fibrous tissue [348]. The Col1a2 gene contains an alternative, cartilagespecific transcription start site with a 179-bp promoter in the third intron, which is transcribed in cartilage into a mRNA with an open reading frame, but does not translate into a collagenous protein [306, 307]. The role of this alternative transcript which is rapidly turned over in cartilage is still unclear. Of particular interest would be the mechanism controlling the shift from the use of the fibroblast-specific promoter of pro α2(I) in chondroprogenitor cells to

19

the use of the cartilage-specific promoter in differentiated chondrocytes. 2. Type II Collagen Genes

Unlike type I or III collagen, the expression of type II collagen is restricted to only a few tissues, predominantly to cartilage, vitreous humor, embryonic cardiac jelly, and some embryonic epithelia [71, 90]; therefore, the tissuespecific regulation of Col2a1 expression has received particular interest in the past. Horton et al. [349] observed that CAT reporter gene constructs containing only Col2a1 promoter fragments upstream of the transcription start site showed low expression levels, which was significantly increased when an 800-bp segment of the first intron was included. Cartilage-specific enhancer elements were located within a 182-bp sequence in the first intron of the murine Col2a1 gene [350] or a 119-bp fragment of the rat Col2a1 gene [351]. The enhancer element consists of three HighMobility Group (HMG) binding sequences located within a 48-bp element between +1878 to +2345 bp in the first intron [350, 352]. Four tandem repeats of this 48-bp element or 12 tandem copies of an 18-bp sub-element of this enhancer, in combination with a 309-bp Col2a1 promoter, allowed cell-specific expression of a β-galactosidase reporter gene in mouse chondrocytes [353]. The critical role of this enhancer is underlined by the finding that cartilagespecific expression of Col2a1 enhancer-reporter genes in transgenic mice even occurs when the 309-bp Col2a1 promoter is replaced by a minimal β-globin promoter [352]. The localization of this enhancer element allowed the identification of SOX-9 as a major transcription factor regulating type II collagen expression in chondrocytes [354] (see also below). Furthermore, a zinc-finger protein CII2FP was described, which binds to the enhancer within the first intron of Col2a1 and to Sp1 [355], but the role of this protein in the control of Col2a1 expression remains to be elucidated. The complexity of the transcriptional regulation of the type II collagen gene is underlined by the finding of two silencer elements in the rat Col2a1 promoter which seem to inhibit Col2a1 expression in nonchondrocytic cells [355]. 3. TYPE III COLLAGEN GENES

In the mouse Col3a1 gene two positive and one negative regulatory domains were located in the proximal promoter region. A 12-bp stretch binding at −122 to −106 bp to an AP-1-like protein enhances transcription in DNAtransfection assays, and a 22-bp segment B (−61 to −83 bp upstream of the transcription start site) binds a nuclear factor (BBF) extracted from Hela cells [356]. A negative regulatory element located between −350 and −300 bp repressed the activity of the Col 3a1 promoter [357].

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KLAUS VON DER MARK 4. Type V and XI Collagen Genes

In contrast to the genes of type I, II, and III collagen, the promoter of the human of α1(V) collagen gene COL5A1 [358] lacks obvious TATA and CAAT boxes and has a number of features characteristic of the promoters of housekeeping genes [359]. It contains multiple transcription start sites, a high GC content and a number of consensus sites for the potential binding of the transcription factor SP1. The promoter seems to consist of an array of cis-acting elements, with a minimal promoter 212 bp upstream of the major transcription start site. The COL5A2 gene seems more closely related to the COL3A1 gene, but little is known of its regulating sequences [360]. Expression of the human COL5A3 gene is regulated by the transcription factor CBF/NF-Y binding to two repressors in the TATAless core promoter [361]. Similar to the COL5A1 gene, the genes of the human type XI collagens COL11A1 and COL11A2 lack TATA boxes, indicating that they are regulated similarly to housekeeping genes [362, 363]. The human Col11A1 gene has several minor transcriptional start sites clustered around one major start site at 318 bp upstream of the ATG codon [362, 363]. A transcriptional complex containing potential binding sites for ubiquitous nuclear proteins such as AP2 and Sp 1 is closely related to a similar complex in the proximal promoter of the human COL11A2 gene. The COL11A2 gene spans 30.5 kb with a minimum of 62 exons. Similarly to COL5A2, its promoter is GC-rich with two potential Sp1binding sites [167, 364]. A chondrocyte-specific enhancer element in the murine Col11a2 gene which binds Sox9 and resembles the Col2a1 enhancer confers cartilage-specific expression [365].

B. Type VI Collagen Genes The genes of the three type VI collagen subunits are composed of more than 60 small exons that are multiples of nine bp similar to those of the fibril-forming collagens. Of these, 19 exons in the range between 27–90 bp code for the triple helical part of chicken and human α1(VI) and α2(VI) [216, 366, 367]. Multiple splice variants have been described for α2(VI) and α3(VI) in chicken and human: in the chicken α2(VI) the major splice variant contains a vWFA domain at the carboxyterminal globule which is replaced by a FN-type III repeat in the minor splice variant expressed only in aortic tissue [368]. The human α2(VI) exists in three splice variants owing to alternative splicing at the C-terminus [216]. The chicken α3(VI) shows extensive size heterogeneity, with five bands in SDS-PAGE due to alternative splicing of vWFA

modules in the globular domain [369]. The α3(VI) splice variants show different tissue distribution depending on age and developmental stage [215]. In the chicken Col6a1 gene two major and several minor transcription sites were found [370]. The proximal promoter which lacks TATA or CAAT boxes contains three regulatory C-GGAGGG (GA box) boxes that bind several transcription factors including Sp1, AP1 [371, 372], and AP2 [373]. These elements are responsible for up-regulation of α1(VI) expression during C2C12 myoblast differentiation [371]. Expression of the murine Col6a1 promoterreporter gene constructs in transgenic mice indicated that the −215 bp minimal promoter is not sufficient for expression in tissue [370, 374]. Additional cis-acting elements between −0.4 to 1.4 kb are required for tissue expression. The cis-acting elements at –4.6 to −5.4 kb are responsible for α1(VI) expression in joints, while the region at −6.2 to −7.5 kb upstream of the transcription start site is required for expression in muscle, meninges, and other tissues except lung and uterus [370, 374]. Two promoters control the transcription of the human COL6A1 gene [373]. The chicken Col6a1 collagen promoter encompasses four regulatory sites; S1, S2, and S3 are recognized by transcription factors of the Sp1-family, while site X binds a 43 kDa transcription factor [375]. This type of cis-acting regulatory element found in the type VI collagen genes differs completely from those found in the genes of fibril-forming collagens. This is consistent with the distinct regulation of type VI collagen expression in tissues and cell culture in comparison to collagens I, II, or III. All three type VI collagen genes are up-regulated in fibroblasts when grown at increasing densities in culture, while Col I and III levels do not change [376]. Conversely, the phorbolester and cocarcinogen PMA reduces Col I and III levels, but not Col VI expression. Similar to Col I and III, however, Col VI expression is increased in systemic sclerosis and in keloids [377]. A strong up-regulation of collagen VI expression has been observed during chondrocyte differentiation [226]. Only in one study expression of α1(VI) in osteoblast-like cells been documented; it is up-regulated by interleukin 4 [378]. Interestingly, the human COL6A1 gene is not always coregulated with COL6A2 or COL6A3. Only α3(VI) expression, but not α1(VI) nor α2(VI), are up-regulated by TGFβ [379] or down-regulated by IFNγ [218] .

C. Type X Collagen Genes Unlike genes coding for fibril-forming collagens or type VI collagen chains, the genes for type X collagen contain only three exons that are fairly conserved in all

CHAPTER 1 Structure, Biosynthesis and Gene Regulation of Collagens in Cartilage and Bone

four species investigated: chicken [380], mouse [381–383], bovine [384], and human [385, 386]. Exon 1 of Col10a1 contains an 80–114 bp 5′ UTR sequences, exon 159–169 bp, while exon 3 with 2136–2900 bp codes for the majority of the triple helical part, the entire C-terminal NC1-1 domain and up to 939 (human) 3′ UTR sequences [387]. Nuclear run on experiments indicate that collagen X expression is regulated strictly at the transcriptional level [388]. As expected from the restricted temporal and spatial expression of type X collagen in the fetal cartilage growth plate [389, 390], strong positive and negative regulatory cisacting elements have been described in the promoter of collagen X genes, with major differences, however, between mammalian and chicken genes [387]. In the chicken, multiple negative elements in the proximal promoter (−580 bp) restrict expression of the Col10a1 gene to hypertrophic chondrocytes [391–393]. Also, a positive regulatory BMP-responsive 529-bp element was located 2.4–2.9 kb upstream of the Col10a1 transcription start site, which was found necessary and sufficient for BMP-stimulated Col10a1 expression in prehypertrophic chicken chondrocytes [394]. In the human COL10A1 promoter a strong enhancer element located between −1821 and −2400 bp upstream of the transcription site [395–397] stimulated transcription of reporter gene constructs in hypertrophic chondrocytes. In the mouse Col10a1 promoter an additional 2000-bp segment containing LINE elements, B1 and B2 and LTR elements, are inserted [398] moving the enhancer further upstream. This enhancer is highly homologous to the human and bovine enhancer and promotes specific expression of a COL10A1 reporter gene in hypertrophic cartilage both in in vitro reporter gene assays and in transgenic mice in vivo [398]. The enhancer contains an AP-1 site that is 100% conserved in mouse, human, and bovine Col10a1 enhancers. In vitro it is recognized by the AP-1 factors Fos B and Fra 1 [398]. Expression of type X collagen is down-regulated by PTH and PTHrP [397, 399] via the adenylate cyclase and PKA pathway which activates c-fos expression [397]. The proximal promoter regions of the Col10a1 genes contain a TATA box, one ETS binding site and an AP-2 binding site that are conserved between mouse, bovine and human species [386, 387]. Data by Harada et al. [400] indicate that the murine Col10a1 promoter is up-regulated by osteogenic protein 1, which is mediated by a MEF2-like sequence. The murine Col10a contains a second TATA box, and correspondingly, a second transcription start site, which is utilized only in the newborn mouse [382], while CCAAT boxes at −120 bp were found so far only in the mammalian Col10a1 genes. Furthermore, the proximal promoter of the murine Col10a1 gene contains OSE2 binding sites [401].

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VII. FACTORS REGULATING COLLAGEN BIOSYNTHESIS A. Bone Collagen Regulation With collagens providing 90% of the organic mass of bone, many mechanisms and factors regulating growth, turnover, or repair of bone are likely to be involved in the regulation of type I and V collagen synthesis by osteoblasts or their turnover by osteoclasts and proteases. As shown above, collagen biosynthesis in bone is regulated at transcriptional, translational, and post-translational level. Here I will review briefly current knowledge on the role of growth factors, hormones, cytokines, oncogenes, and transcription factors in regulating collagen synthesis, but additional chapters in this volume are devoted to collagen turnover by proteases and the role of growth factors, hormones, and cartilage on other aspects of bone metabolism. In view of the ubiquitous distribution of type I and V collagen in the vertebrate organism, most factors regulating these collagens are not tissue specific. Recently, however, two bone specific transcription factors Cbfa1/ runx2 [402–404] and osterix [405] have been discovered which bind to distinct cis-acting elements in the promoters of type I collagen and other bone-specific genes. 1. Growth Factors

TGFβ factors are one of the longest-known and bestdescribed factors regulating expression of collagen types, although their specific effects on collagen expression seem to vary considerably in different cell culture systems. Generally, TGFβ1 stimulates type I and III collagen synthesis in fibroblasts, osteoblasts, hepatocytes, and many other cell types [342]. Incubation of cells with TGFβ1 in vitro causes enhanced α1(I) mRNA levels by stimulating gene transcription and by supporting the stability and translational activity of collagen mRNA [308, 406]. TGFβ3 stimulates collagen synthesis both in TGFβ1-dependent and -independent pathways [407] (see Table II). In hepatocytes, TGFβ-stimulated transcription of COL1A2 involves binding of the transcription factors SP1 and Smad 2, one of the TGFβ receptor regulated Smad factors, to the proximal promoter of the COL1A2 gene [408, 409]. In human skin fibroblasts, Smad-3 transmits the TGFβ signals from the receptor to the COL1A2 promoter, while Smad 7, an inhibitory Smad factor, may be involved in autocrine negative feedback in the regulation of COL1A2 promoter activity by TGFβ [410]. Bone morphogenic proteins (BMPs), in particular BMP-2, -3, -4, and -7, which belong to the TGFβ superfamily, induce bone formation in embryonic and adult stem cells, mostly via enchondral ossification, but also directly

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KLAUS VON DER MARK Table II.

Cytokines, growth factors, oncogenes and transcription factors modulating collagen biosynthesis

Factor

Collagens affected

TGFβ1

I, III

TGFβ3

II II VI α1(I)

BMP -2, -3, -4, -7

I (?)

PDGF EGF IGF-I

II; X V I II

Il-1

I, III

IFNγ TNFα

II I II I

Vit.D3 Glucocorticoid

I α1(I)

PTH Retinoic acid

I X I, III

Ascorbate

X All collagens

c-fos

I

Cbfa1/Runx2

I X

SOX9

II, IX, XI

Cellular targets

Type of response

Growth factors Fibroblasts, osteoblasts Increases α1(I) + α1(III) mRNA level & stability, hepatoblasts, dediff. enhances translational activity of α2(I) mRNA chondrocytes Chondrocytes Suppression of Col2a1 expression Chondrocytes Up-regulation of Col2a1 expression Fibroblasts Up-regulates α3(VI), down-regulates α1 + α2(VI) Fibroblasts Stimulates α1(I) mRNA levels; down-regulates in presence of TGFβ1 Osteoblasts Induces osteoblast differentiation, stimulation of collagen I Mediated through smads and cbfa1β Chondrocytes Up-regulation of col2a1 expression in chondrocytic lines Gingival fibroblasts Stimulation of collagen protein synthesis Osteoblasts Inhibits collagen synthesis Chondrocytes Stimulates Col2a1 synthesis

Dediff. chondrocytes, fibroblasts Chondrocytes Fibroblasts Chondrocytes Fibroblasts

Cytokines Enhanced mRNA levels Suppresses CO2A1 transcription Suppression of collagen synthesis + mRNA levels Suppression of α1(II) mRNA levels Suppression of α1(I) mRNA

Hormones and other factors Osteoblasts Decreases α1(I) mRNA and protein levels Calvaria fibroblasts Suppresses collagen production in long-term cultures, reduced transcription rate of α1(I), multifactorial complex effects Fetal rat calvaria Suppression of α1(I) mRNA levels Hypertr. chondrocytes Reduced α1(X) mRNA levels Chondrocytes Induces switch from α1(II) to α1(I), α1(II), and α1(III) mRNA in chondrocytes Chondrocytes Transient stimulation of α1(X) expression Fibroblasts, Enhances prolyl and lysyl hydroxylation chondrocytes, stimulation of α2(I) mRNA transcription, enhanced osteoblasts, etc. mRNA stability Transcription factors and oncogenes c-fos overexpression inhibits α1(I) mRNA levels in osteoblasts Osteoblasts Induces transcription of Col1a1 in osteoblasts by binding to OSE2 elements in the promoter Hypertr. chondrocytes Stimulates Col10a1 expression by binding to OSE2 elements in the Col10a1 promoter Chondrocytes Induces Col2a1 and Col11a1 expression by binding to enhancer Osteoblasts

by membranous bone formation [411–413] (see also Chapter 6). BMP-induced osteoblast differentiation involves the Smad signaling pathway and receptors of the BMPRI group [414]. In BMP-treated cell and organ cultures the induction of osteogenesis is associated with a steep increase on type I collagen synthesis, as a result of enhanced osteoblast differentiation and proliferation. Recently Takagi et al.

Reference 308, 342, 406 408, 409, 410 461, 462 459, 460 379 407 411–415

466–468 416 417 482

439 490, 491 439 492 440–443

338, 418, 419 421, 422, 426

420 397, 488, 489 474–476 474, 477 494

419, 437 402, 403 430–432 401, 433 450–452

[415] have shown that BMP-2 induces the expression of osteogenic transcription factors such as Cbfa1/runx2 (see below), osterix, AJ18 or Msx2. There is no evidence, however, for a stimulation and direct regulation of type I collagen synthesis by BMPs as is the case with TGFβ. Another growth factor which has been shown to enhance collagen type I synthesis in gingival fibroblasts is PDGF

CHAPTER 1 Structure, Biosynthesis and Gene Regulation of Collagens in Cartilage and Bone

[416], while EGF inhibits collagen type I synthesis in osteoblasts [417]. 2. Steroid Hormones

Despite its essential role on bone mineralization, vitamin D3 (1,25-dihydroxycholecalciferol or 1,25(OH)2D3) inhibits type I collagen synthesis in vitro by suppressing the transcription of Col1a1 and Col1a2 genes [338, 418, 419]. Also, parathyroid hormone (PTH), another factor involved in calcium metabolism of bone, has been shown to inhibit type I collagen expression [420]. Glucocorticoids generally suppress synthesis of collagens in fibroblasts [421, 422] and in cartilage [423]. In calvarial bone dexamethasone suppresses collagen production in long-term cultures, but after an initial phase of stimulation [424, 425]. The effect of glucocorticoids can be complex in the presence of other factors, e.g. it potentiates the stimulatory effect of IGF-I on collagen synthesis [425, 426]. Estrogen, however, has been shown to stimulate type I collagen in osteoblasts predominantly in intermediate stages of cell differentiation [427]. 3. Transcription Factors and Oncogenes

A number of transcription factors are involved in controlling differentiation of preosteoblasts to osteoblasts. Dlx5, a homeodomain protein, is one of several Dlx proteins that are expressed in different craniofacial regions and in the limb during development [428]. Overexpression of Dlx5 in osteoblasts from chicken calvaria enhanced expression of type I collagen [429]. Cbfa1/Runx2 has been identified as a Runt-related, osteoblast-specific transcription factor which is essential for osteoblast differentiation and regulation of osteoblastspecific genes such as osteocalcin, osteopontin, and Col1a1 and Col1a2 genes [402, 403, 430–432]. Cbfa1 knockout mice initially develop a nearly complete skeleton. Endochondral and membraneous ossification are, however, completely impaired, indicating that, in the absence of Cbfa1, osteogenic differentiation is blocked at the nonossified preosteoblastic stage. The role of Cbfa1 in the regulation of bone-specific gene expression was shown by its specific binding to consensus sequences in the promoters of the mouse osteocalcin and Col1a1 genes [402]. A consensus Cbfa1 binding site called OSE2 is located at approximately –1350 bp in the promoter of human, mouse, and rat Col1a1 genes [337], but additional OSE2 elements are located also in the proximal promoter. Of several OSE2 elements in the Col1a2 promoter, only one is conserved in other species; this elements binds Cbfa1 and confers osteoblast-specific activity to the minimum Col1a1 promoter in reporter gene assays [337]. In view of its early expression in skeletal development, Cbfa1 seems

23

also involved in the regulation of genes that are expressed before overt osteogenic differentiation [337]. Furthermore, cbfa1/runx2 is one of the major factors regulating expression of type X collagen in hypertrophic chondrocytes [401, 433]. Recently, another osteogenic transcription factor called osterix, which belongs to the zinc finger family, has been identified. It is essential for bone formation [405], but acts downstream of Cbfa1. Similar to Cbfa1 null mice, osterix-deficient mice develop a cartilaginous skeleton with impaired endochondral ossification and complete absence of membraneous ossification. Osterix stimulates type I collagen gene expression in C2C12 cells [405], but it is nuclear whether it binds directly to regulatory elements in the Col1a1 gene. Also, the nuclear matrix seems involved in the regulation of type I collagen gene expression in osteoblasts. In reporter gene assays using UMR-106 osteoblasts it was shown that NP/NMP4 nuclear matrix proteins bind to a 3.5 kb promoter fragment of the human COL1A1 gene [434]. There is ample evidence for a key role of the AP-1 factors c-fos and Fos B in osteoblast differentiation [435, 436], but only few data are available on the regulation of type I collagen in osteoblasts by Ap-1 factors. Kuroki et al. [419, 437] have shown that overexpression of c-fos in the osteoblast cell line MC3T3-E1 inhibits α1(I) mRNA expression. Interestingly, vitamin D, which also inhibits type I collagen synthesis (see above), reverts the inhibitory effect of c-fos to a level of control cells, indicating inhibition of vitamin D binding to the vitamin D receptor [419]. 4. Cytokines

Most inflammatory cytokines such as interleukin-1β or interferon-α inhibit type I collagen synthesis [438]. Suppression of transcription of COL1A2 in hepatocytes by IFNα is exerted through the interaction of p300 with Stat1 [439]. Also, IFNγ plays an important physiological role in inflammation by down-regulating collagen gene expression. Interferon γ (IFNγ) and TNFα both suppress type I collagen mRNA levels in fibroblasts [440–443]. In analyzing the mechanism of IFNγ-induced collagen expression in lung fibroblasts, a trimeric DNA-binding complex named RFX5 was identified which is induced by IFNγ and binds to the Col1a1 transcription start site, thus repressing collagen expression [444, 445]. 5. Regulation of Type V Collagen

Little is known on factors which specifically regulate type V collagen synthesis. There is, however, experimental evidence from DNaseI footprinting and electrophoretic mobility shift, as well as from chromatin immunoprecipitation assays, that the CCAAT-binding

24

KLAUS VON DER MARK

transcription factor CBF/NF-Y regulates the core promoter of the human COL5A1 gene [361].

B. Cartilage Collagen Regulation 1. SOX9

Wright et al. [446] reported on the expression of SOX9, a transcription factor of the SRY (sex-determining region, Y-chromosome) family, in cartilaginous anlagen of the mouse embryo. In situ hybridization studies showed that Sox9 expression parallels expression of the Col2a1 gene in all chondroprogenitor cells and in early, but not hypertrophic, chondrocytes [447, 448], suggesting that it may be involved in chondrogenic development. Furthermore, mutations in the human Sox9 gene cause severe skeletal malformations in patients with campomelic dysplasia [449]. SOX9 has a DNA-binding domain that is homologous to the HMG (High Mobility Group) DNA-binding domain of the SRY gene [450]. Lefebvre et al. [451] and Bell et al. [452] finally demonstrated that SOX9 is a potent activator of Col2a1 gene expression, as it binds to a HMG box consensus sequence CATTCAT present in the 48-bp enhancer of the Col2a1 gene and strongly enhances transcription of Col2a1 reporter genes [354]. In mouse chondrocytes, high levels of SOX9 protein correlate with type II collagenexpressing cells, while dedifferentiated as well as hypertrophic chondrocytes lack SOX9 [354]. The strong chondrogenic potency of SOX9 is further supported by the observation that ectopic expression of SOX9 in transgenic mice activated Col2a1 expression in noncartilaginous tissues [452]. Transactivation of the Col2a1 enhancer by SOX9 is enhanced after phosphorylation of SOX9 by cAMP-dependent protein kinase A [453]. The key role of SOX9 in chondrogenesis is underlined by the fact that haploinsufficiency of SOX9 in Sox9 −/+ heterozygous mice results in defective cartilage primordia [454]. Sox9 gene deletion is lethal in the homozygotes, while in Sox9 −/− chimera all Sox9 −/− cells are excluded from cartilage [455]. SOX9 not only activates transcription of Col2a1, but also of other cartilage matrix genes such as Col11a1 [365], Col9a1, and aggrecan. The transcriptional complex includes also SOX5, L-SOX5, a larger splice variant, and SOX6, which are also essential for chondrogenic development, but cannot activate collagen type II gene expression in the absence of SOX9 [456]. 2. BMP and TGFb Family Proteins

TGFβ, which is the main stimulator of type I collagen expression, has also been shown to induce chondrogenic differentiation in rat periosteum in vivo and in vitro [457].

Conflicting reports exist, however, concerning the effect of TGFβ on type II expression in mature chondrocytes. Redini et al. [458] and Galera et al. [459, 460], have shown up-regulation of type II collagen expression in rabbit articular chondrocytes by TGFβ, while others report that TGFβ1 inhibits type II collagen [461]. Inhibition occurs in a protein kinase C-dependent manner and involves regulatory sequences in the promoter and first intron [462]. Bone morphogenic proteins have been shown to induce chondrogenic differentiation in limb bud mesenchymal cells [463, 464], in periosteal stem cells [465] and in various undifferentiated stem cell lines such as C3H10T1/2, ATDC5, and others [411, 466–468]. As a result of chondrogenic differentiation, type II collagen is up-regulated, and there are also data indicating that type II collagen expression in adult articular chondrocytes in agar cultures is stimulated by BMP-2. In reporter gene studies using MC615 chondrocytes and C3H10T1/2 cells, it was shown that Smad1 and 5 are involved in transactivation of Col2a1 reporter genes after stimulation with BMP-4 [469]. The target genes of Smad1 or 5 that are responsible for upregulation of type II collagen expression have not been identified yet, but several lines of evidence indicate that SOX9 is one of them. BMP responsive cis-acting elements have not been described in the type II collagen gene, but in the type X collagen gene. Several BMP factors, including BMP-4, -6, and -7, promote differentiation of proliferating to hypertrophic chondrocytes [399, 470, 471] as indicated by a steep increase in type X collagen expression. Several functional BMP-responsive elements [472] are, however, not active in the human or bovine COL10A1 gene. 3. Retinoic Acid

Retinoic acid (RA) has rigorous effects on both chondrogenic differentiation and on the stability of the chondrocyte phenotype, dependent on the RA dosis [473, 474]. At high concentrations (10–7 M) RA causes chondrocyte dedifferentiation, associated with a switch from type II to type I and III collagen expression [474, 475]. At low doses (10−9–10–8 M) RA stimulates chondrogenic differentiation of limb mesenchymal cells [476] and induces hypertrophic differentiation, including transient up-regulation of type X collagen expression in chondrocytes [474, 477]. RA inhibits limb chondrogenesis and causes malformation at high doses [478, 479]. There is, however, no information on direct interactions of RA receptors with collagen promoter sequences. The RA effect on type X collagen expression may be regulated through the induction of runx2/cbfa1, which not only induces osteogenic differentiation and the expression of bone-specific genes, but also

CHAPTER 1 Structure, Biosynthesis and Gene Regulation of Collagens in Cartilage and Bone

up-regulates expression of type X collagen in chicken and mouse chondrocytes by binding to OSE2 elements [401]. 4. Cytokines and Growth Factors Regulating Cartilage Collagen Synthesis

Insulin-like growth factor I (IGF-I) is one of the main growth factors regulating cartilage growth during skeletal development. It stimulates chondrocyte proliferation, but also proteoglycan synthesis [480, 481] and type II collagen synthesis [482]. FGF2 (= bFGF) inhibits chondrocyte differentiation [483], but no information is available on direct inteference with collagen gene expression. A key hormone regulating chondrocyte differentiation to hypertrophic cells is PTHrP (parathyroid hormonerelated peptide) which is synthesized in the perichondrium and prehypertrophic chondrocytes [484, 485]. Overexpression of PTHrP in transgenic mice delays endochondral ossification and bone development by stimulating proliferation and inhibiting differentiation of chondrocytes in the proliferative zone of the growth plate [486], while premature chondrocyte hypertrophy, including early onset of type X collagen, is observed in PTHrP-deficient mice [487]. In vitro, PTHrP inhibits expression of type X collagen in rabbit, chicken, and bovine hypertrophic chondrocytes [397, 488, 489]. This inhibition is mediated by c-fos, which is induced by PTHrP and involves a PTHrP-sensitive site located in the COL10A1 enhancer [397]. This enhancer contains an AP1 site, which is however not recognized by c-Fos, but by Fos B and Fra1 factors which stimulate the expression of Col10A1 reporter genes [398]. Interleukin 1 (IL-1) increases type I and III collagen mRNA levels in cultures of human chondrocytes [490], probably owing to the presence of dedifferentiated chondrocytes, while it suppresses the α1(II) mRNA levels [490, 491]. IFNγ also reduces type II collagen mRNA levels in chondrocytes [492]. Ascorbic acid is an essential factor in collagen biosynthesis as it is involved in hydroxylation of proline and hydroxyproline (for review see [493]). But beyond this role, ascorbic acid has been shown to enhance collagen α2(I) mRNA levels six-fold and to enhance mRNA stability in chicken tendon fibroblasts [494].

VIII. CONCLUSIONS With growing knowledge on the variety of collagen types, their structure, physiological features, and the manifold ways of their metabolic regulation, we begin to understand not only the unique biomechanical properties of bone and cartilage, but also the mechanisms regulating

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their growth and turnover in development and diseases. In bone, densely packed and intensely cross-linked heterofibrils of type I and V collagen provide the architectural scaffold and the substrate for mineralization, and thus hold responsibility for the enormous resistance of bone to mechanical load, torsion, and tension. In hyaline cartilage, heterofibrils of collagen types II and XI, decorated with FACIT collagens, assemble into cross-linked meshworks which allow the incorporation of highly hydrated hyaluronan–proteoglycan complexes, thus providing the tissue with elasticity combined with high resistance to pressure. With better information on tissue-specific posttranslational modifications of collagens, including prolyl and lysyl-hydroxylation, glycosylation, sulfation, oxidation by lysyloxidases, and cross-linking, we begin to understand the unique biochemical and physiological properties of different collagen types in different tissues and may some day understand why, for example, types I or III collagen cannot replace the similar type II collagen in cartilage. A major extension in our understanding of the role of collagens in bone and cartilage was introduced by cell biological and transgenic mouse experiments, showing that collagens transmit – on top of their biomechanical function – specific information to cells by providing substrates for cell adhesion, proliferation, migration, arrangement in the tissue, differentiation, gene expression (e.g. of collagenases), or cell survival. Owing to recent discoveries of specific interactions of type I, II, III, and VI collagens with α1β1, α2β1, α11β1, and α10β1 integrins, the identification of the integrin-binding triple helical epitope GFQGER in type I collagen and the integrin-mediated signaling pathways, we are now in a position to elucidate entire molecular pathways of collagenregulated cell responses in bone and cartilage and other tissues. Impressed by the apparent eternal stability of mammoth bone discovered after 10000 years in Siberian ice, one should keep in mind, however, that bone and its collagens are subject to continuous renewal and turnover by osteoblasts and osteoclasts in the living organism. Numerous growth factors such as TGFβ, BMPs, FGFs, and IGF, steroid hormones such as vitamin D3, corticoids, and cytokines, are involved in the regulation of collagen biosynthesis and turnover in bone. The regulation of cartilage collagen metabolism by BMPs, IGFI, FGF, PTHrP, retinoic acid, and cytokines is not less complicated and varies in different cartilages and developmental stages. Boneand cartilage-specific transcription factors, such as cbfa1/ runx2 and osterix, or SOX9 for cartilage, respectively, were discovered which regulate not only the expression of

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KLAUS VON DER MARK

collagen genes, but also of other bone- and cartilagespecific genes. Elucidation of the complete pathways of bone- and cartilage-inducing growth factors, such as BMPs and their receptor-mediated regulation of tissuespecific expression of collagens, seems one of the current challenges in bone and cartilage research. This task is matched by the elucidation of the equally complex regulation of collagen degradation by collagenases and the regulation of collagenases by cytokines and cell–matrix interactions, by activation and extracellular inhibitors, as described in detail in Chapter 10. Modern genetic tools have unraveled numerous new collagens also in cartilage and bone. The majority of these collagens, including types XII, XIV, and XVI, XX and XXII, classified as FACIT collagens and a new fibril-forming collagen (type XXIV), are also expressed in cartilage. The functions of these collagens and also of the long-known collagens such as collagen VI in cartilage are, however, yet unclear. Collagen knockout experiments often failed to reveal a clear phenotype, which leaves the field with numerous challenging problems to be solved.

References 1. Boskey, A. L., Wright, T. M., and Blank, R. D. (1999). Collagen and bone strength. J. Bone Miner. Res. 14, 330–335. 2. Kielty, C. M., and Grant, M. E. (2002). The collagen family: structure, assembly, and organisation in the extracellular matrix. In Connective Tisssue and its Heritable Disorders (P. M. Royce, and B. Steinmann, ed.) pp. 159–221, Wiley-Liss, New York. 3. Bruckner, P., and van der Rest, M. (1994). Structure and function of cartilage collagens. Microsc. Res. Tech. 28, 378–384. 4. Eyre, D. (2002). Collagen of articular cartilage. Arthritis Res. 4, 30–35. 5. Niyibizi, C., and Eyre, D. R. (1994). Structural characteristics of cross-linking sites in type V collagen of bone. Chain specificities and heterotypic links to type I collagen. Eur. J. Biochem. 224, 943–950. 6. Mayne, R., and Brewton, R. G. (1993). New members of the collagen superfamily. Curr. Opin. Cell Biol. 5, 883–890. 7. Eyre, D. R., Wu, J. J., Fernandes, R. J., Pietka, T. A., and Weis, M. A. (2002). Recent developments in cartilage research: matrix biology of the collagen II/IX/XI heterofibril network. Biochem. Soc. Trans. 30, 893–899. 8. Ayad, S., Evans, H., Weiss, J. B., and Holt, L. (1984). Type VI collagen but not type V collagen is present in cartilage. Collagen Rel. Res. 4, 165–168. 9. Keene, D. R., and Engvall, E. G. W. (1988). Ultrastructure of type VI collagen in human skin and cartilage suggests an anchoring function for this filamentous network. J. Cell Biol. 107, 1995–2006. 10. Horikawa, O., Nakajima, H., Kikuchi, T., Ichimura, S., Yamada, H., Fujikawa, K., and Toyama, Y. (2004). Distribution of type VI collagen in chondrocyte microenvironment: study of chondrons isolated from human normal and degenerative articular cartilage and cultured chondrocytes. J. Orthop. Sci. 9, 29–36. 11. Poole, C. A., Ayad, S., and Gilbert, R. T. (1992). Chondrons from articular cartilage. V. Immunohistochemical evaluation of type VI collagen organisation in isolated chondrons by light, confocal and electron microscopy. J. Cell. Sci. 103, 1101–1110.

12. von der Mark, K., and Sorokin, L. M. (2002). Adhesive Glycoproteins. In Connective Tissue and its Heritable Disorders (P. M. Royce, and B. Steinmann, eds.), pp. 293–328, Wiley-Liss, New York. 13. Mayne, R., and Burgeson, R. E. (1987). Structure and Function of Collagen Types. In (R. P. Mecham, ed), Academic Press, Inc. Orlando, FL. 14. Bateman, J. F., Lamandé, S. R., and Ramskans, J. A. M. (1996). Collagen superfamily. In “Extrazellular Matrix.” (W. D. Comper, ed.), Vol. 2, pp. 22–67, Harwood Acad. Pulls. 15. van der Rest, M., and Garrone, R. (1991). Collagen family of proteins. Faseb. J. 5, 2814–2823. 16. Piez, K. A. (1998). Molecular and aggregate structure of the collagens. In “Extracellular Matrix Biochemistry” (K. A. Piez, and H. Reddi, eds), Elsevier. 17. Olsen, B. R. (1995). New insights into the function of collagens from genetic analysis. Curr. Opin. Cell. Biol. 7, 720–727. 18. Sandell, L., and Boyd, C. D. (1990). Extracellular Matrix Genes. Academic Press, San Diego, CA. 19. Chu, M. L., and Prockop, D. J. (2002). Collagen: gene structure. In “Connective Tisssue and its Heritable Disorders” (B. Steinmann, and P. M. Royce, eds.), pp. 223–248, Wiley-Liss, New York. 20. Kivirikko, K. I. (1998). Collagen biosynthesis: a mini-review cluster. Matrix Biol. 16, 355–356. 21. Bornstein, P., and Sage, H. (1989). Regulation of collagen gene expression. Prog. Nucleic. Acid. Res. Mol. Biol. 37, 67–106. 22. McLaughlin, S. H., and Bulleid, N. J. (1998). Molecular recognition in procollagen chain assembly. Matrix Biol. 16, 369–377. 23. Prockop, D. J., Sieron, A. L., and Li, S. W. (1998). Procollagen N-proteinase and procollagen C-proteinase. Two unusual metalloproteinases that are essential for procollagen processing probably have important roles in development and cell signaling. Matrix Biol. 16, 399–408. 24. Johnson, L. L., Dyer, R., and Hupe, D. J. (1998). Matrix metalloproteinases. Curr. Opin. Chem. Biol. 2, 466–471. 25. Murphy, G., and Reynolds, J. J. (2002). Extracelllular matrix degradation. In “Connective Tissue and its Heritable Disorders” (P. M. Royce, and B. Steinmann, eds.), pp. 343–384, Wiley-Liss, New York. 26. Myllyharju, J., and Kivirikko, K. I. (2001). Collagens and collagenrelated diseases. Ann. Med. 33, 7–21. 27. Prockop, D. J., and Kivirikko, K. I. (1995). Collagens: molecular biology, diseases, and potentials for therapy. Annu. Rev. Biochem. 64, 403–434. 28. Kuivaniemi, H., and Tromp, G. (1997). Mutations in fibrillar collagens (types I, II, III, and XI), fibril-associated collagen (type IX), and network-forming collagen (type X) cause a spectrum of diseases of bone, cartilage, and blood vessels. Hum. Mut. 9, 300–315. 29. Royce, P. M., and Steinmann, B. (2002). “Connective tissue and its Heritable Disorders,” Wiley Liss, New York. 30. Koch, M., Foley, J. E., Hahn, R., Zhou, P., Burgeson, R. E., Gerecke, D. R., and Gordon, M. K. (2001). Alpha 1(XX) collagen, a new member of the collagen subfamily, fibril-associated collagens with interrupted triple helices. J. Biol. Chem. 276, 23120–23126. 31. Koch, M., Laub, F., Zhou, P., Hahn, R. A., Tanaka, S., Burgeson, R. E., Gerecke, D. R., Ramirez, F., and Gordon, M. K. (2003). Collagen XXIV, a vertebrate fibrillar collagen with structural features of invertebrate collagens: selective expression in developing cornea and bone. J. Biol. Chem. 278, 43236–43244. 32. Koch, M., Schulze, J., Hansen, U., Ashwodt, T., Keene, D. R., Brunken, W. J., Burgeson, R. E., Bruckner, P., and Bruckner-Tuderman, L. (2004). A novel marker of tissue junctions, collagen XXII. J. Biol. Chem. 279, 22514–22521. 33. Chou, M. Y., and Li, H. C. (2002). Genomic organization and characterization of the human type XXI collagen (COL21A1) gene. Genomics 79, 395–401. 34. Banyard, J., Bao, L., and Zetter, B. R. (2003). Type XXIII collagen, a new transmembrane collagen identified in metastatic tumor cells. J. Biol. Chem. 278, 20989–20994.

CHAPTER 1 Structure, Biosynthesis and Gene Regulation of Collagens in Cartilage and Bone 35. Hashimoto, T., Wakabayashi, T., Watanabe, A., Kowa, H., Hosoda, R., Nakamura, A., Kanazawa, I., Arai, T., Takio, K., Mann, D. M., and Iwatsubo, T. (2002). CLAC: a novel Alzheimer amyloid plaque component derived from a transmembrane precursor, CLAC-P/collagen type XXV. Embo. J. 21, 1524–1534. 36. Sato, K., Yomogida, K., Wada, T., Yorihuzi, T., Nishimune, Y., Hosokawa, N., and Nagata, K. (2002). Type XXVI collagen, a new member of the collagen family, is specifically expressed in the testis and ovary. J. Biol. Chem. 277, 37678–37684. 37. Boot-Handford, R. P., Tuckwell, D. S., Plumb, D. A., Rock, C. F., and Poulsom, R. (2003). A novel and highly conserved collagen (pro(alpha)1(XXVII)) with a unique expression pattern and unusual molecular characteristics establishes a new clade within the vertebrate fibrillar collagen family. J. Biol. Chem. 278, 31067–31077. 38. Brodsky, B., and Ramshaw, J. A. (1997). The collagen triple-helix structure. Matrix Biol. 15, 545–554. 39. Berisio, R., Vitagliano, L., Mazzarella, L., and Zagari, A. (2002). Recent progress on collagen triple helix structure, stability and assembly. Protein Pept. Lett. 9, 107–116. 40. Mizuno, K., Hayashi, T., Peyton, D. H., and Bachinger, H. P. (2004). Hydroxylation-induced stabilization of the collagen triple helix. Acetyl-(glycyl-4(R)-hydroxyprolyl-4(R)-hydroxyprolyl)(10)-NH(2). forms a highly stable triple helix. J. Biol. Chem. 279, 38072–38078. 41. Hofmann, H., Fietzek, P. P., and Kuhn, K. (1978). The role of polar and hydrophobic interactions for the molecular packing of type I collagen: a three-dimensional evaluation of the amino acid sequence. J. Mol. Biol. 125, 137–165. 42. Berg, R. A., and Prockop, D. J. (1973). The thermal transition of a nonhydroxylated form of collagen. Evidence for a role for hydroxyproline in stabilizing the triple-helix of collagen. Biochem. Biophys. Res. Commun. 52, 115–120. 43. Schmid, T. M., and Linsenmayer, T. F. (1984). Denaturation-renaturation properties of two molecular forms of short-chain cartilage collagen. Biochem. 23, 553–558. 44. Risteli, J., Baechinger, H. P., Engel, J., Furthmayr, H., and Timpl, R. (1980). 7S-collagen: characterization of an unusual basement membrane structure. Eur. J. Biochem. 108, 239–250. 45. Kuehn, K., Wiedemann, H., Timpl, R., Risteli, J., Dieringer, H., Voss, T., and Glanville, R. (1981). Macromolecular structure of basement membrane collagen. FEBS Lett. 125, 123–128. 46. Bruckner, P., and Prockop, D. J. (1981). Proteolytic enzymes as probes for the triple-helical conformation of procollagen. Analyt. Biochem. 110, 360–368. 47. Kodama, T., Doi, T., Suzuki, H., Takahashi, K., Wada, Y., and Gordon, S. (1996). Collagenous macrophage scavenger receptors. Curr. Opin. Lipidol. 7, 287–291. 48. Elomaa, O., Sankala, M., Pikkarainen, T., Bergmann, U., Tuuttila, A., Raatikainen-Ahokas, A., Sariola, H., and Tryggvason, K. (1998). Structure of the human macrophage MARCO receptor and characterization of its bacteria-binding region. J. Biol. Chem. 273, 4530–4538. 49. Fessler, J. H., Doege, K. J., Duncan, K. G., and Fessler, L. I. (1985). Biosynthesis of collagen. J. Cell Biochem. 28, 31–37. 50. Fessler, J. H., Shigaki, N., and Fessler, L. I. (1985). Biosynthesis and properties of procollagens V. Ann. Acad. Sci. N.Y. 460, 181–186. 51. Fleischmajer, J. S., Perlish, J. S., and Olsen, B. R. (1987). Amino and carboxyl propeptides in bone collagen fibrils during embryogenesis. Cell Tissue Res. 247, 105–109. 52. Kadler, K. E., Hojima, Y., and Prockop, D. J. (1987). Assembly of collagen fibrils de novo by cleavage of the type I PC-collagen with procollagen C-proteinase – assay of critical concentration demonstrates that collagen self-assembly is a classical example of an entropy-driven process. J. Biol. Chem. 262, 15696–15701. 53. Hinek, A., Reiner, A., and Poole, R. (1987). The calcification of cartilage matrix in chondrocyte culture: Studies of the C-propetide of type II collagen (chondrocalcin). J. Cell Biol. 104, 1435–1441.

27

54. Bogin, O., Kvansakul, M., Rom, E., Singer, J., Yayon, A., and Hohenester, E. (2002). Insight into Schmid metaphyseal chondrodysplasia from the crystal structure of the collagen X NC1 domain trimer. Structure 10, 165–173. 55. Schmid, T. M., Cole, A. A., Chen, Q., Bonen, D. K., Luchene, L., and Linsenmayer, T. (1994). Assembly of type X collagen by hypertrophic chondrocytes. In “Extracellular Matrix Assembly and Structure” (P. Yurchenco, D. Birk, and R. Mecham, eds.), pp. 171–206, Academic Press. 56. Frischholz, S., Beier, F., Girkontaite, I., Wagner, K., Pöschl, E., Turnay, J., Mayer, U., and von der Mark, K. (1998). Characterization of human type X procollagen and its NC-1 domain expressed as recombinant proteins in HEK293 cells. J. Biol. Chem. 273, 4547–4555. 57. Shaw, L. M., and Olsen, B. R. (1991). FACIT collagens: diverse molecular bridges in extracellular matrices. Trends. Biochem. Sci. 16, 191–194. 58. Oh, S. P., Warman, M. L., Seldin, M. F., Cheng, S. D., Knoll, J. H., Timmons, S., and Olsen, B. R. (1994). Cloning of cDNA and genomic DNA encoding human type XVIII collagen and localization of the alpha 1(XVIII) collagen gene to mouse chromosome 10 and human chromosome 21. Genomics 19, 494–499. 59. Yamagata, M., Yamada, K. M., Yamada, S. S., Shinomura, T., Tanaka, H., Nishida, Y., Obara, M., and Kimata, K. (1991). The complete primary structure of type XII collagen shows a chimeric molecule with reiterated fibronectin type III motifs, von Willebrand factor A motifs, a domain homologous to a noncollagenous region of type IX collagen, and short collagenous domains with an Arg-Gly-Asp site. J. Cell Biol. 115, 209–221. 60. O’Reilly, M. S., Boehm, T., Shing, Y., Fukai, N., Vasios, G., Lane, W. S., Flynn, E., Birkhead, J. R., Olsen, B. R., and Folkman, J. (1997). Endostatin: an endogenous inhibitor of angiogenesis and tumor growth. Cell 88, 277–285. 61. Hagg, P., Rehn, M., Huhtala, P., Vaisanen, T., Tamminen, M., and Pihlajaniemi, T. (1998). Type XIII collagen is identified as a plasma membrane protein. J. Biol. Chem. 273, 15590–15597. 62. Pihlajaniemi, T., Tamminen, M. (1990). The alpha1 chain of type XIII collagen consists of three collagenous and four non-collagenous domains, and its primary transcript undergoes complex alternative splicing. J. Biol. Chem. 265, 16922–16928. 63. Gordon, M. K., Fitch, J. M., Foley, J. W., Gerecke, D. R., Linsenmayer, C., Birk, D. E., and Linsenmayer, T. F. (1997). Type XVII collagen (BP 180) in the developing avian cornea. Invest. Ophthalmol. Vis. Sci. 38, 153–166. 64. Farjanel, J., Schurmann, G., and Bruckner, P. (2001). Contacts with fibrils containing collagen I, but not collagens II, IX, and XI, can destabilize the cartilage phenotype of chondrocytes. Osteoarthritis Cartilage 9, S55–S63. 65. Tulla, M., Pentikainen, O. T., Viitasalo, T., Kapyla, J., Impola, U., Nykvist, P., Nissinen, L., Johnson, M. S., and Heino, J. (2001). Selective binding of collagen subtypes by integrin alpha 1I, alpha 2I, and alpha 10I domains. J. Biol. Chem. 276, 48206–48212. 66. Tiger, C. F., Fougerousse, F., Grundstrom, G., Velling, T., and Gullberg, D. (2001). Alpha11beta1 integrin is a receptor for interstitial collagens involved in cell migration and collagen reorganization on mesenchymal nonmuscle cells. Dev. Biol. 237, 116–129. 67. Jokinen, J., Dadu, E., Nykvist, P., Kapyla, J., White, D. J., Ivaska, J., Vehvilainen, P., Reunanen, H., Larjava, H., Hakkinen, L., and Heino, J. (2004). Integrin-mediated cell adhesion to type I collagen fibrils. J. Biol. Chem. 279, 31956–31963. 68. Aszodi, A., Bateman, J. F., Gustafsson, E., Boot-Handford, R., and Fassler, R. (2000). Mammalian skeletogenesis and extracellular matrix: what can we learn from knockout mice? Cell. Struct. Funct. 25, 73–84. 69. Aszodi, A., Chan, D., Hunziker, E., Bateman, J. F., and Fassler, R. (1998). Collagen II is essential for the removal of the notochord and the formation of intervertebral discs [In Process Citation]. J. Cell Biol. 143, 1399–1412.

28 70. Hagg, R., Hedbom, E., Mollers, U., Aszodi, A., Fassler, R., and Bruckner, P. (1997). Absence of the alpha1(IX) chain leads to a functional knock-out of the entire collagen IX protein in mice. J. Biol. Chem. 272, 20650–20654. 71. Cheah, K. S., Lau, E. T., Au, P. K., and Tam, P. P. (1991). Expression of the mouse alpha 1(II) collagen gene is not restricted to cartilage during development. Development 111, 945–953. 72. von der Mark, K., von der Mark, H., Timpl, R., and Trelstad, R. L. (1977). Immunofluorescent localization of collagen type-1, type-2 and type-3 in embryonic chick eye. Dev. Biol. 59, 75–85. 73. Linsenmayer, T. F., Smith, G. N., and Hay, E. D. (1977). Synthesis of 2 collagen types by embryonic chick corneal epithelium in vitro. P. Nas. US 74, 39–43. 74. Hay, E. D. (1981). Collagen and embryonic development. In “Cell biology of extracellular matrix” (E. D. Hay, ed.) Plenum Press, N. Y., pp. 379–409. 75. von der Mark, H., von der Mark, K., and Gay, S. (1976). Study of differential collagen synthesis during development of the chick embryo by immunofluorescence. I. Preparation of collagen type I and type II specific antibodies and their application to early stages of the chick embryo. Develop. Biol. 48, 237–249. 76. Niyibizi, C., and Eyre, D. R. (1989). Bone type V collagen: chain composition and location of a trypsin cleavage site. Connect. Tissue Res. 20, 247–250. 77. Hulmes, D. J. (1992). The collagen superfamily – diverse structures and assemblies. Essays. Biochem. 27, 49–67. 78. Ramachandran, G. N. (1988). Stereochemistry of collagen. Int. J. Pept. Protein Res. 31, 1–16. 79. Adachi, E., and Hayashi, T. (1985). In vitro formation of fine fibrils with a D-periodic banding pattern from type V collagen. Collagen Rel. Res. 5, 225–232. 80. Birk, D. E. (2001). Type V collagen: heterotypic type I/V collagen interactions in the regulation of fibril assembly. Micron. 32, 223–237. 81. Glimcher, M. J., Shapiro, F., Ellis, R. D., and Eyre, D. R. (1980). Changes in tissue morphology and collagen composition during the repair of cortical bone in the adult chicken. J. Bone Am. V. 62, 964–973. 82. Lees, S. (2003). Mineralization of type I collagen. Biophys. J. 85, 204–207. 83. Giraud-Guille, M. M., Besseau, L., and Martin, R. (2003). Liquid crystalline assemblies of collagen in bone and in vitro systems. J. Biomech. 36, 1571–1579. 84. Tromp, G., Kuivaniemi, H., Stacey, A., Shikata, H., Baldwin, C. T., Jaenisch, R., and Prockop, D. J. (1988). Structure of a full-length cDNA clone for the prepro alpha 1(I) chain of human type I procollagen. Biochem. J. 253, 633–640. 85. Kuivaniemi, H., Tromp, G., Chu, M. L., and Prockop, D. J. (1988). Structure of a full-length cDNA clone for the prepro alpha 2(I) chain of human type I procollagen. Comparison with the chicken gene confirms unusual patterns of gene conservation. Biochem. J. 252, 633–640. 86. Gerstenfeld, L. C., Chipman, S. D., Kelly, C. M., Hodgens, K. J., Lee, D. D., and Landis, W. J. (1988). Collagen expression, ultrastructural assembly, and mineralization in cultures of chicken embryo osteoblasts. J. Cell Biol. 106, 979–989. 87. Kratochwil, K., Ghaffari-Tabrizi, N., Holzinger, I., and Harbers, K. (1993). Restricted expression of Mov13 mutant alpha 1(I) collagen gene in osteoblasts and its consequences for bone development. Dev. Dyn. 198, 273–283. 88. Fleischmajer, R., MacDonald, E. D., Perlish, J. S., Burgeson, R. E., and Fisher, L. W. (1990). Dermal collagen fibrils are hybrids of type I and type III collagen molecules. J. Struct. Biol. 105, 162–169. 89. Birk, D. E., Fitch, J. M., Babiarz, J. P., and Linsenmayer, T. F. (1988). Collagen type I and type V are present in the same fibril in the avian corneal stroma. J. Cell Biol. 106, 999–1008. 90. von der Mark, K. (1981). Localization of collagen types in tissues. Int. Rev. Conn. Tissue Res. 9, 265–324.

KLAUS VON DER MARK 91. Byers, P. H. (2000). Osteogenesis imperfecta: perspectives and opportunities. Curr. Opin. Pediatr. 12, 603–609. 92. Brenner, R. E., Vetter, U., Nerlich, A., Worsdorfer, O., Teller, W. M., and Muller, P. K. (1989). Biochemical analysis of callus tissue in osteogenesis imperfecta type IV. Evidence for transient overmodification in collagen types I and III. J. Clin. Invest. 84, 915–921. 93. Rao, V. H., Steinmann, B., de Wet, W., and Hollister, D. W. (1989). Decreased thermal denaturation temperature of osteogenesis imperfecta mutant collagen is independent of post-translational overmodifications of lysine and hydroxylysine. J. Biol. Chem. 264, 1793–1798. 94. Montesano, R., Schaller, C., and Orci, L. (1991). Induction of epithelial tubular morphogenesis in vitro by fibroblast-derived soluble factors. Cell 66, 697–711. 95. Emsley, J., Knight, C. G., Farndale, R. W., Barnes, M. J., and Liddington, R. C. (2000). Structural basis of collagen recognition by integrin alpha2beta1. Cell 101, 47–56. 96. Brakebusch, C., and Fassler, R. (2003). The integrin-actin connection, an eternal love affair. Embo. J. 22, 2324–2333. 97. DeMali, K. A., Wennerberg, K., and Burridge, K. (2003). Integrin signaling to the actin cytoskeleton. Curr. Opin. Cell Biol. 15, 572–582. 98. Giancotti, F. G., and Ruoslahti, E. (1999). Integrin signaling. Science 285, 1028–1032. 99. Velling, T., Risteli, J., Wennerberg, K., Mosher, D. F., and Johansson, S. (2002). Polymerization of type I and III collagens is dependent on fibronectin and enhanced by integrins alpha 11beta 1 and alpha 2beta 1. J. Biol. Chem. 277, 37377–37381. 100. Still, J., Glat, P., Silverstein, P., Griswold, J., and Mozingo, D. (2003). The use of a collagen sponge/living cell composite material to treat donor sites in burn patients. Burns 29, 837–841. 101. Ueda, H., Hong, L., Yamamoto, M., Shigeno, K., Inoue, M., Toba, T., Yoshitani, M., Nakamura, T., Tabata, Y., and Shimizu, Y. (2002). Use of collagen sponge incorporating transforming growth factor-beta1 to promote bone repair in skull defects in rabbits. Biomaterials 23, 1003–1010. 102. Yamanouchi, K., Satomura, K., Gotoh, Y., Kitaoka, E., Tobiume, S., Kume, K., and Nagayama, M. (2001). Bone formation by transplanted human osteoblasts cultured within collagen sponge with dexamethasone in vitro. J. Bone Miner. Res. 16, 857–867. 103. Jaenisch, R., Harbers, K., Schnieke, A., Lohler, J., Chumakov, I., Jahner, D., Grotkopp, D., and Hoffmann, E. (1983). Germline integration of moloney murine leukemia virus at the Mov13 locus leads to recessive lethal mutation and early embryonic death. Cell 32, 209–216. 104. Hartung, S., Jaenisch, R., and Breindl, M. (1986). Retrovirus insertion inactivates mouse alpha 1(I) collagen gene by blocking initiation of transcription. Nature 320, 365–367. 105. Kratochwil, K., Dziadek, M., Lohler, J., Harbers, K., and Jaenisch, R. (1986). Normal epithelial branching morphogenesis in the absence of collagen I. Dev. Biol. 117, 596–606. 106. Schwarz, M., Harbers, K., and Kratochwil, K. (1990). Transcription of a mutant collagen-I gene is a cell type and stage-specific marker for odontoblast and osteoblast differentiation. Development 108, 717–726. 107. Kratochwil, K., von der Mark, K., Kollar, E. J., Jaenisch, R., Mooslehner, K., Schwarz, M., Haase, K., Gmacht, I., and Harbers, K. (1992). Retrovirus-induced insertional mutation in Mov13 mice affects collagen-I expression in a tissue-specific manner. Cell 57, 807–816. 108. Burgeson, R. E., Hebda, P. A., Morris, N. P., and Hollister, D. (1982). Human cartilage collagens. Comparison of cartilage collagens with human type V collagen. J. Biol. Chem. 257, 7852–7856. 109. Burgeson, R. E., and Hollister, D. (1979). Collagen heterogeneity in human cartilage: identification of several new collagen chains. Biochem. Biophys. Res. Commun. 87, 1124–1131. 110. Zhidkova, N. I., Brewton, R. G., and Mayne, R. (1993). Molecular cloning of PARP (proline/arginine-rich protein) from human cartilage and subsequent demonstration that PARP is a fragment of the

CHAPTER 1 Structure, Biosynthesis and Gene Regulation of Collagens in Cartilage and Bone

111.

112.

113.

114.

115.

116.

117.

118.

119.

120.

121.

122.

123.

124.

125.

126. 127.

128.

129.

NH2-terminal domain of the collagen alpha 2(XI) chain. FEBS Lett. 326, 25–28. Fessler, L., Kumamoto, C., Meis, M., and Fessler, J. (1981). Assembly and processing of procollagen V (AB) in chick blood vessels and other tissues. J. Biol. Chem. 256, 9640–9645. Fessler, L. I., Brosh, S., Chapin, S., and Fessler, J. H. (1986). Tyrosine sulfation in precursors of collagen V. J. Biol. Chem. 261, 5034–5040. Wu, J. J., Lark, M. W., Chun, L. E., and Eyre, D. R. (1991). Sites of stromelysin cleavage in collagen types II, IX, X, and XI of cartilage. J. Biol. Chem. 266, 5625–5628. Haralson, M. A., Mitchell, W. M., Rhodes, R. K., Kresina, T. F., Gay, R., and Miller, E. J. (1980). Chinese hamster lung cells synthesize and confine to the cellular domain a collagen composed solely of B chains. Proc. Natl. Acad. Sci. USA 77, 5206–5210. Eyre, D., and Wu, J.-J. (1987). Type XI or 1alpha, 2alpha, 3alpha collagen. In: “Stucture and Function of Collagen Types” (R. Mayne and R.E. Burgeson, eds.), pp. 261–281. Niyibizi, C., and Eyre, D. R. (1989). Identification of the cartilage alpha 1(XI) chain in type V collagen from bovine bone. FEBS Lett. 242, 314–318. Mayne, R., Brewton, R. G., Mayne, P. M., and Baker, J. R. (1993). Isolation and characterization of the chains of type V/type XI collagen present in bovine vitreous. J. Biol. Chem. 268, 9381–9386. von der Mark, K., and Oecalan, M. (1982). Immunofluorescent localization of type V collagen in the chick embryo with monoclonal antibodies. Collagen Rel. Res. 2, 541–555. Linsenmayer, T. F., Fitch, J. M., Schmid, T. M., Zak, N. B., Gibney, E., Sanderson, R. D., and Mayne, R. (1983). Monoclonal antibodies against chicken type V collagen: Production, specificity, and use for immunocytochemical localization in embryonic cornea and other organs. J. Cell Biol. 96, 124–132. Mayne, R. (1989). Cartilage collagens – what is their function, and are they involved in articular disease. Arthritis Rheumatism 32, 241–246. von der Mark, K., and von der Mark, H. (1977). Role of 3 genetically distinct collagen types in endochondral ossification and calcification of cartilage. J. Bone Joint Res. 59, 458–464. Eyre, D. R., Brickley-Parsons, D. M., and Glimcher, M. J. (1978). Predominance of type I collagen at the surface of avian articular cartilage. FEBS Lett. 85, 259–263. Goldwasser, M., Astley, T., van der Rest, M., and Glorieux, F. H. (1982). Analysis of the type of collagen present in osteoarthritic human cartilage. Clin. Orthop. Rev. Res. 167, 296–302. Ronziere, M. C., Ricard-Blum, S., Tiollier, J., Hartmann, D. J., Garrone, R., and Herbage, D. (1990). Comparative analysis of collagens solubilized from human foetal, and normal and osteoarthritic adult articular cartilage, with emphasis on type VI collagen. Biochim. Biophys. Acta 1038, 222–230. Aigner, T., Bertling, W., Stöb, H., Weseloh, G., and von der Mark, K. (1993). Independent expression of fibril-forming collagens I, II, and III in chondrocytes in human osteoarthritic cartilage. J. Clin. Invest. 91, 829–837. Wotton, S. F., and Duance, V. C. (1994). Type III collagen in normal human articular cartilage. Histochem. J. 26, 412–416. Young, R. D., Lawrence, P. A., Duance, V. C., Aigner, T., and Monaghan, P. (2000). Immunolocalization of collagen types II and III in single fibrils of human articular cartilage. J. Histochem. Cytochem. 48, 423–432. Eerola, I., Salminen, H., Lammi, P., Lammi, M., von der Mark, K., Vuorio, E., and Saamanen, A. M. (1998). Type X collagen, a natural component of mouse articular cartilage: association with growth, aging, and osteoarthritis. Arthritis Rheum. 41, 1287–1295. Lammi, P. E., Lammi, M. J., Hyttinen, M. M., Panula, H., Kiviranta, I., and Helminen, H. J. (2002). Site-specific immunostaining for type X

130.

131.

132.

133.

134.

135.

136.

137.

138.

139.

140.

141.

142.

143.

144.

145.

146.

29

collagen in noncalcified articular cartilage of canine stifle knee joint. Bone 31, 690–696. Rucklidge, G. J., Milne, G., and Robins, S. P. (1996). Collagen type X: a component of the surface of normal human, pig, and rat articular cartilage. Biochem. Biophys. Res. Commun. 224, 297–302. von der Mark, K., Kirsch, T., Nerlich, A., Kuss, A., Weseloh, G., Glueckert, K., and Stoess, H. (1992). Type X collagen synthesis in human osteoarthritic cartilage: Indication of chondrocyte hypertrophy. Arthritis Rheum. 35, 806–811. Aigner, T., Reichenberger, E., Bertling, W., Kirsch, T., Stöb, H., and von der Mark, K. (1993). Type X collagen expression in osteoarthritic and rheumatoid articular cartilage. Virchows Arch. 63, 205–211. Kwan, K. M., Pang, M. K., Zhou, S., Cowan, S. K., Kong, R. Y., Pfordte, T., Olsen, B. R., Sillence, D. O., Tam, P. P., and Cheah, K. S. (1997). Abnormal compartmentalization of cartilage matrix components in mice lacking collagen X: implications for function. J. Cell Biol. 136, 459–471. Jacenko, O., Chan, D., Franklin, A., Ito, S., Underhill, C. B., Bateman, J. F., and Campbell, M. R. (2001). A dominant interference collagen X mutation disrupts hypertrophic chondrocyte pericellular matrix and glycosaminoglycan and proteoglycan distribution in transgenic mice. Am. J. Pathol. 159, 2257–2269. Jacenko, O., Roberts, D. W., Campbell, M. R., McManus, P. M., Gress, C. J., and Tao, Z. (2002). Linking hematopoiesis to endochondral skeletogenesis through analysis of mice transgenic for collagen X. Am. J. Pathol. 160, 2019–2034. Topping, R. E., Bolander, M. E., and Balian, G. (1994). Type X collagen in fracture callus and the effects of experimental diabetes. Clin. Orthop. 308, 220–228. Trelstad, R. L., Kang, A. H., Igarashi, S., and Gross, J. (1970). Isolation of two distinct collagens from chick cartilage. Biochemistry 9, 4993–4998. Miller, E. J. (1971). Isolation and characterization of a collagen from chick cartilage containing three identical alpha chains. Biochemistry 10, 1652–1659. Ryan, M. C., and Sandell, L. J. (1990). Differential expression of a cysteine-rich domain in the amino- terminal propeptide of type II procollagen by alternative splicing of messenger-RNA. J. Biol. Chem. 265, 10334–10339. Sandell, L. J., Nalin, A. M., and Reife, R. A. (1994). Alternative splice form of type II procollagen mRNA (IIA) is predominant in skeletal precursors and non-cartilaginous tissues during early mouse development. Dev. Dyn. 199, 129–140. Madsen, K., Moskalewski, S., von der Mark, K., and Friberg, U. (1983). Synthesis of proteoglycans, collagen, and elastin by cultures of rabbit auricular chondrocytes – relation to age of the donor. Dev. Biol. 96, 63–73. Eyre, D. R., and Muir, H. (1976). Types I and II collagens in intervertebral disc. Interchanging radial distributions in annulus fibrosus. Biochem. J. 157, 267–270. Yang, C. L., Rui, H., Mosler, S., Notbohm, H., Sawaryn, A., and Muller, P. K. (1993). Collagen II from articular cartilage and annulus fibrosus. Structural and functional implication of tissue specific posttranslational modifications of collagen molecules. Eur. J. Biochem. 213, 1297–1302. Newsome, D. A., Linsenmayer, T. F., and Trelstad, R. L. (1976). Vitreous body collagen. Evidence for a dual origin from the neural retina and hyalocytes. J. Cell Biol. 71, 59–67. Trelstad, R. L., and Kang, A. H. (1974). Collagen heterogeneity in the avian eye: lens, vitreous body, cornea and sclera. Exp. Eye Res. 18, 395–406. Thorogood, P., Bee, J., and von der Mark, K. (1986). Transient expression of collagen type II at epitheliomesenchymal interfaces during morphogenesis of the cartilaginous neurocranium. Develop. Biol. 116, 497–509.

30 147. Wood, A., Ashhurst, D. E., Corbett, A., and Thorogood, P. (1991). The transient expression of type II collagen at tissue interfaces during mammalian craniofacial development. Development 111, 955–968. 148. Mundlos, S., and Olsen, B. R. (1997). Heritable diseases of the skeleton. Part II: Molecular insights into skeletal development-matrix components and their homeostasis. Faseb. J. 11, 227–233. 149. Spranger, J., Winterpacht, A., and Zabel, B. (1994). The type II collagenopathies: a spectrum of chondrodysplasias. Eur. J. Pediatr. 153, 56–65. 150. Byers, P. H. (1994). Molecular genetics of chondrodysplasias, including clues to development, structure, and function. Curr. Opin. Rheumatol. 6, 345–350. 151. Chan, D., Cole, W. G., Chow, C. W., Mundlos, S., and Bateman, J. F. (1995). COL2A1 mutation in achondrogenesis type II results in the replacement of type II collagen type I and III collagens in cartilage. J. Biol. Chem. 270, 1747–1753. 152. Mundlos, S., Chan, D., McGill, J., and Bateman, J. F. (1996). An alpha 1(II) Gly913 to Cys substitution prevents the matrix incorporation of type II collagen which is replaced with type I and III collagens in cartilage from a patient with hypochondrogenesis. Am. J. Med. Genet. 63, 129–136. 153. Aszodi, A., Hunziker, E. B., Olsen, B. R., and Fassler, R. (2001). The role of collagen II and cartilage fibril-associated molecules in skeletal development. Osteoarthritis Cartilage 9, Suppl A, S150–159. 154. Conrad, G. W., Kelly, P. T., von der Mark, K., and Edelhauser, H. F. (1981). A comparative study of elasmobranch corneal and scleral collagens. Expt. Eye Res. 32, 659–672. 155. Holmdahl, R., Malmstrom, V., and Vuorio, E. (1993). Autoimmune recognition of cartilage collagens. Ann. Med. 25, 251–264. 156. Stuart, J. M., Watson, W. C., and Kang, A. H. (1988). Collagen autoimmunity and arthritis. Faseb. J. 2, 2950–2956. 157. Watson, W. C., Cremer, M. A., Wooley, P. H., and Townes, A. S. (1986). Assessment of the potential pathogenicity of type II collagen autoantibodies in patients with rheumatoid arthritis. Evidence of restricted IgG3 subclass expression and activation of complement C5 to C5a. Arthritis Rheum. 29, 1316–1321. 158. Holmdahl, R., Andersson, M., Goldschmidt, T. J., Gustafsson, K., Jansson, L., and Mo, J. A. (1990). Type II collagen autoimmunity in animals and provocations leading to arthritis. Immunol. Rev. 118, 193–232. 159. Andersson, M., Cremer, M. A., Terato, K., Burkhardt, H., and Holmdahl, R. (1991). Analysis of type II collagen reactive T cells in the mouse. II. Different localization of immunodominant T cell epitopes on heterologous and autologous type II collagen. Scand. J. Immunol. 33, 505–510. 160. Schulte, S., Unger, C., Mo, J. A., Wendler, O., Bauer, E., Frischholz, S., von der Mark, K., Kalden, J. R., Holmdahl, R., and Burkhardt, H. (1998). Arthritis-related B cell epitopes in collagen II are conformationdependent and sterically privileged in accessible sites of cartilage collagen fibrils. J. Biol. Chem. 273, 1551–1561. 161. Burkhardt, H., Koller, T., Engstrom, A., Nandakumar, K. S., Turnay, J., Kraetsch, H. G., Kalden, J. R., and Holmdahl, R. (2002). Epitopespecific recognition of type II collagen by rheumatoid arthritis antibodies is shared with recognition by antibodies that are arthritogenic in collagen-induced arthritis in the mouse. Arthritis Rheum. 46, 2339–2348. 162. Backlund, J., Carlsen, S., Hoger, T., Holm, B., Fugger, L., Kihlberg, J., Burkhardt, H., and Holmdahl, R. (2002). Predominant selection of T cells specific for the glycosylated collagen type II epitope (263–270) in humanized transgenic mice and in rheumatoid arthritis. Proc. Natl. Acad. Sci. USA 99, 9960–9965. 163. Morris, N. P., and Baechinger, H. P. (1987). Type XI collagen is a heterotrimer with the composition (1alpha, 2alpha, 3alpha) retaining non-triple-helical domains. J. Biol. Chem. 262, 11345–11350.

KLAUS VON DER MARK 164. Wu, J. J., and Eyre, D. R. (1995). Structural analysis of cross-linking domains in cartilage type XI collagen. Insights on polymeric assembly. J. Biol. Chem. 270, 18865–18870. 165. Oxford, J. T., DeScala, J., Morris, N., Gregory, K., Medeck, R., Irwin, K., Oxford, R., Brown, R., Mercer, L., and Cusack, S. (2004). Interaction between amino propeptides of type XI procollagen alpha1 chains. J. Biol. Chem. 279, 10939–10945. 166. Davies, G. B., Oxford, J. T., Hausafus, L. C., Smoody, B. F., and Morris, N. P. (1998). Temporal and spatial expression of alternative splice-forms of the alpha1(XI) collagen gene in fetal rat cartilage. Dev. Dyn. 213, 12–26. 167. Lui, V. C. H., Ng, L. J., Sat, E. W. Y., Nicholls, J., and Cheah, K. S. E. (1996). Extensive alternative splicing within the amino-propeptide coding domain of alpha2(XI) procollagen mRNAs. Expression of transcripts encoding truncated pro-alpha chains. J. Biol. Chem. 271, 16945–16951. 168. Tsumaki, N., and Kimura, T. (1995). Differential expression of an acidic domain in the amino- terminal propeptide of mouse pro-alpha 2(XI) collagen by complex alternative splicing. J. Biol. Chem. 270, 2372–2378. 169. Neame, P. J., Young, C. N., and Treep, J. T. (1990). Isolation and primary structure of PARP, a 24-kDa proline- and arginine-rich protein from bovine cartilage closely related to the NH2-terminal domain in collagen alpha 1 (XI). J. Biol. Chem. 265, 20401–20408. 170. Mendler, M., Eich-Bender, S. G., Vaughan, L., Winterhalter, K. H., and Bruckner, P. (1989). Cartilage contains mixed fibrils of collagen types II, IX and XI. J. Cell Biol. 108, 191–197. 171. Blaschke, U. K., Eikenberry, E. F., Hulmes, D. J., Galla, H. J., and Bruckner, P. (2000). Collagen XI nucleates self-assembly and limits lateral growth of cartilage fibrils. J. Biol. Chem. 275, 10370–10378. 172. Gregory, K. E., Oxford, J. T., Chen, Y., Gambee, J. E., Gygi, S. P., Aebersold, R., Neame, P. J., Mechling, D. E., Bachinger, H. P., and Morris, N. P. (2000). Structural organization of distinct domains within the non-collagenous N-terminal region of collagen type XI. J. Biol. Chem. 275, 11498–11506. 173. Li, Y., Lacerda, D. A., Warman, M. L., Beier, D. R., Yoshioka, H., Ninomiya, Y., Oxford, J. T., Morris, N. P., Andrikopoulos, K., and Ramirez, F. (1995). A fibrillar collagen gene, Col11a1, is essential for skeletal morphogenesis. Cell 80, 423–430. 174. Vikkula, M., Mariman, E. C., Lui, V. C., Zhidkova, N. I., Tiller, G. E., Goldring, M. B., van Beersum, S. E., de Waal Malefijt, M. C., van den Hoogen, F. H., and Ropers, H. H. (1995). Autosomal dominant and recessive osteochondrodysplasias associated with the COL11A2 locus. Cell 80, 431–437. 175. Shimokomaki, M., Duance, V. C., and Bailey, A. J. (1980). Identification of a new disulphide bonded collagen from cartilage. FEBS Lett. 121, 51–54. 176. Reese, C. A., and Mayne, R. (1981). Minor collagens of chicken hyaline cartilage. Biochem. 20, 5443–5448. 177. Ninomiya, Y., and Olsen, B. R. (1984). Synthesis and characterization of cDNA encoding a cartilage-specific short collagen. Proc. Natl. Acad. Sci. USA 81, 3014–3018. 178. von der Mark, K., van Menxel, M., and Wiedemann, H. (1984). Isolation and characterization of a precursor form of M-collagen from embryonic chicken cartilage. Eur. J. Biochem. 138, 629–633. 179. Mayne, R., and von der Mark, K. (1983). Collagens of cartilage. Biochem. Cartilage I, 181–214. 180. van der Rest, M., Mayne, R., Ninomiya, Y., Seidah, N. G., Chretien, M., and Olsen, B. R. (1985). The structure of type IX collagen. J. Biol. Chem. 260, 220–225. 181. Vaughan, L., Mendler, M., Huber, S., Bruckner, P., Winterhalter, K. H., Irwin, M. I., and Mayne, R. (1988). D-periodic distribution of collagen type IX along cartilage fibrils. J. Cell Biol. 106, 998–997.

CHAPTER 1 Structure, Biosynthesis and Gene Regulation of Collagens in Cartilage and Bone 182. van der Rest, M., Mayne, R. (1988). Type IX collagen proteoglycan from cartilage is covalently cross-linked to type II collagen. J. Biol. Chem. 263, 1615–1618. 183. Wu, J. J., Woods, P. E., and Eyre, D. R. (1992). Identification of crosslinking sites in bovine cartilage type IX collagen reveals an antiparallel type II-type IX molecular relationship and type IX to type IX bonding. J. Biol. Chem. 267, 23007–23014. 184. Vaughan, L., Winterhalter, K. H., and Bruckner, P. (1985). Proteoglycan Lt from chicken embryo sternum identified as type IX collagen. J. Biol. Chem. 260, 4758–4763. 185. Noro, A., Kimata, K., Oike, Y., Shinomura, T., Maeda, N., Yano, S., Takahashi, N., and Suzuki, S. (1983). Isolation and characterization of a third proteoglycan (PG-Lt) from chick embryo cartilage which contains disulfide-bonded collagenous polypeptide. J. Biol. Chem. 258, 9323–9331. 186. Huber, S., Winterhalter, K. H., and Vaughan, L. (1988). Isolation and sequence-analysis of the glycosaminoglycan attachment site of type-IX collagen. J. Biol. Chem. 263, 752–756. 187. Brewton, R. G., Wright, D. W., and Mayne, R. (1991). Structural and functional comparison of type IX collagen- proteoglycan from chicken cartilage and vitreous humor. J. Biol. Chem. 266, 4752–4757. 188. Nishimura, I., Muragaki,Y., and Olsen, B. R. (1989). Tissue-specific forms of type IX collagen-proteoglycan arise from the use of two widely separated promoters. J. Biol. Chem. 264, 20033–20041. 189. Muragaki, Y., Nishimura, I., Henney, A., Ninomiya, Y., and Olsen, B. R. (1990). The alpha 1(IX) collagen gene gives rise to two different transcripts in both mouse embryonic and human fetal RNA. Proc. Natl. Acad. Sci. USA 87, 2400–2404. 190. Fassler, R., Schnegelsberg, P. N., Dausman, J., Shinya, T., Muragaki, Y., McCarthy, M. T., Olsen, B. R., and Jaenisch, R. (1994). Mice lacking α1(IX) collagen develop non-inflammatory degenerative joint disease. Proc. Natl. Acad. Sci. USA 91, 5070–5074. 191. Watt, S. L., Lunstrum, G. P., McDonough, A. M., Keene, D. R., Burgeson, R. E., and Morris, N. P. (1992). Characterization of collagen types XII and XIV from fetal bovine cartilage. J. Biol. Chem. 267, 20093–20099. 192. Gordon, M. K., Gerecke, D. R., and Olsen, B. R. (1987). Type XII collagen: Distinct extracellular matrix component discovered by cRNA cloning. Proc. Natl. Acad. Sci. USA 84, 6040–6044. 193. Dublet, B., and van der Rest, M. (1991). Type XIV collagen, a new homotrimeric molecule extracted from bovine skin and tendon, with a triple helical disulfide-bonded domain homologous to type IX and type XII collagens. J. Biol. Chem. 266, 6853–6858. 194. Lunstrum, G. P., Morris, N. P., McDonough, A. M., Keene, D. R., and Burgeson, R. E. (1991). Identification and partial characterization of two type XII-like collagen molecules. J. Cell Biol. 113, 963–969. 195. Keene, D. R., Lunstrum, G. P., Morris, N. P., Stoddard, D. W., and Burgeson, R. E. (1991). Two type XII-like collagens localize to the surface of banded collagen fibrils. J. Cell Biol. 113, 971–978. 196. Lunstrum, G. P., McDonough, A. M., Marinkovich, M. P., Keene, D. R., Morris, N. P., and Burgeson, R. E. (1992). Identification and partial purification of a large, variant form of type XII collagen. J. Biol. Chem. 267, 20087–20092. 197. Gerecke, D. R., Olson, P. F., Koch, M., Knoll, J. H., Taylor, R., Hudson, D. L., Champliaud, M. F., Olsen, B. R., and Burgeson, R. E. (1997). Complete primary structure of two splice variants of collagen XII, and assignment of alpha 1(XII) collagen (COL12A1), alpha 1(IX) collagen (COL9A1), and alpha 1(XIX) collagen (COL19A1) to human chromosome 6q12- q13. Genomics 41, 236–242. 198. Aubert-Foucher, E., Font, B., Eichenberger, D., Goldschmidt, D., Lethias, C., and van der Rest, M. (1992). Purification and characterization of native type XIV collagen. J. Biol. Chem. 267, 15759–15764. 199. Schuppan, D., Cantaluppi, M. C., Becker, J., Veit, A., Bunte, T., Troyer, D., Schuppan, F., Schmid, M., Ackermann, R., and Hahn, E. G.

200.

201.

202.

203.

204.

205.

206.

207.

208.

209.

210.

211.

212.

213.

214.

215.

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(1990). Undulin, an extracellular matrix glycoprotein associated with collagen fibrils. J. Biol. Chem. 265, 8823–8832. Walchli, C., Koch, M., Chiquet, M., Odermatt, B. F., and Trueb, B. (1994). Tissue-specific expression of the fibril-associated collagens XII and XIV. J. Cell Sci. 107, 669–681. Berthod, F., Germain, L., Guignard, R., Lethias, C., Garrone, R., Damour, O., van der Rest, M., and Auger, F. A. (1997). Differential expression of collagens XII and XIV in human skin and in reconstructed skin. J. Invest. Dermatol. 108, 737–742. Font, B., Aubert-Foucher, E., Goldschmidt, D., Eichenberger, D., and van der Rest, M. (1993). Binding of collagen XIV with the dermatan sulfate side chain of decorin. J. Biol. Chem. 268, 25015–25018. Ehnis, T., Dieterich, W., Bauer, M., Kresse, H., and Schuppan, D. (1997). Localization of a binding site for the proteoglycan decorin on collagen XIV (undulin). J. Biol. Chem. 272, 20414–20419. Kassner, A., Hansen, U., Miosge, N., Reinhardt, D. P., Aigner, T., Bruckner-Tuderman, L., Bruckner, P., and Grassel, S. (2003). Discrete integration of collagen XVI into tissue-specific collagen fibrils or beaded microfibrils. Matrix Biol. 22, 131–143. Kassner, A., Tiedemann, K., Notbohm, H., Ludwig, T., Morgelin, M., Reinhardt, D. P., Chu, M. L., Bruckner, P., and Grassel, S. (2004). Molecular structure and interaction of recombinant human type XVI collagen. J. Mol. Biol. 339, 835–853. Pan, T. C., Zhang, R. Z., Mattei, M. G., Timpl, R., and Chu, M. L. (1992). Cloning and chromosomal location of human alpha 1(XVI) collagen. Proc. Natl. Acad. Sci. USA 89, 6565–6569. Myers, J. C., Li, D., Amenta, P. S., Clark, C. C., Nagaswami, C., and Weisel, J. W. (2003). Type XIX collagen purified from human umbilical cord is characterized by multiple sharp kinks delineating collagenous subdomains and by intermolecular aggregates via globular, disulfidelinked, and heparin-binding amino termini. J. Biol. Chem. 278, 32047–32057. Sumiyoshi, H., Inoguchi, K., Khaleduzzaman, M., Ninomiya, Y., and Yoshioka, H. (1997). Ubiquitous expression of the alpha1(XIX) collagen gene (Col19a1) during mouse embryogenesis becomes restricted to a few tissues in the adult organism. J. Biol. Chem. 272, 17104–17111. Tuckwell, D. (2002). Identification and analysis of collagen alpha 1(XXI), a novel member of the FACIT collagen family. Matrix Biol. 21, 63–66. Timpl, R., and Engel, J. (1987). Type VI collagen. In: “Structure and Function of Collagen Types.” (R. Mayne and R. E. Burgeson, eds.), pp. 105–143, Academic Press. Chu, M. L., Pan, T. C., Conway, D., Kuo, H. J., Glanville, R. W., Timpl, R., Mann, K., and Deutzmann, R. (1989). Sequence analysis of alpha 1(VI) and alpha 2(VI) chains of human type VI collagen reveals internal triplication of globular domains similar to the A domains of von Willebrand factor and two alpha 2(VI) chain variants that differ in the carboxy terminus. EMBO J. 8, 1939–1946. Bonaldo, P., Russo, V., Bucciotti, F., Bressan, G. M., and Colombatti, A. (1989). Alpha 1 chain of chick type VI collagen. The complete cDNA sequence reveals a hybrid molecule made of one short collagen and three von Willebrand factor type A-like domains. J. Biol. Chem. 264, 5575–5580. Koller, E., Winterhalter, K. H., and Trueb, B. (1989). The globular domains of type VI collagen are related to the collagen-binding domains of cartilage matrix protein and von Willebrand factor. EMBO J. 8, 1073–1077. Ball, S., Bella, J., Kielty, C., and Shuttleworth, A. (2003). Structural basis of type VI collagen dimer formation. J. Biol. Chem. 278, 15326–15332. Doliana, R., Mucignat, M. T., Segat, D., Zanussi, S., Fabbro, C., Lakshmi, T. R., and Colombatti, A. (1998). Alternative splicing of VWFA modules generates variants of type VI collagen alpha 3 chain with a distinctive expression pattern in embryonic chicken tissues and potentially different adhesive function. Matrix Biol. 16, 427–442.

32 216. Saitta, B., Stokes, D. G., Vissing, H., Timpl, R., and Chu, M. L. (1990). Alternative splicing of the human alpha 2(VI) collagen gene generates multiple mRNA transcripts which predict three protein variants with distinct carboxyl termini. J. Biol. Chem. 265, 6473–6480. 217. Mayer, U., Poschl, E., Nischt, R., Specks, U., Pan, T. C., Chu, M. L., and Timpl, R. (1994). Recombinant expression and properties of the Kunitz-type protease-inhibitor module from human type VI collagen alpha 3(VI) chain. Eur. J. Biochem. 225, 573–580. 218. Heckmann, M., Aumailley, M., Hatamochi, A., Chu, M.-L., Timpl, R., and Krieg, T. (1989). Down-regulation of alpha3(VI) chain expression by gamma- interferon dereases synthesis and deposition of collagen type VI. Eur. J. Biochem. 182, 719–726. 219. Doane, K. J., Howell, S. J., and Birk, D. E. (1998). Identification and functional characterization of two type VI collagen receptors, alpha 3 beta 1 integrin and NG2, during avian corneal stromal development. Invest. Ophthalmol. Vis. Sci. 39, 263–275. 220. Kuo, H. J., Maslen, C. L., Keene, D. R., and Glanville, R. W. (1997). Type VI collagen anchors endothelial basement membranes by interacting with type IV collagen. J. Biol. Chem. 272, 26522–26529. 221. Furthmayr, H., Wiedemann, H., Timpl, R., Odermatt, E., and Engel, J. (1983). Electron-microscopical approach to a structural model of intima collagen. Biochem. J. 211, 303–311. 222. Bruns, R. R., Press, W., Engvall, E. T., and Gross, J. (1986). Type VI collagen in extracellular, 100-nm periodic filaments and fibrils: identification by immunoelectron microscopy. J. Cell Biol. 103, 393–404. 223. von der Mark, H., Aumailley, M., Wick, G., Fleischmajer, R., and Timpl, R. (1984). Immunochemistry, genuine size and tissue localization of collagen VI. Eur. J. Biochem. 142, 493–502. 224. Wiberg, C., Klatt, A. R., Wagener, R., Paulsson, M., Bateman, J. F., Heinegard, D., and Morgelin, M. (2003). Complexes of matrilin-1 and biglycan or decorin connect collagen VI microfibrils to both collagen II and aggrecan. J. Biol. Chem. 278, 37698–37704. 225. Wiberg, C., Heinegard, D., Wenglen, C., Timpl, R., and Morgelin, M. (2002). Biglycan organizes collagen VI into hexagonal-like networks resembling tissue structures. J. Biol. Chem. 277, 49120–49126. 226. Quarto, R., Dozin, B., Bonaldo, P., Cancedda, R., and Colombatti, A. (1993). Type VI collagen expression is upregulated in the early events of chondrocyte differentiation. Development 117, 245–251. 227. Swoboda, B., Pullig, O., Kirsch, T., Kladny, B., Steinhauser, B., and Weseloh, G. (1998). Increased content of type-VI collagen epitopes in human osteoarthritic cartilage: quantitation by inhibition ELISA. J. Orthop. Res. 16, 96–99. 228. Kielty, C. M., Cummings, C., Whittaker, S. P., Shuttleworth, C. A., and Grant, M. E. (1991). Isolation and ultrastructural analysis of microfibrillar structures from foetal bovine elastic tissues. Relative abundance and supramolecular architecture of type VI collagen assemblies and fibrillin. J. Cell. Sci. 99, 797–807. 229. Aumailley, M., Mann, K., von der Mark, H., and Timpl, R. (1989). Cell attachment properties of collagen type VI and Arg-Gly-Asp dependent binding to its alpha2(VI) and alpha3(VI) chains. Exp. Cell Res. 181, 463–474. 230. Gibson, G., Schor, S. L., and Grant, M. E. (1982). Effects of matrix molecules on chondrocyte gene expression. Synthesis of a low molecular weight collagen species by cells cultured within collagen gels. J. Cell Biol. 93, 767–774. 231. Capasso, O., Tajana, G., and Cancedda, R. (1984). Location of 64K collagen producing chondrocytes in developing chick embryo tibiae. Mol. Cell. Biol. 4, 1163–1168. 232. Schmid, T. M., and Linsenmayer, T. F. (1987). Type X collagen. In: “Structure and Function of Collagen types” (R. Mayne and R. E. Burgeson, eds.), Academic Press, pp. 223–259. 233. Aigner, T., Frischholz, S., Dertinger, S., Beier, F., Girkontaite, I., and von der Mark, K. (1997). Type X collagen expression and hypertrophic differentiation in chondrogenic neoplasias. Histochem. Cell Biol. 107, 435–440.

KLAUS VON DER MARK 234. Girkontaite, J., Frischholz, S., Lammi, P., Wagner, K., Swoboda, B., Aigner, T., and von der Mark, K. (1996). Immunolocalization of type X collagen in normal fetal and adult osteoarthritic cartilage with monoclonal antibodies. Matrix Biol. 15, 231–238. 235. Walker, G. D., Fischer, M., Gannon, J., Thompson, R. C., and Oegama, T. R. (1995). Expression of type X collagen in osteoarthritis. J. Orthopaedic Res. 13, 4–12. 236. Yamaguchi, N., Mayne, R., and Ninomiya, Y. (1991). The alpha 1 (VIII) collagen gene is homologous to the alpha 1 (X) collagen gene and contains a large exon encoding the entire triple helical and carboxyl-terminal non-triple helical domains of the alpha 1 (VIII) polypeptide. J. Biol. Chem. 266, 4508–4513. 237. Sawada, H., Konomi, H., and Hirosawa, K. (1990). Characterization of the collagen in the hexagonal lattice of Descemet’s membrane: its relation to type VIII collagen. J. Cell Biol. 110, 219–227. 238. Kwan, A. P. L., Cummings, C. E., Chapman, J. A., and Grant, M. E. (1991). Macromolecular organization of chicken type X collagen in vitro. J. Cell Biol. 114, 597–605. 239. Brass, A., Kadler, K. E., Thomas, J. T., Grant, M. E., and Boot-Handford, R. P. (1992). The fibrillar collagens, collagen VIII, collagen X and the C1q complement proteins share a similar domain in their C-terminal non-collagenous regions. FEBS Lett. 303, 126–128. 240. Marks, D. S., Gregory, C. A., Wallis, G. A., Brass, A., Kadler, K. E., and Boot-Handford, R. P. (1999). Metaphyseal chondrodysplasia type Schmid mutations are predicted to occur in two distinct threedimensional clusters within type X collagen NC1 domains that retain the ability to trimerize. J. Biol. Chem. 274, 3632–3641. 241. Bonaventure, J., Chaminade, F., and Maroteaux, P. (1995). Mutations in three subdomains of the carboxy-terminal region of collagen type X account for most of the Schmid metaphyseal dysplasias. Hum. Genet. 96, 58–64. 242. Chan, D., and Jacenko, O. (1998). Phenotypic and biochemical consequences of collagen X mutations in mice and humans. Matrix Biol. 17, 169–184. 243. Chan, D., Weng, Y. M., Golub, S., and Bateman, J. F. (1996). Type X collagen NC1 mutations produced by site-directed mutagenesis prevent in vitro assembly. Ann. Acad. Sci. N.Y. 785, 231–233. 244. Wallis, G. A., Rash, B., Sykes, B., Bonaventure, J., Maroteaux, P., Zabel, B., Wynne-Davies, R., Grant, M. E., and Boot-Handford, R. P. (1996). Mutations within the gene encoding the alpha 1 (X) chain of type X collagen (COL10A1) cause metaphyseal chondrodysplasia type Schmid but not several other forms of metaphyseal chondrodysplasia. J. Med. Genet. 33, 450–457. 245. Warman, M. L., Abbott, M., Apte, S. S., Hefferon, T., McIntosh, I., Cohn, D. H., Hecht, J. T., Olsen, B. R., and Francomano, C. A. (1993). A type X collagen mutation causes Schmid metaphyseal chondrodysplasia. Nat. Gen. 5, 79–82. 246. Dublet, B., Vernet, T., and van der Rest, M. (1999). Schmid’s metaphyseal chondrodysplasia mutations interfere with folding of the C-terminal domain of human collagen X expressed in Escherichia coli. J. Biol. Chem. 274, 18909–18915. 247. Jacenko, O., LuValle, P. A., and Olsen, B. R. (1993). Spondylometaphyseal dysplasia in mice carrying a dominant negative mutation in a matrix protein specific for cartilage to bone transition. Nature 365, 56–61. 248. Rosati, R., Horan, G. S., Pinero, G. J., Garofalo, S., Keene, D. R., Horton, W. A., Vuorio, E., deCrombrugghe, B., and Behringer, R. R. (1994). Normal long bone growth and development in type X collagennull mice. Nat. Gen. 8, 129–135. 249. Gress, C. J., and Jacenko, O. (2000). Growth plate compressions and altered hematopoiesis in collagen X null mice. J. Cell. Biol. 149, 983–993. 250. Hagg, P. M., Horelli-Kuitunen, N., Eklund, L., Palotie, A., and Pihlajaniemi, T. (1997). Cloning of mouse type XV collagen sequences and mapping of the corresponding gene to 4B1–3. Comparison of

CHAPTER 1 Structure, Biosynthesis and Gene Regulation of Collagens in Cartilage and Bone

251.

252.

253.

254.

255.

256.

257.

258.

259.

260.

261.

262.

263.

264.

265.

mouse and human alpha 1 (XV) collagen sequences indicates divergence in the number of small collagenous domains. Genomics 45, 31–41. Rehn, M., Hintikka, E., and Pihlajaniemi, T. (1996). Characterization of the mouse gene for the alpha 1 chain of type XVIII collagen (Col18a1) reveals that the three variant N- terminal polypeptide forms are transcribed from two widely separated promoters. Genomics 32, 436–446. Oh, S. P., Kamagata, Y., Muragaki, Y., Timmons, S., Ooshima, A., and Olsen, B. R. (1994). Isolation and sequencing of cDNAs for proteins with multiple domains of Gly-Xaa-Yaa repeats identify a distinct family of collagenous proteins. Proc. Natl. Acad. Sci. U.S.A. 91, 4229–4233. Van den Bergh, F., and Giudice, G. J. (2003). BP180 (type XVII collagen) and its role in cutaneous biology and disease. Adv. Dermatol. 19, 37–71. Ricard-Blum, S., Dublet, B. and van der Rest, M. (2000). Unconventional Collagens Types VI, VII, VIII, IX, X, XIV, XVI and XIX (S. RicardBlum, B. Dublet and M. van der Rest, eds.), Oxford University Press, p. 155. Tu, H., Sasaki, T., Snellman, A., Gohring, W., Pirila, P., Timpl, R., and Pihlajaniemi, T. (2002). The type XIII collagen ectodomain is a 150-nm rod and capable of binding to fibronectin, nidogen-2, perlecan, and heparin. J. Biol. Chem. 277, 23092–23099. Nykvist, P., Tu, H., Ivaska, J., Kapyla, J., Pihlajaniemi, T., and Heino, J. (2000). Distinct recognition of collagen subtypes by alpha(1)beta(1) and alpha(2)beta(1) integrins. Alpha(1)beta(1) mediates cell adhesion to type XIII collagen. J. Biol. Chem. 275, 8255–8261. Juvonen, M., and Pihlajaniemi, T. (1992) Characterization of the spectrum of alternative splicing of alpha 1 (XIII) collagen transcripts in HT-1080 cells and calvarial tissue resulted in identification of two previously unidentified alternatively spliced sequences, one previously unidentified exon, and nine new mRNA variants. J. Biol. Chem. 267, 24693–24699. Rehn, M., and Pihlajaniemi, T. (1994). Alpha 1(XVIII), a collagen chain with frequent interruptions in the collagenous sequence, a distinct tissue distribution, and homology with type XV collagen. Proc. Natl. Acad. Sci. U.S.A. 91, 4234–4238. Rehn, M., Hintikka, E., and Pihlajaniemi, T. (1994). Primary structure of the alpha 1 chain of mouse type XVIII collagen, partial structure of the corresponding gene, and comparison of the alpha 1(XVIII) chain with its homologue, the alpha 1(XV) collagen chain. J. Biol. Chem. 269, 13929–13935. Pufe, T., Petersen, W. J., Miosge, N., Goldring, M. B., Mentlein, R., Varoga, D. J., and Tillmann, B. N. (2004). Endostatin/collagen XVIIIan inhibitor of angiogenesis-is expressed in cartilage and fibrocartilage. Matrix Biol. 23, 267–276. Sasaki, T., Fukai, N., Mann, K., Gohring, W., Olsen, B. R., and Timpl, R. (1998). Structure, function and tissue forms of the C-terminal globular domain of collagen XVIII containing the angiogenesis inhibitor endostatin. EMBO J. 17, 4249–4256. Olsen, B. R. (1981). Collagen Biosynthesis. In “Cell Biology of Extracellular Matrix” (E. D. Hay, ed), pp. 139–177, Plenum Press, New York. Ryan, M. C., Sieraski, M., and Sandell, L. J. (1990). The human type II procollagen gene: identification of an additional protein-coding domain and location of potential regulatory sequences in the promoter and first intron. Genomics 8, 41–48. Peltonen, S., Rehn, M., and Pihlajaniemi, T. (1997). Alternative splicing of mouse alpha1(XIII) collagen RNAs results in at least 17 different transcripts, predicting alpha1(XIII) collagen chains with length varying between 651 and 710 amino acid residues. DNA Cell Biol. 16, 227–234. Chu, M. L., de Wet, W., Bernard, M., and Ramirez, F. (1985). Fine structural analysis of the human pro-alpha 1 (I) collagen gene.

266.

267.

268.

269. 270.

271.

272.

273.

274.

275.

276.

277. 278.

279.

280.

281.

282.

33

Promoter structure, AluI repeats, and polymorphic transcripts. J. Biol. Chem. 260, 2315–2320. Focht, R. J., and Adams, S. L. (1984). Tissue specificity of type I collagen gene expression is determined at both transcriptional and post-transcriptional levels. Mol. Cell. Biol. 4, 1843–1852. Mauch, C., Hatamochi, A., Scharffetter, K., and Krieg, T. (1988). Regulation of collagen synthesis in fibroblasts within a three- dimensional collagen gel. Exp. Cell Res. 178, 493–503. Eckes, B., Mauch, C., Huppe, G., and Krieg, T. (1993). Downregulation of collagen synthesis in fibroblasts within three- dimensional collagen lattices involves transcriptional and posttranscriptional mechanisms. FEBS Lett. 318, 129–133. Kivirikko, K. I., and Myllyharju, J. (1998). Prolyl 4-hydroxylases and their protein disulfide isomerase subunit. Matrix Biol. 16, 357–368. Kivirikko, K. I., and Myllyla, R. (1982). Posttranslational enzymes in the biosynthesis of collagen: intracellular enzymes. Methods Enzymol 82, 245–304. Kivirikko, K. I., and Pihlajaniemi, T. (1998). Collagen hydroxylases and the protein disulfide isomerase subunit of prolyl 4-hydroxylases. Adv. Enzymol. Relat. Areas Mol. Biol. 72, 325–398. Vranka, J. A., Sakai, L. Y., and Bachinger, H. P. (2004). Prolyl 3-hydroxylase 1, enzyme characterization and identification of a novel family of enzymes. J. Biol. Chem. 279, 23615–23621. Tryggvason, K., Majamaa, K., Risteli, J., and Kivirikko, K. I. (1979). Partial purification and characterization of chick-embryo prolyl 3-hydroxylase. Biochem. J. 183, 303–307. Helaakoski, T., Veijola, J., Vuori, K., Rehn, M., Chow, L. T., TaillonMiller, P., Kivirikko, K. I., and Pihlajaniemi, T. (1994). Structure and expression of the human gene for the alpha subunit of prolyl 4-hydroxylase. The two alternatively spliced types of mRNA correspond to two homologous exons the sequences of which are expressed in a variety of tissues. J. Biol. Chem. 269, 27847–27854. Annunen, P., Helaakoski, T., Myllyharju, J., Veijola, J., Pihlajaniemi, T., and Kivirikko, K. I. (1997). Cloning of the human prolyl 4-hydroxylase alpha subunit isoform alpha(II) and characterization of the type II enzyme tetramer. The alpha(I) and alpha(II) subunits do not form a mixed alpha(I)alpha(II)beta2 tetramer. J. Biol. Chem. 272, 17342–17348. Chvapil, M., and Ryan, J. N. (1972). Diverse effect of 1,10-phenanthroline and 2,2′-dipyridyl on the synthesis of collagen and noncollagenous proteins in in vivo and in vitro systems. Biochim. Biophys. Acta 273, 208–211. Bienkowski, R. S. (1989). Intracellular degradation of newly synthesized collagen. Revis. Biol. Celular. 21, 423–443. Hanauske-Abel, H. M. (1991). Prolyl 4-hydroxylase, a target enzyme for drug development. Design of suppressive agents and the in vitro effects of inhibitors and proinhibitors. J. Hepatol. 13 Suppl 3:S8–15; discussion S16. Tschank, G., Brocks, D. G., Engelbart, K., Mohr, J., Baader, E., Gunzler, V., and Hanauske-Abel, H. M. (1991). Inhibition of prolyl hydroxylation and procollagen processing in chick-embryo calvaria by a derivative of pyridine-2,4- dicarboxylate. Characterization of the diethyl ester as a proinhibitor. Biochem. J. 275, 469–476. Myllyla, R., Pihlajaniemi, T., Pajunen, L., Turpeenniemi-Hujanen, T., and Kivirikko, K. I. (1991). Molecular cloning of chick lysyl hydroxylase. Little homology in primary structure to the two types of subunit of prolyl 4- hydroxylase. J. Biol. Chem. 266, 2805–2810. Rautavuoma, K., Takaluoma, K., Passoja, K., Pirskanen, A., Kvist, A. P., Kivirikko, K. I., and Myllyharju, J. (2002). Characterization of three fragments that constitute the monomers of the human lysyl hydroxylase isoenzymes 1–3. The 30-kDa N-terminal fragment is not required for lysyl hydroxylase activity. J. Biol. Chem. 277, 23084–23091. Smith-Mungo, L. I., and Kagan, H. M. (1998). Lysyl oxidase: properties, regulation and multiple functions in biology. Matrix Biol. 16, 387–398.

34 283. Knott, L., and Bailey, A. J. (1998). Collagen cross-links in mineralizing tissues: a review of their chemistry, function, and clinical relevance. Bone 22, 181–187. 284. Blumenkrantz, N., Assad, R., and Peterkofsky, B. (1984). Characterization of collagen hydroxylysyl glycosyltransferases as mainly intramembranous microsomal enzymes. J. Biol. Chem. 259, 854–859. 285. Clark, C. C. (1979). The distribution and initial characterization of oligosaccharide units on the COOH-terminal propeptide extensions of the pro-alpha 1 and pro-alpha 2 chains of type I procollagen. J. Biol. Chem. 254, 10798–10802. 286. Doege, K. J., and Fessler, J. H. (1986). Folding of carboxyl domain and assembly of procollagen I. J. Biol. Chem. 261, 8924–8935. 287. Freedman, R. B., Hirst, T. R., and Tuite, M. F. (1994). Protein disulphide isomerase: building bridges in protein folding. Trends. Biochem. Sci. 19, 331–336. 288. Lang, K., Schmid, F. X., and Fischer, G. (1987). Catalysis of protein folding by prolyl isomerase. Nature 329, 268–270. 289. Bulleid, N. J., Dalley, J. A., and Lees, J. F. (1997). The C-propeptide domain of procollagen can be replaced with a transmembrane domain without affecting trimer formation or collagen triple helix folding during biosynthesis. EMBO J. 16, 6694–6701. 290. Steinmann, B., Bruckner, P., and Superti-Furga, A. (1991). Cyclosporin A slows collagen triple-helix formation in vivo: indirect evidence for a physiologic role of peptidyl-prolyl cis-trans-isomerase. J. Biol. Chem. 266, 1299–1303. 291. Nakai, A., Satoh, M., Hirayoshi, K., and Nagata, K. (1992). Involvement of the stress protein HSP47 in procollagen processing in the endoplasmic reticulum. J. Cell Biol. 117, 903–914. 292. Nagata, K., and Hosokawa, N. (1996). Regulation and function of collagen-specific molecular chaperone, HSP47. Cell Struct. Funct. 21, 425–430. 293. Nagata, K. (1996). Hsp47: a collagen-specific molecular chaperone. Trends. Biochem. Sci. 21, 22–26. 294. Natsume, T., Koide, T., Yokota, S., Hirayoshi, K., and Nagata, K. (1994). Interactions between collagen-binding stress protein HSP47 and collagen. Analysis of kinetic parameters by surface plasmon resonance biosensor. J. Biol. Chem. 269, 31224–31228. 295. Hattori, T., Von Der Mark, K., Kawaki, H., Yutani, Y., Kubota, S., Nakanishi, T., Eberspaecher, H., De Crombrugghe, B., and Takigawa, M. (2004). Downregulation of rheumatoid arthritis-related antigen RAA47 (HSP47/colligin-2) in chondrocytic cell lines induces apoptosis and cell-surface expression of RA-A47 in association with CD9. J. Cell Physiol. 30, 30. 296. Rawlings, N. D., and Barrett, A. J. (1995). Evolutionary families of metallopeptidases. Methods Enzymol. 248, 183–228. 297. Hojima, Y., McKenzie, J. A., van der Rest, M., and Prockop, D. J. (1989). Type I procollagen N-proteinase from chick embryo tendons. Purification of a new 500-kDa form of the enzyme and identification of the catalytically active polypeptides. J. Biol. Chem. 264, 11336–11345. 298. Colige, A., Li, S. W., Sieron, A. L., Nusgens, B. V., Prockop, D. J., and Lapiere, C. M. (1997). cDNA cloning and expression of bovine procollagen I N-proteinase: a new member of the superfamily of zincmetalloproteinases with binding sites for cells and other matrix components. Proc. Natl. Acad. Sci. U.S.A. 94, 2374–2379. 299. Tuderman, L., and Prockop, D. J. (1982). Procollagen N-proteinase. Properties of the enzyme purified from chick embryo tendons. Eur. J. Biochem. 125, 545–549. 300. Li, S. W., Sieron, A. L., Fertala, A., Hojima, Y., Arnold, W. V., and Prockop, D. J. (1996). The C-proteinase that processes procollagens to fibrillar collagens is identical to the protein previously identified as bone morphogenic protein-1. Proc. Natl. Acad. Sci. U.S.A. 93, 5127–5130. 301. Suzuki, N., Labosky, P. A., Furuta, Y., Hargett, L., Dunn, R., Fogo, A. B., Takahara, K., Peters, D. M., Greenspan, D. S., and Hogan, B. L. (1996).

KLAUS VON DER MARK

302.

303.

304.

305.

306.

307.

308.

309.

310.

311.

312.

313.

314.

315.

316.

317.

318.

319.

Failure of ventral body wall closure in mouse embryos lacking a procollagen C-proteinase encoded by Bmp1, a mammalian gene related to Drosophila tolloid. Development 122, 3587–3595. Helseth, D. L., Jr., and Veis, A. (1984). Cathepsin D-mediated processing of procollagen: lysosomal enzyme involvement in secretory processing of procollagen. Proc. Natl. Acad. Sci. U.S.A. 81, 3302–3306. Hamalainen, E. R., Kemppainen, R., Pihlajaniemi, T., and Kivirikko, K. I. (1993). Structure of the human lysyl oxidase gene. Genomics 17, 544–548. Ramirez, F., and diLiberto, M. (1990). Complex and diversified regulatory programs control the expression of vertebrate collagen genes. FASEB J. 4, 1616–1623. Finer, M. H., Gerstenfeld, L. C., Young, D., Doty, P., and Boedtker, H. (1985). Collagen expression in embryonic chicken chondrocytes treated with phorbol myristate acetate. Mol. Cell Biol. 5, 1415–1424. Bennett, V. D., and Adams, S. L. (1987). Characterization of the translational control mechanism preventing synthesis of alpha 2(I) collagen in chicken vertebral chondroblasts. J. Biol. Chem. 262, 14806–14814. Bennett, V. D., and Adams, S. L. (1990). Identification of a cartilagespecific promoter within intron-2 of the chick alpha-2(i) collagen gene. J. Biol. Chem. 265, 2223–2230. Penttinen, R. P., Kobayashi, S., and Bornstein, P. (1988). Transforming growth factor beta increases mRNA for matrix proteins both in the presence and in the absence of changes in mRNA stability. Proc. Natl. Acad. Sci. U.S.A. 85, 1105–1108. Speth, C., and Oberbaumer, I. (1993). Expression of basement membrane proteins: evidence for complex post-transcriptional control mechanisms. Exp. Cell Res. 204, 302–310. Rossi, P., and de Crombrugghe, B. (1987). Formation of a type I collagen RNA dimer by intermolecular base-pairing of a conserved sequence around the translation initiation site. Nucleic Acids Res. 15, 8935–8956. Bornstein, P., McKay, J., Devarayalu, S., and Cook, S. C. (1988). A highly conserved, 5′ untranslated, inverted repeat sequence is ineffective in translational control of the alpha 1(I) collagen gene [published erratum appears in Nucleic Acids Res 1988 Dec 9; 16(23):11399]. Nucleic. Acids. Res. 16, 9721–9736. Paglia, L., Wilczek, J., de Leon, L. D., Martin, G. R., Horlein, D., and Muller, P. (1979). Inhibition of procollagen cell-free synthesis by amino-terminal extension peptides. Biochemistry 18, 5030–5034. Paglia, L. M., Wiestner, M., Duchene, M., Ouellette, L. A., Horlein, D., Martin, G. R., and Muller, P. K. (1981). Effects of procollagen peptides on the translation of type II collagen messenger ribonucleic acid and on collagen biosynthesis in chondrocytes. Biochemistry 20, 3523–3527. Horlein, D., McPherson, J., Goh, S. H., and Bornstein, P. (1981). Regulation of protein synthesis: translational control by procollagenderived fragments. Proc. Natl. Acad. Sci. U.S.A. 78, 6163–6167. Schlumberger, W., Thie, M., Volmer, H., Rauterberg, J., and Robenek, H. (1988). Binding and uptake of Col 1(I), a peptide capable of inhibiting collagen synthesis in fibroblasts. Eur. J. Cell Biol. 46, 244–252. Fouser, L., Sage, E. H., Clark, J., and Bornstein, P. (1991). Feedback regulation of collagen gene expression: a Trojan horse approach. Proc. Natl. Acad. Sci. U.S.A. 88, 10158–10162. Wu, C. H., Donovan, C. B., and Wu, G. Y. (1986). Evidence for pretranslational regulation of collagen synthesis by procollagen propeptides. J. Biol. Chem. 261, 10482–10484. Aycock, R. S., Raghow, R., Stricklin, G. P., Seyer, J. M., and Kang, A. H. (1986). Post-transcriptional inhibition of collagen and fibronectin synthesis by a synthetic homolog of a portion of the carboxyl- terminal propeptide of human type I collagen. J. Biol. Chem. 261, 14355–14360. Mizuno, M., Fujisawa, R., and Kuboki, Y. (2000). The effect of carboxyl-terminal propeptide of type I collagen (c-propeptide) on

CHAPTER 1 Structure, Biosynthesis and Gene Regulation of Collagens in Cartilage and Bone

320.

321. 322.

323.

324.

325.

326. 327.

328.

329.

330.

331.

332.

333.

334.

335.

collagen synthesis of preosteoblasts and osteoblasts. Calcif. Tissue Int. 67, 391–399. Nakata, K., Miyamoto, S., Bernier, S., Tanaka, M., Utani, A., Krebsbach, P., Rhodes, C., and Yamada, Y. (1996). The c-propeptide of type II procollagen binds to the enhancer region of the type II procollagen gene and regulates its transcription. Ann. Acad. Sci. N.Y. 785, 307–308. Rossert, J. A., and Garrett, L. A. (1995). Regulation of type I collagen synthesis. Kidney Int. Suppl. 49, S34–38. Bornstein, P. (1996). Regulation of expression of the alpha 1 (I) collagen gene: a critical appraisal of the role of the first intron. Matrix Biol. 15, 3–10. Frischauf, A. M., Lehrach, H., Rosner, C., and Boedtker, H. (1978). Procollagen complementary DNA, a probe for messenger RNA purification and the number of type I collagen genes. Biochemistry 17, 3243–3249. Boedtker, H., Frischauf, A. M., and Lehrach, H. (1976). Isolation and translation of calvaria procollagen messenger ribonucleic acids. Biochemistry 15, 4765–4770. Yamamoto, T., Sobel, M. E., Adams, S. L., Avvedimento, V. E., DiLauro, R., Pastan, I., de Crombrugghe, B., Showalter, A., Pesciotta, D., Fietzek, P., and Olsen, B. (1980). Construction of a recombinant bacterial plasmid containing pro-alpha 1(I) collagen DNA sequences. J. Biol. Chem. 255, 2612–2615. Vuorio, E., and de Crombrugghe, B. (1990). The family of collagen genes. Annu. Rev. Biochem. 59, 837–872. Chu, M. L., de Wet, W., Bernard, M., Ding, J. F., Morabito, M., Myers, J., Williams, C., and Ramirez, F. (1984). Human pro alpha 1(I) collagen gene structure reveals evolutionary conservation of a pattern of introns and exons. Nature 310, 337–340. Dewet, W., Bernard, M., Bensochanda, V., Chu, M. L., Dickson, L., Weil, D., and Ramirez, F. (1987). Organization of the human proalpha 2(I) collagen gene. J. Biol. Chem. 262, 16032–16036. Chuang, P. T., and McMahon, A. P. (1999). Vertebrate Hedgehog signalling modulated by induction of a Hedgehog-binding protein. Nature 397, 617–621. Sokolov, B. P., Mays, P. K., Khillan, J. S., and Prockop, D. J. (1993). Tissue- and development-specific expression in transgenic mice of a type I procollagen (COL1A1) minigene construct with 2.3 kb of the promoter region and 2 kb of the 3′-flanking region. Specificity is independent of the putative regulatory sequences in the first intron. Biochemistry 32, 9242–9249. Sokolov, B. P., Ala-Kokko, L., Dhulipala, R., Arita, M., Khillan, J. S., and Prockop, D. J. (1995). Tissue-specific expression of the gene for type I procollagen (COL1A1) in transgenic mice. Only 476 base pairs of the promoter are required if collagen genes are used as reporters. J. Biol. Chem. 270, 9622–9629. Rossert, J. A., Chen, S. S., Eberspaecher, H., Smith, C. N., and de Crombrugghe, B. (1996). Identification of a minimal sequence of the mouse pro-alpha 1(I) collagen promoter that confers high-level osteoblast expression in transgenic mice and that binds a protein selectively present in osteoblasts. Proc. Natl. Acad. Sci. U.S.A. 93, 1027–1031. Rossert, J., Eberspaecher, H., and de Crombrugghe, B. (1995). Separate cis-acting DNA elements of the mouse pro-alpha 1(I) collagen promoter direct expression of reporter genes to different type I collagen-producing cells in transgenic mice. J. Cell Biol. 129, 1421–1432. Bornstein, P., and McKay, J. (1988). The first intron of the alpha 1(I) collagen gene contains several transcriptional regulatory elements. J. Biol. Chem. 263, 1603–1606. Slack, J. L., Liska, D. J., and Bornstein, P. (1991). An upstream regulatory region mediates high-level, tissue- specific expression of the human alpha 1(I) collagen gene in transgenic mice. Mol. Cell Biol. 11, 2066–2074.

35

336. Liska, D. J., Robinson, V. R., and Bornstein, P. (1992). Elements in the first intron of the alpha 1(I) collagen gene interact with Sp1 to regulate gene expression. Gene Expr. 2, 379–389. 337. Kern, B., Shen, J., Starbuck, M., and Karsenty, G. (2001). Cbfa1 contributes to the osteoblast-specific expression of type I collagen genes. J. Biol. Chem. 276, 7101–7107. 338. Pavlin, D., Bedalov, A., Kronenberg, M. S., Kream, B. E., Rowe, D. W., Smith, C. L., Pike, J. W., and Lichtler, A. C. (1994). Analysis of regulatory regions in the COL1A1 gene responsible for 1,25- dihydroxyvitamin D3-mediated transcriptional repression in osteoblastic cells. J. Cell Biochem. 56, 490–501. 339. Meisler, N., Keefer, K. A., Ehrlich, H. P., Yager, D. R., Myers-Parrelli, J., and Cutroneo, K. R. (1997). Dexamethasone abrogates the fibrogenic effect of transforming growth factor-beta in rat granuloma and granulation tissue fibroblasts. J. Invest. Dermatol. 108, 285–289. 340. Meisler, N., Shull, S., Xie, R., Long, G. L., Absher, M., Connolly, J. P., and Cutroneo, K. R. (1995). Glucocorticoids coordinately regulate type I collagen pro alpha 1 promoter activity through both the glucocorticoid and transforming growth factor beta response elements: a novel mechanism of glucocorticoid regulation of eukaryotic genes. J. Cell Biochem. 59, 376–388. 341. Rossi, P., and deCrombrugghe, B. (1987). Identification of a cellspecific transcriptional enhancer in the first intron of the mouse alpha 2 (type I) collagen gene. Proc. Natl. Acad. Sci. USA 84, 5590–5594. 342. Ignotz, R. A., Endo, T., and Massague, J. (1987). Regulation of fibronectin and type I collagen mRNA levels by transforming growth factor-beta. J. Biol. Chem. 262, 6443–6446. 343. Ritzenthaler, J. D., Goldstein, R. H., Fine, A., and Smith, B. D. (1993). Regulation of the alpha 1(I) collagen promoter via a transforming growth factor-beta activation element. J. Biol. Chem. 268, 13625–13631. 344. D’Souza, R. N., Niederreither, K., and de Crombrugghe, B. (1993). Osteoblast-specific expression of the alpha 2(I) collagen promoter in transgenic mice: correlation with the distribution of TGF-beta 1. J. Bone Miner. Res. 8, 1127–1136. 345. Chung, K. Y., Agarwal, A., Uitto, J., and Mauviel, A. (1996). An AP-1 binding sequence is essential for regulation of the human alpha2(I) collagen (COL1A2) promoter activity by transforming growth factorbeta. J. Biol. Chem. 271, 3272–3278. 346. Greenwel, P., Inagaki, Y., Hu, W., Walsh, M., and Ramirez, F. (1997). Sp1 is required for the early response of alpha2(I) collagen to transforming growth factor-beta1. J. Biol. Chem. 272, 19738–19745. 347. Higashi, K., Kouba, D. J., Song, Y. J., Uitto, J., and Mauviel, A. (1998). A proximal element within the human alpha 2(I) collagen (COL1A2) promoter, distinct from the tumor necrosis factor-alpha response element, mediates transcriptional repression by interferongamma. Matrix Biol. 16, 447–456. 348. Bou-Gharios, G., Garrett, L. A., Rossert, J., Niederreither, K., Eberspaecher, H., Smith, C., Black, C., and Crombrugghe, B. (1996). A potent far-upstream enhancer in the mouse pro alpha 2(I) collagen gene regulates expression of reporter genes in transgenic mice. J. Cell Biol. 134, 1333–1344. 349. Horton, W., Miyashita, T., Kohno, K., Hassell, J. R., and Yamada, Y. (1987). Identification of a phenotype-specific enhancer in the first intron of the rat collagen II gene. Proc. Natl. Acad. Sci. USA 84, 8864–8868. 350. Zhou, G., Garofalo, S., Mukhopadhyay, K., Lefebvre, V., Smith, C. N., Eberspaecher, H., and de Crombrugghe, B. (1995). A 182 bp fragment of the mouse pro alpha 1(II) collagen gene is sufficient to direct chondrocyte expression in transgenic mice. J. Cell Sci. 108, 3677–3684. 351. Mukhopadhyay, K., Lefebvre, V., Zhou, G., Garofalo, S., Kimura, J. H., and de Crombrugghe, B. (1995). Use of a new rat chondrosarcoma cell line to delineate a 119- base pair chondrocyte-specific enhancer

36

352.

353.

354.

355.

356.

357.

358.

359. 360.

361.

362.

363.

364.

365.

366.

367.

KLAUS VON DER MARK element and to define active promoter segments in the mouse proalpha 1(II) collagen gene. J. Biol. Chem. 270, 27711–27719. Zhou, G., Lefebvre, V., Zhang, Z., Eberspaecher, H., and de Crombrugghe, B. (1998). Three high mobility group-like sequences within a 48-base pair enhancer of the Col2a1 gene are required for cartilage-specific expression in vivo. J. Biol. Chem. 273, 14989–14997. Lefebvre, V., Zhou, G., Mukhopadhyay, K., Smith, C. N., Zhang, Z., Eberspaecher, H., Zhou, X., Sinha, S., Maity, S. N., and de Crombrugghe, B. (1996). An 18-base-pair sequence in the mouse proalpha1(II) collagen gene is sufficient for expression in cartilage and binds nuclear proteins that are selectively expressed in chondrocytes. Mol. Cell Biol. 16, 4512–4523. Lefebvre, V., and de Crombrugghe, B. (1998). Toward understanding SOX9 function in chondrocyte differentiation. Matrix Biol. 16, 529–540. Savagner, P., Krebsbach, P. H., Hatano, O., Miyashita, T., Liebman, J., and Yamada, Y. (1995). Collagen II promoter and enhancer interact synergistically through Sp1 and distinct nuclear factors. DNA Cell Biol. 14, 501–510. Ruteshouser, E. C., and de Crombrugghe, B. (1992). Purification of BBF, a DNA-binding protein recognizing a positive cis-acting element in the mouse alpha 1(III) collagen promoter. J. Biol. Chem. 267, 14398–14404. Mudryj, M., and de Crombrugghe, B. (1988). Deletion analysis of the mouse alpha 1(III) collagen promoter. Nucleic. Acids. Res. 16, 7513–7526. Takahara, K., Hoffman, G. G., and Greenspan, D. S. (1995). Complete structural organization of the human alpha 1 (V) collagen gene (COL5A1): divergence from the conserved organization of other characterized fibrillar collagen genes. Genomics 29, 588–597. Lee, S., and Greenspan, D. S. (1995). Transcriptional promoter of the human alpha 1(V) collagen gene (COL5A1). Biochem. J. 310, 15–22. Truter, S., Di Liberto, M., Inagaki, Y., and Ramirez, F. (1992). Identification of an upstream regulatory region essential for cell typespecific transcription of the pro-alpha 2(V) collagen gene (COL5A2). J. Biol. Chem. 267, 25389–25395. Nagato, H., Matsuo, N., Sumiyoshi, H., Sakata-Takatani, K., Nasu, M., Yoshioka, H. (2004). The transcription factor CCAAT-binding -factor CBF/NF-Y and two repressors regulate the core promoter of the human Proa3(V) collagen gene(COL5A3). J. Biol. Chem. 279, in press. Yoshioka, H., Greenwel, P., Inoguchi, K., Truter, S., Inagaki, Y., Ninomiya, Y., and Ramirez, F. (1995). Structural and functional analysis of the promoter of the human alpha 1(XI) collagen gene. J. Biol. Chem. 270, 418–424. Yoshioka, H., Inoguchi, K., Khaleduzzaman, M., Ninomiya, Y., Andrikopoulos, K., and Ramirez, F. (1995). Coding sequence and alternative splicing of the mouse alpha 1(XI) collagen gene (Col11a1). Genomics 28, 337–340. Lui, V. C., Ng, L. J., Sat, E. W., and Cheah, K. S. (1996). The human alpha 2(XI) collagen gene (COL11A2): completion of coding information, identification of the promoter sequence, and precise localization within the major histocompatibility complex reveal overlap with the KE5 gene. Genomics 32, 401–412. Bridgewater, L. C., Lefebvre, V., and de Crombrugghe, B. (1998). Chondrocyte-specific enhancer elements in the Col11a2 gene resemble the Col2a1 tissue-specific enhancer. J. Biol. Chem. 273, 14998–15006. Hayman, A. R., Koppel, J., and Trueb, B. (1991). Complete structure of the chicken alpha 2(VI) collagen gene. Eur. J. Biochem. 197, 177–184. Trikka, D., Davis, T., Lapenta, V., Brahe, C., and Kessling, A. M. (1997). Human COL6A1: genomic characterization of the globular domains, structural and evolutionary comparison with COL6A2. Mamm. Genome. 8, 342–345.

368. Walchli, C., Marcionelli, R., Odermatt, B. F., Peltonen, J., Vuorio, E., and Trueb, B. (1996). Expression and distribution of two alternatively spliced transcripts from the chicken alpha 2 (VI) collagen gene. J. Cell Biochem. 63, 207–220. 369. Doliana, R., Bonaldo, P., and Colombatti, A. (1990). Multiple forms of chicken alpha 3(VI) collagen chain generated by alternative splicing in type A repeated domains. J. Cell Biol. 111, 2197–2205. 370. Braghetta, P., Fabbro, C., Piccolo, S., Marvulli, D., Bonaldo, P., Volpin, D., and Bressan, G. M. (1996). Distinct regions control transcriptional activation of the alpha1(VI) collagen promoter in different tissues of transgenic mice. J. Cell Biol. 135, 1163–1177. 371. Piccolo, S., Bonaldo, P., Vitale, P., Volpin, D., and Bressan, G. M. (1995). Transcriptional activation of the alpha 1(VI) collagen gene during myoblast differentiation is mediated by multiple GA boxes. J. Biol. Chem. 270, 19583–19590. 372. Fabbro, C., Braghetta, P., Girotto, D., Piccolo, S., Volpin, D., and Bressan, G. M. (1999). Cell type-specific transcription of the alpha1(VI) collagen gene. Role of the ap1 binding site and of the core promoter. J. Biol. Chem. 274, 1759–1766. 373. Saitta, B., and Chu, M. L. (1995). Characterization of the human alpha 1(VI) collagen promoter and its comparison with human alpha 2(VI) promoters. Eur. J. Biochem. 234, 542–549. 374. Braghetta, P., Vitale, P., Piccolo, S., Bonaldo, P., Fabbro, C., Girotto, D., Volpin, D., and Bressan, G. M. (1997). Tissue-specific expression of promoter regions of the alpha1(VI) collagen gene in cell cultures and transgenic mice. Eur. J. Biochem. 247, 200–208. 375. Willimann, T. E., Maier, R., and Trueb, B. (1995). A novel transcription factor and two members of the Sp 1 multigene family regulate the activity of the alpha 2 (VI) collagen promoter. Matrix Biol. 14, 653–663. 376. Hatamochi, A., Aumailley, M., Mauch, C., Chu, M.-L., Timpl, R., and Krieg, T. (1989). Regulation of collagen VI expression in fibroblasts. Effects of cell density, cell-matrix interactions, and chemical transformation. J. Biol. Chem. 264, 3494–3499. 377. Peltonen, J., Hsiao, L. L., Jaakkola, S., Sollberg, S., Aumailley, M., Timpl, R., Chu, M. L., and Uitto, J. (1991). Activation of collagen gene expression in keloids: co- localization of type I and VI collagen and transforming growth factor-beta 1 mRNA. J. Invest. Dermatol. 97, 240–248. 378. Harumiya, S., Gibson, M. A., and Koshihara, Y. (2002). Antisense suppression of collagen VI synthesis results in reduced expression of collagen I in normal human osteoblast-like cells. Biosci. Biotechnol. Biochem. 66, 2743–2747. 379. Heckmann, M., Aumailley, M., Chu, M. L., Timpl, R., and Krieg, T. (1992). Effect of transforming growth factor-beta on collagen type VI expression in human dermal fibroblasts. FEBS Lett. 310, 79–82. 380. Luvalle, P., Ninomiya, Y., and Rosenblum, N. D. (1988). The type X collagen gene. J. Biol. Chem. 263, 18378–18385. 381. Elima, K., Eerola, I., Rosati, R., Metsäranta, M., Garofalo, S., Perälä, M., deCrombrugghe, B., and Vuorio, E. (1993). The mouse collagen X gene: complete nucleotide sequence, exon structure and expression pattern. Biochem. J. 289, 247–253. 382. Kong, R. Y., Kwan, K. M., Lau, E. T., Thomas, J. T., BootHandford, R. P., Grant, M. E., and Cheah, K. S. (1993). Intron-exon structure, alternative use of promoter and expression of the mouse collagen X gene, COL10A-1. Eur. J. Biochem. 213, 99–111. 383. Apte, S. S., and Olsen, B. R. (1993). Characterization of the mouse type X collagen gene. Matrix 13, 165–179. 384. Thomas, J. T., Kwan, A. P. L., Grant, M. E., and Boot-Handford, R. P. (1991). Isolation of cDNAs encoding the complete sequence of bovine type X collagen. Biochem. J. 273, 141–148. 385. Thomas, J. T., Cresswell, C. J., Rash, B., Nicolai, H., Jones, T., Solomon, E., Grant, M. E., and Boot-Handford, R. P. (1991). The human collagen X gene. Biochem. J. 280, 617–623. 386. Reichenberger, E., Beier, F., Luvalle, P., Olsen, B. R., von der Mark, K., and Bertling, W. (1992). Genomic organisation and full length cDNA sequence of human collagen X. FEBS Lett. 311, 305–310.

CHAPTER 1 Structure, Biosynthesis and Gene Regulation of Collagens in Cartilage and Bone 387. Beier, F., Eerola, I., Vuorio, E., Luvalle, P., Reichenberger, E., Bertling, W., von der Mark, K., and Lammi, M. J. (1996). Variability in the upstream promoter and intron sequences of the human, mouse and chick type X collagen genes. Matrix Biol. 15, 415–422. 388. Luvalle, P., Daniels, K., Hay, E. D., and Olsen, B. (1992). Type X collagen is transcriptionally activated and specifically localized during sternal cartilage maturation. Matrix 12, 404–413. 389. Schmid, I. M., and Linsenmayer, T. F. (1985). Immunohistochemical localization of short chain cartilage collagen (type-X) in avian tissues. J. Cell Biol. 100, 598–605. 390. Reichenberger, E., Aigner, T., von der Mark, K., Stoess, H., and Bertling, W. (1991). In situ hybridization studies on the expression of type X collagen in fetal human cartilage. Develop. Biol. 148, 562–572. 391. Luvalle, P., Iwamoto, M., Fanning, P., Pacifici, M., and Olsen, B. R. (1993). Multiple negative elements in a gene that codes for an extracellular matrix protein, collagen X, restrict expression to hypertrophic chondrocytes. J. Cell Biol. 121, 1173–1179. 392. Dourado, G., and Luvalle, P. (1998). Proximal DNA elements mediate repressor activity conferred by the distal portion of the chicken collagen X promoter. J. Cell Biochem. 70, 507–516. 393. Long, F., Sonenshein, G. E., and Linsenmayer, T. F. (1998). Multiple transcriptional elements in the avian type X collagen gene. Identification of Sp1 family proteins as regulators for high level expression in hypertrophic chondrocytes. J. Biol. Chem. 273, 6542–6549. 394. Volk, S. W., Luvalle, P., Leask, T., and Leboy, P. S. (1998). A BMP responsive transcriptional region in the chicken type X collagen gene. J. Bone Miner. Res. 13, 1521–1529. 395. Beier, F., Vornehm, S., Pöschl, E., von der Mark, K., and Lammi, M. J. (1997). Localization of silencer and enhancer elements in the human type X collagen gene. J. Cell Biochem. 66, 210–218. 396. Chambers, D., Young, D. A., Howard, C., Thomas, J. T., Boam, D. S., Grant, M. E., Wallis, G. A., and Boot-Handford, R. P. (2002). An enhancer complex confers both high-level and cell-specific expression of the human type X collagen gene. FEBS Lett. 531, 505–508. 397. Riemer, S., Gebhard, S., Beier, F., Poschl, E., and von der Mark, K. (2002). Role of c-fos in the regulation of type X collagen gene expression by PTH and PTHrP: localization of a PTH/PTHrP-responsive region in the human COL10A1 enhancer. J. Cell Biochem. 86, 688–699. 398. Gebhard, S., Poschl, E., Riemer, S., Bauer, E., Hattori, T., Eberspaecher, H., Zhang, Z., Lefebvre, V., de Crombrugghe, B., and von der Mark, K. (2004). A highly conserved enhancer in mammalian type X collagen genes drives high levels of tissue-specific expression in hypertrophic cartilage in vitro and in vivo. Matrix Biol. 23, 309–322. 399. O’Keefe, R. J., Loveys, L. S., Hicks, D. G., Reynolds, P. R., Crabb, I. D., Puzas, J. E., and Rosier, R. N. (1997). Differential regulation of type-II and type-X collagen synthesis by parathyroid hormone-related protein in chick growth-plate chondrocytes. J. Orthop. Res. 15, 162–174. 400. Harada, S., Sampath, T. K., and Rodan, G. A. (1996). Identification of an AP1-like response region for osteogenic protein-1 in type X collagen promoter. Ann. Acad. Sci. N.Y. 785, 259–260. 401. Zheng, Q., Zhou, G., Morello, R., Chen, Y., Garcia-Rojas, X., and Lee, B. (2003). Type X collagen gene regulation by Runx2 contributes directly to its hypertrophic chondrocyte-specific expression in vivo. J. Cell Biol. 162, 833–842. 402. Ducy, P., Zhang, R., Geoffroy, V., Ridall, A. L., and Karsenty, G. (1997). Osf2/Cbfa1: a transcriptional activator of osteoblast differentiation. Cell 89, 747–754. 403. Otto, F., Thornell, A. P., Crompton, T., Denzel, A., Gilmour, K. C., Rosewell, I. R., Stamp, G. W., Beddington, R. S., Mundlos, S., Olsen, B. R., Selby, P. B., and Owen, M. J. (1997). Cbfa1, a candidate

404.

405.

406.

407.

408.

409.

410.

411.

412.

413. 414.

415.

416.

417.

37

gene for cleidocranial dysplasia syndrome, is essential for osteoblast differentiation and bone development. Cell 89, 765–771. Komori, T., Yagi, H., Nomura, S., Yamaguchi, A., Sasaki, K., Deguchi, K., Shimizu, Y., Bronson, R. T., Gao, Y. H., Inada, M., Sato, M., Okamoto, R., Kitamura, Y., Yoshiki, S., and Kishimoto, T. (1997). Targeted disruption of Cbfa1 results in a complete lack of bone formation owing to maturational arrest of osteoblasts. Cell 89, 755–764. Nakashima, K., Zhou, X., Kunkel, G., Zhang, Z., Deng, J. M., Behringer, R. R., and de Crombrugghe, B. (2002). The novel zinc finger-containing transcription factor osterix is required for osteoblast differentiation and bone formation. Cell 108, 17–29. Slavin, J., Unemori, E., Hunt, T. K., and Amento, E. (1994). Transforming growth factor beta (TGF-beta) and dexamethasone have direct opposing effects on collagen metabolism in low passage human dermal fibroblasts in vitro. Growth Factors. 11, 205–213. Murata, H., Zhou, L., Ochoa, S., Hasan, A., Badiavas, E., and Falanga, V. (1997). TGF-beta3 stimulates and regulates collagen synthesis through TGF-beta1- dependent and independent mechanisms. J. Invest. Dermatol. 108, 258–262. Inagaki, Y., Truter, S., and Ramirez, F. (1994). Transforming growth factor-beta stimulates alpha 2(I) collagen gene expression through a cis-acting element that contains an Sp1-binding site. J. Biol. Chem. 269, 14828–14834. Inagaki, Y., Nemoto, T., Nakao, A., Dijke, P., Kobayashi, K., Takehara, K., and Greenwel, P. (2001). Interaction between GC box binding factors and Smad proteins modulates cell lineage-specific alpha 2(I) collagen gene transcription. J. Biol. Chem. 276, 16573–16579. Chen, S. J., Yuan, W., Lo, S., Trojanowska, M., and Varga, J. (2000). Interaction of smad3 with a proximal smad-binding element of the human alpha2(I) procollagen gene promoter required for transcriptional activation by TGF-beta. J. Cell Physiol. 183, 381–392. Wang, E. A., Rosen, V., D’Alessando, J., Bauduy, M., Cordes, P., Harada, T., Israk, D. I., Hewick, R. M., Kern, K. M., LaDan, P., Luxenberg, D. P., McQuaid, D., Mountsatsos, J. K., Nove, J., and Wozney, J. M. (1990). Recombinant human bone morphogenetic protein induces bone formation. PNAS 87, 2220–2224. Sampath, T. K., Maliakal, J. C., Hauschka, P. V., Jones, W. K., Sasak, H., Tucker, R. F., White, K. H., Coughlin, J. E., Tucker, M. M., and Pang, R. H. (1992). Recombinant human osteogenic protein-1 (hOP-1) induces new bone formation in vivo with a specific activity comparable with natural bovine osteogenic protein and stimulates osteoblast proliferation and differentiation in vitro. J. Biol. Chem 267, 20352–20362. Reddi, A. H. (1998). Initiation of fracture repair by bone morphogenetic proteins. Clin. Orthop. S66–72. Fujii, M., Takeda, K., Imamura, T., Aoki, H., Sampath, T. K., Enomoto, S., Kawabata, M., Kato, M., Ichijo, H., and Miyazono, K. (1999). Roles of bone morphogenetic protein type I receptors and Smad proteins in osteoblast and chondroblast differentiation. Mol. Biol. Cell 10, 3801–3813. Takagi, M., Kamiya, N., Takahashi, T., Ito, S., Hasegawa, M., Suzuki, N., and Nakanishi, K. (2004). Effects of bone morphogenetic protein-2 and transforming growth factor beta1 on gene expression of transcription factors, AJ18 and Runx2 in cultured osteoblastic cells. J. Mol. Histol. 35, 81–90. Lepisto, J., Peltonen, J., Vaha-Kreula, M., Soderstrom, K., Niinikoski, J., and Laato, M. (1996). Selective modulation of collagen gene expression by different isoforms of platelet-derived growth factor in experimental wound healing. Cell Tissue Res. 286, 449–455. Hata, R., Sunada, H., Arai, K., Sato, T., Ninomiya, Y., Nagai, Y., and Senoo, H. (1988). Regulation of collagen metabolism and cell growth by epidermal growth factor and ascorbate in cultured human skin fibroblasts [published erratum appears in Eur J Biochem 1988 Aug 15;175(3): 679]. Eur. J. Biochem. 173, 261–267.

38 418. Rowe, D. W., and Kream, B. E. (1982). Regulation of collagen synthesis in fetal rat calvaria by 1,25- dihydroxyvitamin D3. J. Biol. Chem. 257, 8009–8015. 419. Kuroki, Y., Shiozawa, S., Kano, J., and Chihara, K. (1995). Competition between c-fos and 1,25(OH)2 vitamin D3 in the transcriptional control of type I collagen synthesis in MC3T3-E1 osteoblastic cells. J. Cell Physiol. 164, 459–464. 420. Kream, B. E., Lafrancis, D., Petersen, D. N., Woody, C., Clark, S., Rowe, D. W., and Lichtler, A. (1993). Parathyroid hormone represses alpha 1(I) collagen promoter activity in cultured calvariae from neonatal transgenic mice. Mol. Endocrinol. 7, 399–408. 421. Russell, S. B., Trupin, J. S., Myers, J. C., Broquist, A. H., Smith, J. C., Myles, M. E., and Russell, J. D. (1989). Differential glucocorticoid regulation of collagen mRNAs in human dermal fibroblasts. Keloidderived and fetal fibroblasts are refractory to down-regulation. J. Biol. Chem. 264, 13730–13735. 422. Oikarinen, A., Autio, P., Vuori, J., Vaananen, K., Risteli, L., Kiistala, U., and Risteli, J. (1992). Systemic glucocorticoid treatment decreases serum concentrations of carboxyterminal propeptide of type I procollagen and aminoterminal propeptide of type III procollagen. Br. J. Dermatol. 126, 172–178. 423. Silbermann, M. M. M. (1987). Dexamethasone impairs growth and collagen synthesis in condylar cartilage in vitro. Bone and Mineral 2, 87–106. 424. Advani, S., Lafrancis, D., Bogdanovic, E., Taxel, P., Raisz, L. G., and Kream, B. E. (1997). Dexamethasone suppresses in vivo levels of bone collagen synthesis in neonatal mice. Bone 20, 41–46. 425. Kream, B. E., Tetradis, S., Lafrancis, D., Fall, P. M., Feyen, J. H., and Raisz, L. G. (1997). Modulation of the effects of glucocorticoids on collagen synthesis in fetal rat calvariae by insulin-like growth factor binding protein-2. J. Bone Miner. Res. 12, 889–895. 426. Bird, J. L., and Tyler, J. A. (1994). Dexamethasone potentiates the stimulatory effect of insulin-like growth factor-I on collagen production in cultured human fibroblasts. J. Endocrinol. 142, 571–579. 427. Ireland, D. C., Bord, S., Beavan, S. R., and Compston, J. E. (2002). Effects of estrogen on collagen synthesis by cultured human osteoblasts depend on the rate of cellular differentiation. J. Cell Biochem. 86, 251–257. 428. Simeone, A., Acampora, D., Pannese, M., D’Esposito, M., Stornaiuolo, A., Gulisano, M., Mallamaci, A., Kastury, K., Druck, T., and Huebner, K. (1994). Cloning and characterization of two members of the vertebrate Dlx gene family. Proc. Natl Acad. Sci. USA 91, 2250–2254. 429. Tadic, T., Dodig, M., Erceg, I., Marijanovic, I., Mina, M., Kalajzic, Z., Velonis, D., Kronenberg, M. S., Kosher, R. A., Ferrari, D., and Lichtler, A. C. (2002). Overexpression of Dlx5 in chicken calvarial cells accelerates osteoblastic differentiation. J. Bone Miner. Res. 17, 1008–1014. 430. Ducy, P. (2000). Cbfa1: a molecular switch in osteoblast biology. Dev. Dyn. 219, 461–471. 431. Karsenty, G., Ducy, P., Starbuck, M., Priemel, M., Shen, J., Geoffroy, V., and Amling, M. (1999). Cbfa1 as a regulator of osteoblast differentiation and function. Bone 25, 107–108. 432. Komori, T. (2000). A fundamental transcription factor for bone and cartilage. Biochem. Biophys. Res. Commun. 276, 813–816. 433. Kim, I. S., Otto, F., Zabel, B., and Mundlos, S. (1999). Regulation of chondrocyte differentiation by Cbfa1. Mech. Dev. 80, 159–170. 434. Thunyakitpisal, P., Alvarez, M., Tokunaga, K., Onyia, J. E., Hock, J., Ohashi, N., Feister, H., Rhodes, S. J., and Bidwell, J. P. (2001). Cloning and functional analysis of a family of nuclear matrix transcription factors (NP/NMP4) that regulate type I collagen expression in osteoblasts. J. Bone Miner. Res. 16, 10–23. 435. Wagner, E. F. (2002). Functions of AP1 (Fos/Jun) in bone development. Ann. Rheum. Dis. 61, ii40–42.

KLAUS VON DER MARK 436. Grigoriadis, A. E., Wang, Z. Q., and Wagner, E. F. (1995). Fos and bone cell development: lessons from a nuclear oncogene. Trends Genet. 11, 436–441. 437. Kuroki, Y., Shiozawa, S., Sugimoto, T., and Fujita, T. (1992). Constitutive expression of c-fos gene inhibits type 1 collagen synthesis in transfected osteoblasts. Biochem. Biophys. Res. Commun. 182, 1389–1394. 438. Jinnin, M., Ihn, H., Yamane, K., and Tamaki, K. (2004). Interleukin13 stimulates the transcription of the human alpha2(I) collagen gene in human dermal fibroblasts. J. Biol. Chem. 279, 41783–41791. 439. Inagaki, Y., Nemoto, T., Kushida, M., Sheng, Y., Higashi, K., Ikeda, K., Kawada, N., Shirasaki, F., Takehara, K., Sugiyama, K., Fujii, M., Yamauchi, H., Nakao, A., de Crombrugghe, B., Watanabe, T., and Okazaki, I. (2003). Interferon alfa down-regulates collagen gene transcription and suppresses experimental hepatic fibrosis in mice. Hepatology 38, 890–899. 440. Rosenbloom, J., Feldman, G., Freundlich, B., and Jimenez, S. A. (1986). Inhibition of excessive scleroderma fibroblast collagen production by recombinant gamma-interferon. Association with a coordinate decrease in types I and III procollagen messenger RNA levels. Arthritis Rheum. 29, 851–856. 441. Scharffetter, K., Heckmann, M., Hatamochi, A., Mauch, C., Stein, B., Riethmuller, G., Ziegler-Heitbrock, H. W., and Krieg, T. (1989). Synergistic effect of tumor necrosis factor-alpha and interferongamma on collagen synthesis of human skin fibroblasts in vitro. Exp. Cell Res. 181, 409–419. 442. Kahari, V. M., Chen, Y. Q., Su, M. W., Ramirez, F., and Uitto, J. (1990). Tumor necrosis factor-alpha and interferon-gamma suppress the activation of human type I collagen gene expression by transforming growth factor-beta 1. Evidence for two distinct mechanisms of inhibition at the transcriptional and posttranscriptional levels. J. Clin. Invest. 86, 1489–1495. 443. Bird, J. L., and Tyler, J. A. (1995). Tumour necrosis factor alpha, interferon gamma and dexamethasone regulate IGF-I-maintained collagen production in cultured human fibroblasts. J. Endocrinol. 147, 167–176. 444. Xu, Y., Wang, L., Buttice, G., Sengupta, P. K., and Smith, B. D. (2003). Interferon gamma repression of collagen (COL1A2) transcription is mediated by the RFX5 complex. J. Biol. Chem. 278, 49134–49144. 445. Sengupta, P. K., Fargo, J., and Smith, B. D. (2002). The RFX family interacts at the collagen (COL1A2) start site and represses transcription. J. Biol. Chem. 277, 24926–24937. 446. Wright, E., Hargrave, M. R., Christiansen, J., Cooper, L., Kun, J., Evans, T., Gangadharan, U., Greenfield, A., and Koopman, P. (1995). The Sry-related gene Sox9 is expressed during chondrogenesis in mouse embryos. Nat. Genet. 9, 15–20. 447. Zhao, Q., Eberspaecher, H., Lefebvre, V., and de Crombrugghe, B. (1997). Parallel expression of Sox9 and Col2a1 in cells undergoing chondrogenesis. Dev. Dyn. 209, 377–386. 448. Ng, L. J., Wheatley, S., Muscat, G. E., Conway-Campbell, J., Bowles, J., Wright, E., Bell, D. M., Tam, P. P., Cheah, K. S., and Koopman, P. (1997). SOX9 binds DNA, activates transcription, and coexpresses with type II collagen during chondrogenesis in the mouse. Dev. Biol. 183, 108–121. 449. Foster, J. W. (1996). Mutations in SOX9 cause both autosomal sex reversal and campomelic dysplasia. Acta Paediatr. Jpn. 38, 405–411. 450. Giese, K., Pagel, J., and Grosschedl, R. (1994). Distinct DNA-binding properties of the high mobility group domain of murine and human SRY sex-determining factors. Proc. Natl Acad. Sci. USA 91, 3368–3372. 451. Lefebvre, V., Huang, W., Harley, V. R., Goodfellow, P. N., and de Crombrugghe, B. (1997). SOX9 is a potent activator of the chondrocyte-specific enhancer of the pro alpha1(II) collagen gene. Mol. Cell Biol. 17, 2336–2346.

CHAPTER 1 Structure, Biosynthesis and Gene Regulation of Collagens in Cartilage and Bone 452. Bell, D. M., Leung, K. K., Wheatley, S. C., Ng, L. J., Zhou, S., Ling, K. W., Sham, M. H., Koopman, P., Tam, P. P., and Cheah, K. S. (1997). SOX9 directly regulates the type-II collagen gene. Nat. Gen. 16, 174–178. 453. Huang, W., Zhou, X., Lefebvre, V., and de Crombrugghe, B. (2000). Phosphorylation of SOX9 by cyclic AMP-dependent protein kinase A enhances SOX9’s ability to transactivate a Col2a1 chondrocytespecific enhancer. Mol. Cell. Biol. 20, 4149–4158. 454. Bi, W., Huang, W., Whitworth, D. J., Deng, J. M., Zhang, Z., Behringer, R. R., and de Crombrugghe, B. (2001). Haploinsufficiency of Sox9 results in defective cartilage primordia and premature skeletal mineralization. Proc. Natl. Acad. Sci. USA 98, 6698–6703. 455. Bi, W., Deng, J. M., Zhang, Z., Behringer, R. R., and de Crombrugghe, B. (1999). Sox9 is required for cartilage formation. Nat. Genet. 22, 85–89. 456. Lefebvre, V., Behringer, R. R., and de Crombrugghe, B. (2001). L-Sox5, Sox6 and Sox9 control essential steps of the chondrocyte differentiation pathway. Osteoarthritis Cartilage 9 Suppl A, S69–75. 457. Joyce, M. E., Roberts, A. B., Sporn, M. B., and Bolander, M. E. (1990). Transforming growth factor-β and the initiation of chondrogenesis and osteogenesis in the rat femur. J. Cell Biol. 110, 2195–2207. 458. Redini, F., Galera, P., Mauviel, A., Loyau, G., and Pujol, J. P. (1988). Transforming growth factor β stimulates collagen and glycosaminoglycan biosynthesis in cultured rabbit articular chondrocytes. FEBS Lett. 234, 172–176. 459. Galera, P., Vivien, D., Pronost, S., Bonaventure, J., Redini, F., Loyau, G., and Pujol, J. P. (1992). Transforming growth factor-beta 1 (TGF-beta 1) up-regulation of collagen type II in primary cultures of rabbit articular chondrocytes (RAC) involves increased mRNA levels without affecting mRNA stability and procollagen processing. J. Cell Physiol. 153, 596–606. 460. Pujol, J. P., Galera, P., Pronost, S., Boumediene, K., Vivien, D., Macro, M., Min, W., Redini, F., Penfornis, H., and Daireaux, M. (1994). Transforming growth factor-beta (TGF-beta) and articular chondrocytes. Ann. Endocrinol. (Paris) 55, 109–120. 461. Chadjichristos, C., Ghayor, C., Herrouin, J. F., Ala-Kokko, L., Suske, G., Pujol, J. P., and Galera, P. (2002). Down-regulation of human type II collagen gene expression by transforming growth factor-beta 1 (TGFbeta 1) in articular chondrocytes involves SP3/SP1 ratio. J. Biol. Chem. 277, 43903–43917. 462. Bradham, D. M., in der Wiesche, B., Precht, P., Balakir, R., and Horton, W. (1994). Transrepression of type II collagen by TGF-beta and FGF is protein kinase C dependent and is mediated through regulatory sequences in the promoter and first intron. J. Cell Physiol. 158, 61–68. 463. Chen, P., Carrington, J. L., Hammonds, R. G., and Reddi, A. H. (1991). Stimulation of chondrogenesis in limb bud mesoderm cells by recombinant human bone morphogenetic protein 2B (BMP-2B) and modulation by transforming growth factor β1 and β2. Exp. Cell. Res. 195, 509–515. 464. Pizette, S., and Niswander, L. (2000). BMPs are required at two steps of limb chondrogenesis: formation of prechondrogenic condensations and their differentiation into chondrocytes. Dev. Biol. 219, 237–249. 465. Hanada, K., Solchaga, L. A., Caplan, A. I., Hering, T. M., Goldberg, V. M., Yoo, J. U., and Johnstone, B. (2001). BMP-2 induction and TGF-beta 1 modulation of rat periosteal cell chondrogenesis. J. Cell Biochem. 81, 284–294. 466. Shukunami, C., Ohta, Y., Sakuda, M., and Hiraki, Y. (1998). Sequential progression of the differentiation program by bone morphogenetic protein-2 in chondrogenic cell line ATDC5. Exp. Cell Res. 241, 1–11. 467. Valcourt, U., Ronziere, M. C., Winkler, P., Rosen, V., Herbage, D., and Mallein-Gerin, F. (1999). Different effects of bone morphogenetic proteins 2, 4, 12, and 13 on the expression of cartilage and

468.

469.

470.

471.

472.

473. 474.

475.

476.

477.

478. 479.

480.

481.

482.

483.

484.

485.

39

bone markers in the MC615 chondrocyte cell line. Exp. Cell Res. 251, 264–274. Ahrens, M., Ankenbauer, T., Schroder, D., Hollnagel, A., Mayer, H., and Gross, G. (1993). Expression of human bone morphogenetic proteins-2 or -4 in murine mesenchymal progenitor C3H10T1/2 cells induces differentiation into distinct mesenchymal cell lineages. DNA Cell Biol. 12, 871–880. Hatakeyama, Y., Nguyen, J., Wang, X., Nuckolls, G. H., and Shum, L. (2003). Smad signaling in mesenchymal and chondroprogenitor cells. J. Bone Joint Surg. Am. 85-A Suppl 3, 13–18. Leboy, P. S., Sullivan, T. A., Nooreyazdan, M., and Venezian, R. A. (1997). Rapid chondrocyte maturation by serum-free culture with BMP-2 and ascorbic acid. J. Cell Biochem. 66, 394–403. Volk, S. W., D’Angelo, M., Diefenderfer, D., and Leboy, P. S. (2000). Utilization of bone morphogenetic protein receptors during chondrocyte maturation. J. Bone Miner. Res. 15, 1630–1639. Drissi, M. H., Li, X., Sheu, T. J., Zuscik, M. J., Schwarz, E. M., Puzas, J. E., Rosier, R. N., and O’Keefe, R. J. (2003). Runx2/Cbfa1 stimulation by retinoic acid is potentiated by BMP2 signaling through interaction with Smad1 on the collagen X promoter in chondrocytes. J. Cell Biochem. 90, 1287–1298. Benya, P. D., and Shaffer, J. D. (1982). Dedifferentiated chondrocytes reexpress the differentiated collagen phenotype. Cell 30, 215–224. Dietz, U., Aigner, T., Bertling, W., and von der Mark, K. (1993). Alterations of collagen mRNA expression during retinoic acid induced chondrocyte modulation: Absence of untranslated alpha1(I) mRNA in hyaline chondrocytes. J. Cell Biochem. 52, 57–68. Benya, P., and Padilla, P. D. (1986). Modulation of the rabbit chondrocyte phenotype by retinoic acid terminates type II collagen synthesis without inducing type I collagen: The modulated phenotype differs from that produced by subculture. Develop. Biol. 118, 296–305. Paulsen, D. F., Langille, R. M., Dress, V., and Solursh, M. (1988). Selective stimulation of in vitro limb-bud chondrogenesis by retinoic acid. Differentiation 39, 123–130. Pacifici, M., Golden, E. B., Iwamoto, M., and Adams, S. L. (1991). Retinoic acid treatment induces type X collagen gene expression in cultured chick chondrocytes. Exp. Cell Res. 195, 38–46. Lee, G. S., Kochhar, D. M., and Collins, M. D. (2004). Retinoidinduced limb malformations. Curr. Pharm. Des. 10, 2657–2699. Cho, S. H., Oh, C. D., Kim, S. J., Kim, I. C., and Chun, J. S. (2003). Retinoic acid inhibits chondrogenesis of mesenchymal cells by sustaining expression of N-cadherin and its associated proteins. J. Cell Biochem. 89, 837–847. Luyten, F. P., Hascall, V. C., Nissley, S. P., Morales, T. I., and Reddi, A. H. (1988). Insulin-like growth factors maintain steady-state metabolism of proteoglycans in bovine articular cartilage explants. Arch Biochem. Biophys. 267, 416–425. Loeser, R. F., Todd, M. D., and Seely, B. L. (2003). Prolonged treatment of human osteoarthritic chondrocytes with insulin-like growth factor-I stimulates proteoglycan synthesis but not proteoglycan matrix accumulation in alginate cultures. J. Rheumatol. 30, 1565–1570. Sonal, D. (2001). Prevention of IGF-1 and TGFbeta stimulated type II collagen and decorin expression by bFGF and identification of IGF-1 mRNA transcripts in articular chondrocytes. Matrix Biol. 20, 233–242. Wroblewski, J., and Edwall-Arvidsson, C. (1995). Inhibitory effects of basic fibroblast growth factor on chondrocyte differentiation. J. Bone Miner. Res. 10, 735–742. Vortkamp, A., Lee, K., Lanske, B., Segre, G. V., Kronenberg, H. M., and Tabin, C. J. (1996). Regulation of rate of cartilage differentiation by Indian hedgehog and PTH-related protein. Science 273, 613–622. Vortkamp, A. (2000). The Indian hedgehog–PTHrP system in bone development. Ernst. Schering Res. Found Workshop, 191–209.

40 486. Weir, E. C., Philbrick, W. M., Amling, M., Neff, L. A., Baron, R., and Broadus, A. E. (1996). Targeted overexpression of parathyroid hormone-related peptide in chondrocytes causes chondrodysplasia and delayed endochondral bone formation. Proc. Natl Acad. Sci. U.S.A. 93, 10240–10245. 487. Amizuka, N., Warshawsky, H., Henderson, J. E., Goltzman, D., and Karaplis, A. C. (1994). Parathyroid hormone-related peptidedepleted mice show abnormal epiphyseal cartilage development and altered endochondral bone formation. J. Cell Biol. 126, 1611–1623. 488. Iwamoto, M., Jikko, A., Murakami, H., Shimazu, A., Nakashima, K., Takigawa, M., Baba, H., Suzuki, I., and Kato, Y. (1994). Changes in parathyroid hormone receptors during chondrocyte differentiation. J. Biol. Chem. 269, 17245–17251. 489. Crabb, I. D., O’Keefe, R. J., Puzas, J. E., and Rosier, R. N. (1992). Differential effects of parathyroid hormone on chick growth plate and articular chondrocytes. Calcif. Tissue Int. 50, 61–66.

KLAUS VON DER MARK 490. Goldring, M., Birkhead, J., Sandell, L., Kimura, T., and Krane, S. (1988). Interleukin 1 supresses expression of cartilage-specific types II and IX collagens and increases types I and III collagens in human chondrocytes. J. Clin. Inv. 82, 2026–2037. 491. Goldring, M. B., Fukuo, K., Birkhead, J. R., Dudek, E., and Sandell, L. J. (1994). Transcriptional suppression by interleukin-1 and interferon- gamma of type II collagen gene expression in human chondrocytes. J. Cell Biochem. 54, 85–99. 492. Goldring, M. B., Birkhead, J. R., Suen, L. F., Yamin, R., Mizuno, S., Glowacki, J., Arbiser, J. L., and Apperley, J. F. (1994). Interleukin-1 beta-modulated gene expression in immortalized human chondrocytes. J. Clin. Invest. 94, 2307–2316. 493. Pinnell, S. R. (1985). Regulation of collagen biosynthesis by ascorbic acid: a review. Yale J. Biol. Med. 58, 553–559. 494. Schwarz, R. I. (1985). Procollagen secretion meets the minimum requirements for the rate- controlling step in the ascorbate induction of procollagen synthesis. J. Biol. Chem. 260, 3045–3049.

Chapter 2

Fibrillogenesis and Maturation of Collagens Simon P. Robins

Matrix Biochemistry Group, Rowett Research Institute, Bucksburn, Aberdeen AB21 9SB

I. Introduction II. Fibrillogenesis III. Cross-linking

IV. Concluding Remarks References

I. INTRODUCTION

and, in addition to containing mixtures of fibrillar collagens, fibrils often incorporate a number of FACIT (Fibril Associated with Interrupted Triple-helix) collagens, particularly types IX, XII, XIV, and XIX. Such heterotypic fibrils are prevalent in cartilage but their structure and assembly will not be discussed here in detail (see Chapter 25). Proteoglycans, such as decorin, fibromodulin, and biglycan, together with other matrix constituents, have important influences on collagen fibril formation and, although the structure and biosynthesis of these components is discussed elsewhere in this volume (Chapters 4 and 5), their influence on fibrillogenesis will be reviewed briefly. A major factor in the functional capacity of collagen fibrils is the cross-linking process that imparts the tensile properties and resistance to enzymatic degradation. The initial cross-linking reactions and the spontaneous processes that lead to mature collagen fibrils will be discussed in detail, but changes associated with true aging and senescence that might be defined as being deleterious to function are beyond the scope of this review. Several reviews have been published on collagen fibrillogenesis [1, 2] and on cross-linking [3, 4].

The biophysical characteristics of bone and cartilage are governed to a large degree by the properties of their respective collagens, which constitute the majority of the protein content of these tissues. The properties of the collagens are, in turn, a consequence primarily of their fibrillar structure, and the characteristics of collagen fibrils dictate not only the biomechanical properties but also many other metabolic processes, including the susceptibility to degradation. The factors that control collagen fibrillogenesis and the subsequent stabilization of the fibrils are therefore crucial to an understanding of the dynamics of bone and cartilage metabolism. There are some 27 genetically distinct forms of collagen, but for the present discussion we shall be particularly concerned with those types that spontaneously self-assemble to form fibrils: collagens I, II, III, V, and XI. Of these fibrillar collagens, type I is the major component of bone and collagen II is the main collagenous component of cartilage. It has become increasingly clear, however, that collagen fibrils rarely comprise a single collagen type Dynamics of Bone and Cartilage Metabolism

41

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42

Simon P. Robins

II. FIBRILLOGENESIS Most studies of fibril formation have been performed using collagen I and discussion here will be limited to this type of collagen. The main steps involved in fibrillogenesis are illustrated in Figure 1, although studies by Kadler et al. [5] have led to new concepts on the time-course and control of this process during embryonic growth, as will be described in more detail later in this section. As the collagen is synthesized and secreted as a precursor procollagen molecule with globular extensions at both the N- and C-terminal ends (see Chapter 1), the initial events are removal of the extension propeptides, which occurs through the action of discrete proteinases. One function of the propeptides is to prevent inappropriate intracellular fibril formation, since the released triplehelical collagen monomers have an intrinsic property to self assemble into fibrils arranged in a quarter-staggered array. This organization is mainly an entropy-driven process occurring through disruption of ordered solvent molecules at the monomer surface, and enhanced by specific ionic interactions between neighboring molecules to give a 4D overlap (see Fig. 1). This gap and overlap structure gives rise to the 67-nm banding pattern revealed in the electron microscope after negative staining with phosphotungstic acid where the excess stain enters the gap regions of the fibril. Removal of the N-propeptides intact is achieved by a family of proteases including a disintegrin and metalloprotease with thrombospondin motifs (ADAMTS)-2,

which is thought to act on procollagen I [6]. A related enzyme, ADAMTS-3, is prevalent in cartilage [7] and is likely to process procollagen II, although other members of this family of proteases, including ADAMTS-14, have been described [8]. C-proteinase activity, a property of all members of the tolloid family of metalloproteinases including bone morphogenetic protein-1 (BMP-1) [9], cleaves procollagens I, II, and III at a single locus releasing the propeptide intact. A procollagen C-proteinase enhancer (PCPE) has been isolated; this 55-kDa glycoprotein appears to facilitate cleavage by binding to sites either side of the procollagen cleavage site [10]. The N-proteinase binds to one of the FACIT collagens, type XIV [11], and the C-proteinase has been shown to bind the α1β1 and α2β1 integrins [12]. These proteinases may therefore have important biological functions in addition to their role in processing procollagens. Much of the information currently available on the mechanisms of fibril assembly has come from cell-free systems using purified proteinases to generate collagen monomers de novo [13]. Under these conditions, after a relatively short “nucleation” period, intermediate filaments with a blunt (β) end and a pointed (α) end are formed with growth occurring at the pointed end. Subsequently, growth of the fibril from the blunt end is initiated giving rise to a parabola-shaped intermediate fibril. Initial studies showed that the intermediate filaments were bipolar with the N-terminals oriented towards both of the pointed ends, thus necessitating a change in orientation at some

Figure 1 Fibril formation and structure: following secretion of procollagen, the propeptides are cleaved by an N-protease (PNP) and C-protease (BMP-1); intermediate fibrils assemble with collagen monomers in a regular, quarter-staggered array (see inset) that gives rise to 67 nm banding; longer fibrils are formed by fusion of intermediates which may also occur laterally. The assembly process is influenced by interactions of both procollagen and the fibrils with proteoglycans and other matrix molecules (not shown).

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point along the filament. This type of intermediate filament was subsequently shown to occur in vivo [14]. Fibrils produced from acid-extracted collagen monomers are unipolar and the initial presence of the C-propeptide during fibril formation appears to have a major effect [1]. Both longitudinal and lateral growth of the collagen fibril are believed to occur through fusion of these intermediates. Some studies have shown that only unipolar intermediates undergo fusion; indeed, regulation of the formation of these two types of intermediate may have important influences on fibril growth. Other studies of developing chick tendon support the concept of fibril growth by fusion of fibril segments [15], but they also found a more rapid growth in length with development so that, in an 18-day tendon, no small, intermediate fibril segments were observed [16]. At this stage, analyses of serial sections by transmission electron microscopy showed that fibril ends were detected randomly along the fibril. This observation indicates that single fibrils did not span the full length of tendon but, in contrast to earlier stages of development, no fibrils shorter than 60 µm were detected [16]. As will be discussed later, these results also emphasize the importance of inter- as well as intrafibril cross-linking, since the former are essential in stabilizing the fibrils within the tendon fibre. Studies in vivo of fibrillogenesis in embryonic chick tendon have led to novel concepts of how cells may orchestrate the process of tissue assembly [5]. Using sophisticated electron microscopy techniques, the authors showed that procollagen could be processed intracellularly and that initiation of fibrillogenesis occurred in Golgi to plasma membrane carriers that were targeted to protrusions on the cell surface, designated “fibripositors”. These structures appeared to deposit their pre-assembled material onto the growing tendon and maintained the alignment by forming channels between cells parallel to the axis of the tendon. Fibripositors were not observed in collagenous matrices formed in vitro suggesting that, unless novel methods can be devised to maintain the 3-D shape of the cell, culture systems are unable to provide a full model for studying the mechanisms of fibril alignment [5]. Fibripositors were also absent in vivo postnatally suggesting that, for larger fibrils, increases in diameter occur through the accretion of extracellular collagen, as discussed previously in this section.

A. Factors Affecting Fibril Formation The kinetics of removal of the propeptides clearly influences fibril formation [17], and transient retention of the N-propeptides has been suggested as a model to account for preferred fibril diameters at intervals of about

11 nm corresponding to cleavage of the propeptide at the surface [18, 19]. Once the propeptides have been removed, however, there are several other factors which affect the nature and rate of fibril formation. 1. Telopeptides

The short, nonhelical domains at each end of the molecule, the telopeptides, which contain the cross-linking sites (see Section III), are only 15–20 amino acids in length but their removal through enzyme treatment has a dramatic effect on the kinetics of fibril formation [20]. Experiments in which the interactions between collagen molecules were assessed by X-ray diffraction and osmotic stress indicated that collagen fibrils formed after enzymatic removal of telopeptides had a native-like, banded structure, suggesting that all information necessary for fibril assembly was contained within the helical portion [21]. The conclusion from these studies was that the telopeptides performed essentially a catalytic role during fibrillogenesis [21]. Further evidence for the role of telopeptides in fibrillogenesis came from molecular modeling studies which showed that the N-telopeptides adopt an ordered structure after interaction with its helical receptor site [22]. Extension of the models to include quasi-hexagonal packing of collagen molecules showed that each N-telopeptide region can form linkages with two adjacent, aligned helical receptor regions [22], consistent with the highly ordered structure of this domain observed by X-ray diffraction [23, 24]. 2. Other Fibrillar Collagens

Fibril assembly may be modified through the association of different fibrillar collagens to form heterotypic fibrils and the formation of macromolecular alloys may provide a mechanism to produce a diverse range of fibrillar composites, consistent with their varied functional requirements [25, 26]. Collagen I forms hybrid fibrils particularly with collagens III and V, and retention of the N-propeptides of these collagens appears to constitute an additional mechanism for the control of fibril assembly, as has been shown, for example, for collagen III in skin [27]. Studies of the tissues from mice containing a mutant collagen III gene have demonstrated the crucial role of this molecule for adequate collagen type I fibrillogenesis [28]. Collagen V is known to play a central role in regulating fibril assembly in the cornea [25, 29] and, because of the existence of tissue-specific isoforms, this collagen also plays an important role in collagen matrix assembly in many other tissues [25]. Collagen type II also forms heterotypic fibrils, particularly with collagen XI, although the aggregates formed between II/XI are very different from those between collagens I/XI [26]. This observation may also be associated with the much higher content of

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glycosylated hydroxylysine residues in collagen II, which affects directly formation and morphology of the fibrils as shown by experiments with recombinant collagens [30]. Interestingly, in the cartilage matrix produced by a chondrosarcoma cell line lacking N-propeptidase activity, normal covalent cross-linking was observed between collagen II and other collagens including type XI, despite the presence primarily of unprocessed N-procollagen [31]. The importance of heterotypic fibrils in cartilage collagen assembly is discussed in more detail in Chapter 25. 3. FACIT Collagens

The fibril-associated collagens with interrupted triple helix (FACIT) are particularly important in cartilage, but this topic will be dealt with in Chapter 25. For collagen type I, interactions with the FACIT types XII and XIV have major effects in modulating the properties of soft tissues, but these collagens are essentially absent in bone. Collagen types XII and XIV are relatively large molecules with extended globular domains. There is an increasing body of evidence to indicate that these collagens mediate interactions between fibrils and are important in modulating responses to biomechanical stimuli [32–34]. 4. Other Matrix Constituents

Of the many influences from other matrix constituents on collagen fibrillogenesis, that exerted by the family of small, leucine-rich proteoglycans, which includes decorin, fibromodulin, and biglycan (see Chapter 5), has probably received the most attention. Decorin appears to play a major role in collagen fibrillogenesis, as mice with targeted decorin gene disruption were shown to exhibit skin fragility with abnormal morphology characterized by uncontrolled lateral fusion of collagen fibrils [35]. There are, however, conflicting reports on the effects of decorin on fibrillogenesis in vitro, with one model proposing that binding of archshaped decorin molecules to the collagen helix potentially limits lateral accretion of fibrils [36], whereas other studies suggested that the presence of decorin increased fibril diameter [37]. More detailed studies of decorin structure [38] may help to resolve the mechanism of these interactions.

III. CROSS-LINKING The formation of intermolecular cross-links within newly formed collagen fibrils conveys the structural stability necessary for tissue function. The main processes involved in the formation initially of intermediate, chemically reducible cross-links, followed by maturation to more stable, nonreducible bonds, have been established over several years. A number of questions remain, however,

concerning the nature of the mature forms of bonds and the mechanisms of tissue-specific cross-link formation. It is clear that lysine-derived cross-links account for almost all cross-linking in young tissue and that their formation is driven by a single enzymatic process – the action of lysyl oxidase.

A. Lysyl Oxidase Lysyl oxidase (LOX) is known to be one member of a family of enzymes including four isoforms termed lysyl oxidase-like proteins (LOXL1–4), many of which have copper-dependent amine oxidase activity [39]. For collagen, lysyl oxidases catalyze the conversion of lysine or hydroxylysine residues in the N- and C-telopeptides to aldehyde residues. For LOX, lysine tyrosyl quinone, formed by the cross-linking of two amino acid side chains within the enzyme, acts a cofactor necessary for activity [40]. Early studies showed that lysyl oxidase acts extracellularly and requires as a substrate native collagen fibrils in a quarter-staggered array [41]. This observation led to suggestions that amino acid residues at the overlap sites in the helix of adjacent molecules in the fibril participate in the reaction: these sites are known to have highly conserved amino acid sequences [42]. The presence of multiple LOX isoforms suggests an addition level of complexity for the regulation of collagen cross-linking. Studies of MC3T3-E1 osteoblastic cells indicated that LOX and LOX-like protein expression is highly regulated during differentiation, with all isoforms except LOXL2 being expressed during growth and mineralization but with different quantitative differences at each stage [43]. The results of other studies involving lysyl oxidase inhibitors in osteoblastic cells are also consistent with the view that expression of this enzyme may be an important regulator of collagen deposition [44]. LOX is synthesized as a precursor which undergoes post-translational glycosylation and cleavage of the propeptide portion by procollagen C-proteinase/BMP-1 [45]. LOXL1 is processed similarly by BMP-1 but there are no known proteolytically processed forms of LOXL2, LOXL3, or LOXL4 [39]. Expression of LOX is regulated by several cytokines including transforming growth factor-β [46]. Early studies suggested that LOX may also have important functions in tumor suppression [47]. This activity was subsequently shown to reside in the 18–20 kDa, N-propeptide portion of LOX, which is highly basic, possibly facilitating heparin sulfate-mediated re-uptake by cells to accomplish their role [48]. The N-propeptide domains are not well conserved in the LOX-like proteins and the latter are, therefore, unlikely to share tumor suppressor activity [48].

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B. Tissue Specificity of Cross-linking As shown in Figure 2, there are two major pathways of cross-linking depending on whether the residue in the telopeptide is lysine or hydroxylysine. As will become evident later in this section, the presence of hydroxylysine in the telopeptide has a profound effect on the stability of the cross-links initially formed as well as on the final products. The types of cross-link formed are related to the tissue rather than to the collagen type, and the tissue specificity is governed primarily by enzymatic hydroxylation reactions in the telopeptides. Thus, in skin collagen, virtually no hydroxylysine is present in the telopeptides so that cross-linking proceeds through the aldimine route (Fig. 2) and ultimately to histidine-containing compounds, whereas, in cartilage and bone, a large proportion of the telopeptide lysines are hydroxylated giving rise mainly to keto-imine forms of cross-link, which can lead to the formation of pyridinium and pyrrolic cross-links on maturation. Hydroxylation of lysine residues in the telopeptides clearly has crucial effects on cross-linking. Following early studies indicating that the enzyme which hydroxylates lysine residues in the telopeptides is distinct from that which acts on lysine residues destined to be in the helix [49], the enzyme designated telopeptide lysyl hydroxylase was characterized as the longer splice variant of one isotype of lysyl hydroxylase 2 (LH2) [50, 51]. Three isotypes of lysyl hydroxylase (LH1–LH3) have been described [52], but only LH2 has tissue-specific expression of alternatively

spliced forms [53]. Studies of a rare form of osteogenesis imperfecta, Bruck syndrome, indicated that the cause was a lack of telopeptide lysyl hydroxylase activity: this was initially mapped to chromosome 17 [54] but later studies of two more cases showed that the mutation was in LH2 located on chromosome 3 [55], and the earlier report probably, therefore, reflects the heterogeneity of this disorder. Studies in vitro have confirmed that alternatively spliced forms of LH2 direct the pathway of collagen cross-linking [56–58]. It is clear, therefore, that tissue differences in the patterns of collagen cross-linking are due to variations in the expression and activities of telopeptide and helical lysyl hydroxylases. Some hydroxylysine residues in central parts of fibrillar collagens destined to become the helix are further modified enzymatically during synthesis by addition of galactosyl or both galactosyl and glucosyl residues to the hydroxyl group (see Chapter 1). These hydroxylysine glycosides may participate in cross-linking giving rise to glycosylated forms of both the difunctional cross-links and their maturation products.

C. Mechanisms of Cross-linking 1. Intermediate, Difunctional Cross-links

Following the action of lysyl oxidase, the lysine aldehyde (allysine) or hydroxylysine aldehyde (hydroxyallysine) residues undergo spontaneous reactions according

Figure 2 Collagen cross-linking is tissue specific. Intracellular hydroxylation of lysine in the telopeptide gives rise to two pathways of extracellular cross-linking, the lysine aldehyde (allysine) pathway and the hydroxyallysine pathway; the latter results in mainly pyridinium and pyrrolic cross-links in bone and cartilage.

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to the separate pathways indicated in Figure 3. For allysine, reaction with a hydroxylysine or lysine residue in the helix results in the formation of the aldimines, dehydro-hydroxylysinonorleucine (∆-HLNL) or dehydrolysinonorleucine (∆-LNL), respectively. These aldimine bonds provide the main structural stability in newly formed dermal tissue, but are readily cleaved by heat or dilute acids, rendering such tissue completely soluble in dilute acetic acid. Another consequence of the chemical lability of the aldimine bonds is that reduction with borohydride is essential to stabilize the compounds to the hydrolytic processes necessary for their isolation: the use of radiolabeled borohydride was helpful initially in identifying the reduced compounds as HLNL and LNL [59]. The corresponding difunctional cross-links formed through reactions of hydroxyallysine with helical hydroxylysine or lysine residues are stabilized by a spontaneous Amadori rearrangement to produce hydroxylysino-5-ketonorleucine (HLKNL) or lysino-5-keto-norleucine (LKNL), respectively (Fig. 3). This rearrangement renders these compounds stable to the effects of mild acid treatment and peptides containing these cross-links have been isolated without borohydride reduction. After reduction, HLKNL and LKNL are converted to dihydroxy-lysinonorleucine (DHLNL) and hydroxylysinonorleucine (HLNL). The latter compound can therefore be derived from both the allysine and hydroxyallysine pathways but these can be distinguished by identifying the products of a Smith degradation (periodate cleavage followed by borohydride

Figure 3

reduction): in bone about 75% of HLNL is derived from hydroxyallysine [60]. Another difference between the lysine aldehyde and hydroxylysine aldehyde pathways of cross-linking is that the former compound can participate in an “aldol” condensation reaction to give an intramolecular crosslink between the α-chains of a single molecule. The aldol condensation product (ACP) undergoes further reactions that have been detected in tissues after reduction with borohydride (see Fig 3). Hydroxyallysine does not form intramolecular aldols and no corresponding products from this pathway have been detected. The main product derived from the aldol which was identified in borohydride-reduced tissue was the compound histidino-hydroxymerodesmosine (HHMD), involving the addition of hydroxylysine and histidine residues to the ACP [61]. Although evidence was presented that this compound was an artefact of the borohydride reduction process and that the nonreduced form did not constitute a cross-link in vivo [62], contradictory results were reported subsequently [63] and this question has still not been fully resolved. 2. Maturation of Intermediate Cross-links

The reducible, intermediate cross-links decrease in concentration during maturation and aging of the tissues [59]. Initial studies to define the way in which these bonds are modified during maturation focused on a reduction in vivo to give the same cross-links as those produced by borohydride, but these observations have been discounted: similarly, a proposed oxidative pathway of modification

Inter- and intramolecular cross-link formation occurs in different ways depending on the aldehyde involved. Telopeptide lysine aldehyde reacts with lysine or hydroxylysine residues in the helix to form aldimine (Schiff base) cross-links that give rise to LNL and HLNL on reduction with borohydride. When hydroxylysine aldehyde is present in the telopeptide, the products initially formed undergo Amadori rearrangement to form the more stable keto-imines, LKNL and HLKNL, which on reduction give HLNL and DHLNL. Lysine aldehyde also forms an intramolecular cross-link, the Aldol condensation product, which on reduction can produce either a reduced aldol or the tetrafunctional cross-link, HHMD.

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also has not been substantiated. Firm evidence has been obtained that the major maturation products are pyridinium and pyrrolic cross-links in skeletal tissues (Fig. 4) and a histidine adduct in soft tissues, although mechanisms of the maturation processes are not entirely clear, particularly with respect to kinetics and stoichiometry. In addition, immunochemical studies of mature human bone indicated that, at the C-terminal end at least, there appeared to be as yet unidentified cross-links [64]. a. Pyridinium cross-links A fluorescent, 3-hydroxypyridinium compound in hydrolysates of bovine tendon was identified by Fujimoto and colleagues [65] as a trifunctional cross-link, termed pyridinoline, and an analog, named deoxypyridinoline, was discovered in bone [66]. Because the latter cross-link involved reaction of a lysine residue from the helix, it has been termed lysyl-pyridinoline, or LP, to distinguish this compound from pyridinoline which is derived from helical hydroxylysine, and is referred to as hydroxylysylpyridinoline or HP [67]. Two mechanisms for formation of the pyridinium cross-links have been proposed: one involving interaction between two intermediate keto-imine cross-links [68] and the other a reaction between the ketoimine and hydroxylysine aldehyde [69]. The chemistry of these two mechanisms is in fact very similar but there are important functional implications as will be discussed in Section III.E. b. Pyrrolic cross-links The initial suggestion for the presence in collagen of pyrrolic cross-links came from work by Scott and

colleagues [70], based on the formation of colored reaction products of collagen hydrolysates with Ehrlich’s reagent (p-dimethylaminobenzaldehyde in acid solution). Later studies identified pyrrole-containing peptides from tendon and led to a proposed mechanism of cross-link formation [71]. This proposal suggests that in tissues where not all of the telopeptide lysine residues are hydroxylated, lysine aldehyde may interact with the intermediate keto-imine to form a pyrrolic, trifunctional cross-link. According to this mechanism, therefore, pyrrole cross-links should be absent from dermal tissue because of the lack of keto-imines and also absent from cartilage because the telopeptides are fully hydroxylated: this tissue distribution has been verified experimentally [4]. Characterization of pyrrole cross-links was difficult mainly because of their instability during the hydrolytic and chromatographic procedures necessary to isolate them. By targeting the pyrrole with novel Ehrlich’s reagent derivatives followed by mass spectral analysis, direct confirmation for the structure and location of pyrrole cross-links in bone was obtained [72]. These studies showed that, similar to the pyridinium compounds, pyrrole cross-link analogs from both helical hydroxylysyl and lysyl residues were present [72]: consistent with a previous report [73], pyrrole cross-linked peptides were derived mainly from the N-terminal end of collagen. The methods developed to date do not provide precise quantitative data for pyrrole cross-links and further studies are required to provide a comprehensive analysis of the changes in pyrrole cross-linking during development and with age.

Figure 4 Pyridinium and pyrrolic cross-links are the best characterized products of maturation. The difunctional cross-links, hydroxylysino-5-ketonorleucine (HLKNL) and lysino-5-ketonorleucine (LKNL), where R = –OH or –H, respectively, are converted to the different trifunctional cross-links depending on whether the telopeptide aldehydes are hydroxylated.

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c. Kinetics of cross-link maturation There have been relatively few studies of the kinetics of cross-link maturation and most of these have involved administration of radioactively labeled lysine to rats. From these types of study, the half-life of the reducible, difunctional cross-links has been estimated as a few weeks [74]. Several groups have shown that the concentrations of reducible, intermediate cross-links remain much higher in bone than in other tissues, including skin and cartilage. The continual turnover of bone matrix, in contrast to the other tissues, undoubtedly contributes to this observation and results in a higher proportion of recently synthesized collagen being present in bone.

D. Mineralization and Cross-linking The structural features of collagen type I fibrils in bone which allow mineralization to occur have not been fully elucidated, and conflicting reports have been published. It is generally agreed, however, that the types and location of cross-linking are different in bone compared with soft tissues containing collagen type I, such as skin or tendon. Studies of C-terminal cross-linking have suggested that mineralization itself induces structural alterations in the fibril [75], although some of the observed changes may be due to the conditions of tissue processing [76]. Because the concentrations of pyridinium cross-links in mammalian bone are relatively low throughout life in comparison with other tissues, such as cartilage [77], it has been suggested that the presence of mineral arrests maturation of difunctional cross-links into pyridinoline and deoxypyridinoline. Arguing against this contention is the fact that “aging” in vitro of mineralized bone (incubation in physiological saline at 37°C) converts precursor crosslinks to their mature forms [78]; however, earlier studies indicated that there may be differences in the rate of reaction between mineralized and nonmineralized bone [79]. The role of pyrrole cross-links in mammalian bone, although suggested to be functionally the more important mature cross-link in avian bone [4], has not yet been fully evaluated. The natural mineralization of turkey leg tendon at around 12–14 weeks of age has been widely used as a model system for the study of structural changes in collagen type I during the addition of mineral [80]. These changes appear to be strain-induced and are associated with neovascularization [81, 82]. Yamauchi and colleagues showed that the nonmineralized part of the tendon contained primarily pyridinoline cross-links whereas the mineralized portion also contained a high proportion of

deoxypyridinoline [83]. As this altered cross-linking pattern appears to involve changes in the intracellular hydroxylation of lysine residues, these results inferred that mineralization is accompanied by the production of a new matrix with modified post-translational modifications. Similar changes in lysine hydroxylation and cross-link patterns were found to be associated with changes in thermal characteristics of the collagenous matrix [76]. Studies of calcifying callus in dogs also revealed decreases in the ratio of pyridinoline to deoxypyridinoline associated with increased mineralization within the collagen fibrils [84]. These experiments serve to emphasize the fact that collagen type I capable of mineralization exhibits specific patterns of post-translational hydroxylation and fibril structure, which in turn are associated with particular cross-linking patterns, of which the presence of deoxypyridinoline is one characteristic.

E. Cross-linking in Relation to Molecular Packing A large number of studies of the three-dimensional structure of fibrillar collagen over many years culminated in establishing a quasi-hexagonal lattice model as the most appropriate to fit the available data [24]. This model not only satisfies the limitations imposed by the chemistry and location of the intermolecular cross-links [85], but also is consistent with the packing density and crytallinity changes associated with the overlap segments compared with other regions of the fibrils [86]. In conjunction with molecular modeling studies [22], the data reveal a high degree of interconnectivity between microfibrils [87], with a corrugated arrangement of fibrils cross-linked between segment 1 and 5 (Fig. 5). The difunctional, intermediate cross-links are known to stabilize the quarter-staggered overlap (between segments 1 and 5 in Fig. 5). During their maturation to trifunctional pyridinium or pyrrole cross-links, reactions can occur either with an aldehyde from another chain of the same molecule, giving an intrafibril cross-link, or with a reactive group from an adjacent microfibril, giving an interfibrillar bond linking three molecules. Distinguishing between these possibilities biochemically is difficult, but these arrangements will have major effects on the functional properties of the tissue. For mineralized tissues, electron microscopy, combined with computed tomography and three-dimensional image reconstruction methods, have provided evidence that some mineral crystals span the hole regions of adjacent microfibrils [88], a configuration that requires all microfibrils to be in register with their hole regions aligned.

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discovered as the toxic agent released in vivo after chronic ingestion of the sweet pea, Lathyrus oderatus, is now known to be an irreversible inhibitor of the cross-link initiating enzyme, lysyl oxidase, and leads to severe malformations in the skeleton as well as many other abnormalities. Following the discovery of this agent in 1959 [94], βAPN has been used extensively for both in vivo and in vitro experiments to provide noncross-linked matrix proteins for research purposes. In addition to the developmental problems associated with a severe lack of cross-linking, the quantity and nature of cross-link has profound effects on the susceptibility to proteolysis and the mechanical characteristics of collagen fibrils. 1. Susceptibility to Proteolysis Figure 5

Inter- and intrafibrillar cross-links are shown by this schematic representation of quarter-staggered collagen molecules, 4.4D periods in length, where each segment is numbered (upper). One possible arrangement of compressed microfibrils (pentafilaments denoted by the shaded areas) is shown (lower) with a corrugated arrangement of both intra- and interfibrillar cross-link between segments 1 and 5.

F. In situ Analysis of Collagen Cross-linking With improvements in instrumentation and software, Fourier transform infrared imaging (FTIRI) has found applications in the analysis of both mineral and matrix constituents of bone [89–91]. In addition to providing information on the mineral to matrix ratios and mineral crystallinity, spectral analysis can reveal specific data on the environment of different cross-link residues: whilst this information does not directly identify particular cross-links, analysis of isolated peptides with known cross-links has allowed distinctions to be made between difunctional, reducible bonds and pyridinium compounds [89]. These analyses are performed in an array so that an image of cross-link types can be built up in sections of bone. The technique not only provides a novel way to monitor maturational changes in collagen cross-linking in bone but also facilitates studies of the changes that occur in diseases and during therapy [91–93].

G. Biological Consequences of Intermolecular Collagen Cross-linking Inhibition of collagen cross-linking via the dietary lathyrogen, β-amino-proprionitrile (βAPN) provides graphic evidence of the importance of the intermolecular bonds for proper tissue function. The lathyrogen, originally

The triple helical structure of individual monomers within the collagen fibril is resistant to most proteolytic enzymes except specific collagenases or matrix metalloproteinases (see Chapter 11) which cleave through all three chains simultaneously at a point three-quarters distance from the N-terminal end. Other enzymes, such as cathepsins and elastase cleave the telopeptide portions of the molecule between the cross-linking site and the helix, thus depolymerizing the fibril. The presence of the lysinederived cross-links has, however, been shown to affect dramatically the susceptibility to proteolysis of the fibrils. Thus, for fibrils that were formed in vitro from soluble chick bone collagen, the susceptibility to proteolysis was related directly to the concentrations of difunctional crosslinks in the fibrils introduced by the actions of purified lysyl oxidase [95]. Although these studies appeared to show that as little as one cross-link per ten collagen molecules was sufficient to impart a two- to three-fold resistance to collagenase digestion, this may be an underestimate, as the cross-link values represent a figure for the whole fibril but the methods used will have introduced higher concentrations of cross-links at the surface of the fibrils. Maturation of fibrils leads to a lower susceptibility to enzyme degradation and the formation of multifunctional cross-links that link more than two molecules (see Section III.E) is crucial in giving rise to these properties. Distinct from maturational changes, the introduction of cross-links between the helices of adjacent molecules through aging processes, including advanced glycation end-product (AGE) reactions, leads to major changes in the properties of the fibrils [3]. The presence of a fibrillar network cross-linked at the telopeptides means that once cleavage of a peripheral molecule by collagenase has occurred the cleaved chains will remain attached for a period of time to the fibril through the cross-links. At body temperature, the cleaved

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fragments denature and can be detected by specific antibodies that react only with sequence-dependent epitopes, thus providing a means to assess immunocytochemically patterns of degradation in tissues [96]. Similar techniques have been used to detect the degradation patterns in cartilage [97]. 2. Mechanical Properties

There is no doubt that cross-linking in collagen is primarily responsible for the strength of the fibrils but the relative contributions of the different types of cross-link are less clear. Studies of the effects of partial lathyrism on the mechanical strength of embryonic avian bone indicated that a 10% decrease in the concentration of the difunctional keto-imine cross-link resulted in a 15% decrease in bone strength [78]. Similar studies have shown that the concentrations of pyridinoline are also related to the strength of adult bone in the rabbit [98] and rat [99]. It has been suggested that the pyrrole cross-links are more important than the pyridinium cross-links in determining the mechanical properties of bone [78], but this conclusion is based on experiments in avian bone which contain much smaller proportions of the pyridinium cross-links than mammalian bone. In bovine tendon, the concentrations of the pyrrolic and pyridinium cross-links were found to have similar positive correlations with thermal stability [100], suggesting that both types of cross-link were performing a similar function. There appear to be differences in the relative proportions of cross-links involving the N- and C-terminal telopeptides, with about 85% of the pyrroles being present at the N-terminus whereas the pyridinium cross-links were equally distributed [73]. It is possible that the N-telopeptides are involved in more intermicrofibrillar bonding in comparison to the C-terminal cross-links (see Section III.E), a factor that may have effects on biomechanical properties of the tissue. Studies of the age-related changes in bone toughness revealed very little change in the concentrations of pyridinium cross-links, but there were significant increases in pentosidine, a marker for nonenzymatic glycation, associated with decreased bone stiffness and strength [101]. These observations of the effects of in vivo glycation contrast with those for in vitro glycation by ribose that induced increased stiffness of bone together with increased fragility [102].

IV. CONCLUDING REMARKS Intracellular, post-translational modifications of collagen are recognized as key to many of the properties of the fibrillar collagens and hydroxylation of lysine residues, in particular, has far-reaching effects on cross-linking,

tissue strength and the propensity to mineralize. The application of new techniques for microscopy and spectroscopy has contributed to advances in our understanding of collagen fibril formation, mineralization and cross-link maturation. There has been relatively little consideration in this chapter of age-related changes in collagen but these may have important implications for the mechanical properties of the tissue in relation to diseases such as osteoporosis. Of the many biochemical changes that occur, attention has focused on increased hydroxylation of lysine residues and its effects on cross-linking and hydroxylysine glycoside concentrations [103]. The latter may give rise to the production of smaller collagen fibrils [30] that have modified, less robust mechanical properties consistent with the changes seen in osteoporosis. Another age-related change in collagen structure is an aspartyl to isoaspartyl transformation in both the C-terminal [104] and N-terminal [105] telopeptides. The C-telopeptide has proved useful as a biochemical marker (see Chapters 24 and 35) and alterations in the relative amounts of these modified residues have been associated with increased skeletal fragility [106]. With the realization that bone mineral density measurements have a limited ability to reflect fracture risk, much emphasis has been placed on attempting to define biochemical parameters that may provide assessments of “bone quality” in terms of fragility. The ratios of aspartyl/ isoaspartyl residues in the telopeptides have been proposed as one such parameter, as have the maturation rates of collagen cross-links detected by Fourier transform infrared microscopy. One of the challenges of future research is to fully validate these newer measures and to establish real links between the mechanical integrity of bone and biochemical characteristics of the collagen matrix.

Acknowledgments I am grateful to the Scottish Executive Environmental and Rural Affairs Department for support.

References 1. Kadler, K. E., Holmes, D. F., Trotter, J. A., and Chapman, J. A. (1996). Collagen fibril formation. Biochem. J. 316, 1–11. 2. Canty, E. G., and Kadler, K. E. (2002). Collagen fibril biosynthesis in tendon: a review and recent insights. Comp. Biochem. Physiol. A. Mol. Integr. Physiol. 133, 979–985. 3. Reiser, K., McCormick, R. J., and Rucker, R. B. (1992). Enzymatic and nonenzymatic cross-linking of collagen and elastin. FASEB J. 6, 2439–2449. 4. Knott, L., and Bailey, A. J. (1998). Collagen cross-links in mineralizing tissues: a review of their chemistry, function, and clinical relevance. Bone 22, 181–187.

Chapter 2 Fibrillogenesis and Maturation of Collagens 5. Canty, E. G., Lu, Y., Meadows, R. S., Shaw, M. K., Holmes, D. F., and Kadler, K. E. (2004). Coalignment of plasma membrane channels and protrusions (fibripositors) specifies the parallelism of tendon. J. Cell. Biol. 165, 553–563. 6. Colige, A., Sieron, A. L., Li, S. W., Schwarze, U., Petty, E., Wertelecki, W., Wilcox, W., Krakow, D., Cohn, D. H., Reardon, W., Byers, P. H., Lapiere, C. M., Prockop, D. J., and Nusgens, B. V. (1999). Human Ehlers-Danlos syndrome type VII C and bovine dermatosparaxis are caused by mutations in the procollagen I N-proteinase gene. Am. J. Hum. Genet. 65, 308–317. 7. Fernandes, R. J., Hirohata, S., Engle, J. M., Colige, A., Cohn, D. H., Eyre, D. R., and Apte, S. S. (2001). Procollagen II amino propeptide processing by ADAMTS-3. Insights on dermatosparaxis. J. Biol. Chem. 276, 31502–31509. 8. Colige, A., Vandenberghe, I., Thiry, M., Lambert, C. A., Van Beeumen, J., Li, S. W., Prockop, D. J., Lapiere, C. M., and Nusgens, B. V. (2002). Cloning and characterization of ADAMTS-14, a novel ADAMTS displaying high homology with ADAMTS-2 and ADAMTS-3. J. Biol. Chem. 277, 5756–5766. 9. Scott, I. C., Blitz, I. L., Pappano, W. N., Imamura, Y., Clark, T. G., Steiglitz, B. M., Thomas, C. L., Maas, S. A., Takahara, K., Cho, K. W., and Greenspan, D. S. (1999). Mammalian BMP-1/Tolloid-related metalloproteinases, including novel family member mammalian Tolloid-like 2, have differential enzymatic activities and distributions of expression relevant to patterning and skeletogenesis. Dev. Biol. 213, 283–300. 10. Ricard-Blum, S., Bernocco, S., Font, B., Moali, C., Eichenberger, D., Farjanel, J., Burchardt, E. R., van der Rest, M., Kessler, E., and Hulmes, D. J. (2002). Interaction properties of the procollagen C-proteinase enhancer protein shed light on the mechanism of stimulation of BMP-1. J. Biol. Chem. 277, 33864–33869. 11. Colige, A., Beschin, A., Samyn, B., Goebels, Y., Vanbeeumen, J., Nusgens, B., and Lapiere, C. (1995). Characterization and partial amino acid sequencing of a 107- kDa procollagen I N-proteinase purified by affinity chromatography on immobilized type XIV collagen. J. Biol. Chem. 270, 16724–16730. 12. Davies, D., Tuckwell, D. S., Calderwood, D. A., Weston, S. A., Takigawa, M., and Humphries, M. J. (1997). Molecular characterisation of integrin-procollagen C-propeptide interactions. Eur. J. Biochem. 246, 274–282. 13. Kadler, K. E., Hojima, Y., and Prockop, D. J. (1987). Assembly of collagen fibrils de novo by cleavage of the type I pC-collagen with procollagen C-proteinase. Assay of critical concentration demonstrates that collagen self-assembly is a classical example of an entropy-driven process. J. Biol. Chem. 262, 15696–15701. 14. Holmes, D. F. , Lowe, M. P., and Chapman, J. A. (1994). Vertebrate (chick) collagen fibrils formed in vivo can exhibit a reversal in molecular polarity. J. Mol. Biol. 235, 80–83. 15. Birk, D. E., Nurminskaya, M. V., and Zycband, E. I. (1995). Collagen fibrillogenesis in situ: fibril segments undergo post-depositional modifications resulting in linear and lateral growth during matrix development. Dev. Dyn. 202, 229–243. 16. Birk, D. E., Zycband, E. I., Woodruff, S., Winkelmann, D. A., and Trelstad, R. L. (1997). Collagen fibrillogenesis in situ: fibril segments become long fibrils as the developing tendon matures. Dev. Dyn. 208, 291–298. 17. Mellor, S. J., Atkins, G. L., and Hulmes, D. J. S. (1991). Developmental changes in the type I procollagen processing pathway in chick-embryo cornea. Biochem. J. 276, 777–784. 18. Hulmes, D. J. (1983). A possible mechanism for the regulation of collagen fibril diameter in vivo. Coll. Relat. Res. 3, 317–321. 19. Chapman, J. A. (1989). The regulation of size and form in the assembly of collagen fibrils in vivo. Biopolymers 28, 1367–1382. 20. Helseth, D. L., and Veis, A. (1981). Collagen self-assembly in vitro. Differentiating specific telopeptide-dependent interactions using selective enzyme modification and the addition of free amino telopeptide. J. Biol. Chem. 256, 7118–7128.

51 21. Kuznetsova, N., and Leikin, S. (1999). Does the triple helical domain of type I collagen encode molecular recognition and fiber assembly while telopeptides serve as catalytic domains? Effect of proteolytic cleavage on fibrillogenesis and on collagen-collagen interaction in fibers. J. Biol. Chem. 274, 36083–36088. 22. Malone, J. P., George, A., and Veis, A. (2004). Type I collagen N-telopeptides adopt an ordered structure when docked to their helix receptor during fibrillogenesis. Proteins 54, 206–215. 23. Hulmes, D. J. S., Wess, T. J., Prockop, D. J., and Fratzl, P. (1995). Radial packing, order, and disorder in collagen fibrils. Biophysi. J. 68, 1661–1670. 24. Orgel, J. P., Miller, A., Irving, T. C., Fischetti, R. F., Hammersley, A. P., and Wess, T. J. (2001). The in situ supermolecular structure of type I collagen. Structure (Camb) 9, 1061–1069. 25. Birk, D. E. (2001). Type V collagen: heterotypic type I/V collagen interactions in the regulation of fibril assembly. Micron. 32, 223–237. 26. Hansen, U., and Bruckner, P. (2003). Macromolecular specificity of collagen fibrillogenesis: fibrils of collagens I and XI contain a heterotypic alloyed core and a collagen I sheath. J. Biol. Chem. 278, 37352–37359. 27. Fleischmajer, R., MacDonald, E. D., Perlish, J. S., Burgeson, R. E., and Fisher, L. W. (1990). Dermal collagen fibrils are hybrids of type I and type III collagen molecules. J. Struct. Biol. 105, 162–169. 28. Liu, X., Wu, H., Bryne, M., Krane, S., and Jaenisch, R. (1997). Type III collagen is crucial for collagen I fibrillogenesis and for normal cardiovascular development. Proc. Natl Acad. Sci. USA 94, 1852–1856. 29. Linsenmayer, T., Gibney, E., Igoe, F., Gordon, M., Fitch, J., Fessler, L., and Birk, D. (1993). Type V collagen: molecular structure and fibrillar organization of the chicken alpha 1(V) NH2-terminal domain, a putative regulator of corneal fibrillogenesis. J. Cell. Biol. 121, 1181–1189. 30. Notbohm, H., Nokelainen, M., Myllyharju, J., Fietzek, P. P., Muller, P. K., and Kivirikko, K. I. (1999). Recombinant human type II collagens with low and high levels of hydroxylysine and its glycosylated forms show marked differences in fibrillogenesis in vitro. J. Biol. Chem. 274, 8988–8992. 31. Fernandes, R. J., Schmid, T. M., and Eyre, D. R. (2003). Assembly of collagen types II, IX and XI into nascent hetero-fibrils by a rat chondrocyte cell line. Eur. J. Biochem. 270, 3243–3250. 32. Nishiyama, T., McDonough, A. M., Bruns, R. R., and Burgeson, R. E. (1994). Type XII and XIV collagens mediate interactions between banded collagen fibers in vitro and may modulate extracellular matrix deformability. J. Biol. Chem. 269, 28193–28199. 33. Chiquet, M., Renedo, A. S., Huber, F., and Fluck, M. (2003). How do fibroblasts translate mechanical signals into changes in extracellular matrix production? Matrix Biol. 22, 73–80. 34. Jin, X., Iwasa, S., Okada, K., Ooi, A., Mitsui, K., and Mitsumata, M. (2003). Shear stress-induced collagen XII expression is associated with atherogenesis. Biochem. Biophys. Res. Commun. 308, 152–158. 35. Danielson, K. G., Baribault, H., Holmes, D. F., Graham, H., Kadler, K. E., and Iozzo, R. V. (1997). Targeted disruption of decorin leads to abnormal collagen fibril morphology and skin fragility. J. Cell Biol. 136, 729–743. 36. Weber, I. T., Harrison, R. W., and Iozzo, R. V. (1996). Model structure of decorin and implications for collagen fibrillogenesis. J. Biol. Chem. 271, 31767–31770. 37. Kuc, I. M., and Scott, P. G. (1997). Increased diameters of collagen fibrils precipitated in vitro in the presence of decorin from various connective tissues. Connect. Tissue. Res. 36, 287–296. 38. Scott, P. G., McEwan, P. A., Dodd, C. M., Bergmann, E. M., Bishop, P. N., and Bella, J. (2004). Crystal structure of the dimeric protein core of decorin, the archetypal small leucine-rich repeat proteoglycan. Proc. Natl Acad. Sci. USA 101, 15633–15638. 39. Molnar, J., Fong, K. S., He, Q. P., Hayashi, K., Kim, Y., Fong, S. F., Fogelgren, B., Szauter, K. M., Mink, M., and Csiszar, K. (2003). Structural and functional diversity of lysyl oxidase and the LOX-like proteins. Biochim. Biophys. Acta. 1647, 220–224.

52 40. Wang, S. X., Mure, M., Medzihradszky, K. F., Burlingame, A. L., Brown, D. E., Dooley, D. M., Smith, A. J., Kagan, H. M., and Klinman, J. P. (1996). A crosslinked cofactor in lysyl oxidase: redox function for amino acid side chains. Science 273, 1078–1084. 41. Siegel, R. C. (1979). Lysyl oxidase. Int. Rev. Connect. Tissue Res. 8, 73–118. 42. Fietzek, P. P., Allmann, H., Rauterberg, J., and Wachter, E. (1977). Ordering of cyanogen bromide peptides of type III collagen based on their homology to type I collagen: preservation of sites for crosslink formation during evolution. Proc. Natl Acad. Sci. USA 74, 84–86. 43. Atsawasuwan, P., Mochida, Y., Parisuthiman, D., and Yamauchi, M. (2005). Expression of lysyl oxidase isoforms in MC3T3-E1 osteoblastic cells. Biochem. Biophys. Res. Commun. 327, 1042–1046. 44. Hong, H. H., Pischon, N., Santana, R. B., Palamakumbura, A. H., Chase, H. B., Gantz, D., Guo, Y., Uzel, M. I., Ma, D., and Trackman, P. C. (2004). A role for lysyl oxidase regulation in the control of normal collagen deposition in differentiating osteoblast cultures. J. Cell. Physiol. 200, 53–62. 45. Trackman, P., Bedell, H. D., Tang, J., and Kagan, H. (1992). Posttranslational glycosylation and proteolytic processing of a lysyl oxidase precursor. J. Biol. Chem. 267, 8666–8671. 46. Boak, A. M., Roy, R., Berk, J., Taylor, L., Polgar, P., Goldstein, R. H., and Kagan, H. M. (1994). Regulation of lysyl oxidase expression in lung fibroblasts by transforming growth factor-beta 1 and prostaglandin E2. Am. J. Respir. Cell. Mol. Biol. 11, 751–755. 47. Kenyon, K., Contente, S., Trackman, P., Tang, J., Kagan, H., and Friedman, R. (1991). Lysyl Oxidase and rrg Messenger RNA. Science 253, 802. 48. Palamakumbura, A. H., Jeay, S., Guo, Y., Pischon, N., Sommer, P., Sonenshein, G. E., and Trackman, P. C. (2004). The propeptide domain of lysyl oxidase induces phenotypic reversion of ras-transformed cells. J. Biol. Chem. 279, 40593–40600. 49. Royce, P. M., and Barnes, M. J. (1985). Failure of highly purified lysyl hydroxylase to hydroxylate lysyl residues in the non-helical regions of collagen. Biochem. J. 230, 475–480. 50. Mercer, D. K., Nicol, P. F., Kimbembe, C., and Robins, S. P. (2003). Identification, expression, and tissue distribution of the three rat lysyl hydroxylase isoforms. Biochem. Biophys. Res. Commun. 307, 803–809. 51. van der Slot, A. J., Zuurmond, A. M., Bardoel, A. F., Wijmenga, C., Pruijs, H. E., Sillence, D. O., Brinckmann, J., Abraham, D. J., Black, C. M., Verzijl, N., De Groot, J., Hanemaaijer, R., Tekoppele, J. M., Huizinga, T. W., and Bank, R. A. (2003). Identification of PLOD2 as telopeptide lysyl hydroxylase, an important enzyme in fibrosis. J. Biol. Chem. 278, 40967–40972. 52. Valtavaara, M., Szpirer, C., Szpirer, J., and Myllyla, R. (1998). Primary structure, tissue distribution, and chromosomal localization of a novel isoform of lysyl hydroxylase (lysyl hydroxylase 3). J. Biol. Chem. 273, 12881–12886. 53. Yeowell, H. N., and Walker, L. C. (1999). Tissue specificity of a new splice form of the human lysyl hydroxylase 2 gene. Matrix Biol. 18, 179–187. 54. Bank, R. A., Robins, S. P., Wijmenga, C., Breslau-Siderius, L. J., Bardoel, A. F., van der Sluijs, H. A., Pruijs, H. E., and TeKoppele, J. M. (1999). Defective collagen crosslinking in bone, but not in ligament or cartilage, in Bruck syndrome: indications for a bone-specific telopeptide lysyl hydroxylase on chromosome 17. Proc. Natl Acad. Sci. USA 96, 1054–1058. 55. Ha-Vinh, R., Alanay, Y., Bank, R. A., Campos-Xavier, A. B., Zankl, A., Superti-Furga, A., and Bonafe, L. (2004). Phenotypic and molecular characterization of Bruck syndrome (osteogenesis imperfecta with contractures of the large joints) caused by a recessive mutation in PLOD2. Am. J. Med. Genet. A 131A, 115–120. 56. Pornprasertsuk, S., Duarte, W. R., Mochida, Y., and Yamauchi, M. (2004). Lysyl hydroxylase-2b directs collagen cross-linking pathways in MC3T3-E1 cells. J. Bone Miner. Res. 19, 1349–1355.

Simon P. Robins 57. Walker, L. C., Overstreet, M. A., and Yeowell, H. N. (2005). Tissuespecific expression and regulation of the alternatively-spliced forms of lysyl hydroxylase 2 (LH2) in human kidney cells and skin fibroblasts. Matrix Biol. 23, 515–523. 58. Pornprasertsuk, S., Duarte, W. R., Mochida, Y., and Yamauchi, M. (2005). Overexpression of lysyl hydroxylase-2b leads to defective collagen fibrillogenesis and matrix mineralization. J. Bone Miner. Res. 20, 81–87. 59. Bailey, A. J., Robins, S. P., and Balian, G. (1974). Biological significance of the intermolecular crosslinks of collagen. Nature 251, 105–109. 60. Robins, S. P., and Bailey, A. J. (1975). The mechanism of stabilization of the reducible intermediate crosslinks. Biochem. J. 149, 381–385. 61. Tanzer, M. L., Housley, T., Berube, L., Fairweather, R., Franzblau, C., and Gallop, P. M. (1973). Structure of two histidine-containing crosslinks from collagen. J. Biol. Chem. 248, 393–402. 62. Robins, S. P., and Bailey, A. J. (1973). The characterisation of Fraction C, a possible artifact produced during the reduction of collagen fibres with borohydride. Biochem. J. 135, 657–665. 63. Bernstein, P. H., and Mechanic, G. L. (1980). A natural histidine-based imminium cross-link in collagen and its location. J. Biol. Chem. 255, 10414–10422. 64. Eriksen, H. A., Sharp, C. A., Robins, S. P., Sassi, M. L., Risteli, L., and Risteli, J. (2004). Differently cross-linked and uncross-linked carboxyterminal telopeptides of type I collagen in human mineralised bone. Bone 34, 720–727. 65. Fujimoto, D., Moriguchi, T., Ishida, T., and Hayashi, H. (1978). The structure of pyridinoline, a collagen crosslink. Biochem. Biophys. Res. Commun. 84, 52–57. 66. Ogawa, T., Ono, T., Tsuda, M., and Kawanashi, Y. (1982). A novel fluor in insoluble collagen: a crosslinking molecule in collagen molecule. Biochem. Biophys. Res. Commun. 107, 1252–1257. 67. Eyre, D. R. (1984). Cross-linking in collagen and elastin. Ann. Rev. Biochem. 53, 717–748. 68. Eyre, D. R., and Oguchi, H. (1980). Hydroxypyridinium crosslinks of skeletal collagens: their measurement, properties and a proposed pathway of formation. Biochem. Biophys. Res. Commun. 92, 403–410. 69. Robins, S. P. (1983). Crosslinking of collagen: isolation, structural characterization and glycosylation of pyridinoline. Biochem. J. 215, 167–173. 70. Scott, J. E., Hughes, E. W., and Shuttleworth, A. (1981). A collagenassociated Ehrlich chromogen: a pyrrolic cross-link? Biosci. Rep. 1, 611–618. 71. Kuypers, R., Tyler, M., Kurth, L. B., Jenkins, I. D., and Horgan, D. J. (1992). Identification of the loci of the collagen-associated Ehrlich chromogen in type I collagen confirms its role as a trivalent cross-link. Biochem. J. 283, 129–136. 72. Brady, J. D., and Robins, S. P. (2001). Structural characterization of pyrrolic cross-links in collagen using a biotinylated Ehrlich’s reagent. J. Biol. Chem. 276, 18812–18818. 73. Hanson, D., and Eyre, D. (1996). Molecular site specificity of pyridinoline and pyrrole cross- links in type I collagen of human bone. J. Biol. Chem. 271, 26508–26516. 74. Reiser, K. M., and Last, J. A. (1986). Biosynthesis of collagen crosslinks: in vivo labelling of neonatal skin, tendon, and bone in rats. Connect. Tissue Res. 14, 293–306. 75. Otsubo, K., Katz, E. P., Mechanic, G. L., and Yamauchi, M. (1992). Cross-linking connectivity in bone collagen fibrils: the COOH-terminal locus of free aldehyde. Biochemistry (Mosc) 31, 396–402. 76. Knott, L., Tarlton, J. F., and Bailey, A. J. (1997). Chemistry of collagen cross-linking: biochemical changes in collagen during the partial mineralization of turkey leg tendon. Biochem. J. 322, 535–542. 77. Eyre, D. R., Dickson, I. R., and VanNess, K. P. (1988). Collagen crosslinking in human bone and cartilage: age-related changes in the content of mature hydroxypyridinium residues. Biochem. J. 252, 495–500.

Chapter 2 Fibrillogenesis and Maturation of Collagens 78. Knott, L., Whitehead, C. C., Fleming, R. H., and Bailey, A. J. (1995). Biochemical changes in the collagenous matrix of osteoporotic avian bone. Biochem. J. 310, 1045–1051. 79. Eyre, D. R. (1981). Crosslink maturation in bone collagen. Dev. Biochem. 22, 51–55. 80. Traub, W., Arad, T., and Weiner, S. (1989). Three-dimensional ordered distribution of crystals in turkey tendon collagen fibers. Proc. Natl Acad. Sci. USA 86, 9822–9826. 81. Landis, W. J., and Silver, F. H. (2002). The structure and function of normally mineralizing avian tendons. Comp. Biochem. Physiol. A Mol. Integr. Physiol. 133, 1135–1157. 82. Landis, W. J., Kraus, B. L., and Kirker-Head, C. A. (2002). Vascularmineral spatial correlation in the calcifying turkey leg tendon. Connect. Tissue Res. 43, 595–605. 83. Yamauchi, M., and Katz, E. (1993). The post-translational chemistry and molecular packing of mineralizing tendon collagens. Connect. Tissue Res. 29, 81–98. 84. Wassen, M. H., Lammens, J., Tekoppele, J. M., Sakkers, R. J., Liu, Z., Verbout, A. J., and Bank, R. A. (2000). Collagen structure regulates fibril mineralization in osteogenesis as revealed by cross-link patterns in calcifying callus. J. Bone Miner. Res. 15, 1776–1785. 85. Bailey, A. J., Light, N. D., and Atkins, E. D. (1980). Chemical crosslinking restrictions on models for the molecular organization of the collagen fibre. Nature 288, 408–410. 86. Hulmes, D. J., Holmes, D. F., and Cummings, C. (1985). Crystalline regions in collagen fibrils. J. Mol. Biol. 184, 473–477. 87. Wess, T. J. (2005). Collagen fibril form and function. Adv. Prot. Chem. 70, 341–374. 88. Landis, W., Hodgens, K., Song, M., Arena, J., Kiyonaga, S., Marko, M., Owen, C., and McEwen, B. (1996). Mineralization of collagen may occur on fibril surfaces: Evidence from conventional and high-voltage electron microscopy and three-dimensional imaging. J. Struct. Biol. 117, 24–35. 89. Paschalis, E. P., Verdelis, K., Doty, S. B., Boskey, A. L., Mendelsohn, R., and Yamauchi, M. (2001). Spectroscopic characterization of collagen cross-links in bone. J. Bone Miner. Res. 16, 1821–1828. 90. Blank, R. D., Baldini, T. H., Kaufman, M., Bailey, S., Gupta, R., Yershov, Y., Boskey, A. L., Coppersmith, S. N., Demant, P., and Paschalis, E. P. (2003). Spectroscopically determined collagen Pyr/deH-DHLNL cross-link ratio and crystallinity indices differ markedly in recombinant congenic mice with divergent calculated bone tissue strength. Connect. Tissue Res. 44, 134–142. 91. Huang, R. Y., Miller, L. M., Carlson, C. S., and Chance, M. R. (2003). In situ chemistry of osteoporosis revealed by synchrotron infrared microspectroscopy. Bone 33, 514–521. 92. Paschalis, E. P., Recker, R., DiCarlo, E., Doty, S. B., Atti, E., and Boskey, A. L. (2003). Distribution of collagen cross-links in normal human trabecular bone. J. Bone Miner. Res. 18, 1942–1946. 93. Paschalis, E. P., Shane, E., Lyritis, G., Skarantavos, G., Mendelsohn, R., and Boskey, A. L. (2004). Bone fragility and collagen cross-links. J. Bone Miner. Res. 19, 2000–2004.

53 94. Levene, C. I., and Gross, J. (1959). Alterations in state of molecular aggregation of collagen induced in chick embryos by beta-aminopropionitrile (lathyrus factor). J. Exp. Med. 110, 771–790. 95. Vater, C. A., Harris, E. D. J., and Siegel, R. C. (1979). Native crosslinks in collagen fibrils induce resistance to human synovial collagenase. Biochem. J. 181, 639–645. 96. Rucklidge, G. J., Riddoch, G. I., and Robins, S. P. (1986). Immunocytochemical staining of rat tissues with antibodies to denatured type I collagen: a technique for localizing areas of collagen degradation. Collagen Rel. Res. 6, 185–193. 97. Dodge, G. R., and Poole, A. (1989). Immunohistochemical detection and immunochemical analysis of type II collagen degradation in human normal, rheumatoid, and osteoarthritic articular cartilages and in explants of bovine articular cartilage cultured with interleukin 1. J. Clin. Investi. 83, 647–661. 98. Lees, S., Eyre, D. R., and Barnard, S. M. (1990). BAPN dose dependence of mature crosslinking in bone matrix collagen of rabbit compact bone: corresponding sonic velocity and equatorial diffraction spacing. Connect. Tissue Res. 24, 95–105. 99. Oxlund, H., Barckman, M., Ortoft, G., and Andreassen, T. T. (1995). Reduced concentrations of collagen cross-links are associated with reduced strength of bone. Bone 17, 365S–371S. 100. Horgan, D. J., King, N. L., Kurth, L. B., and Kuypers, R. (1990). Collagen crosslinks and their relationship to the thermal properties of calf tendons. Arch. Biochem. Biophys. 281, 21–26. 101. Wang, X., Shen, X., Li, X., and Agrawal, C. M. (2002). Age-related changes in the collagen network and toughness of bone. Bone 31, 1–7. 102. Vashishth, D., Gibson, G. J., Khoury, J. I., Schaffler, M. B., Kimura, J., and Fyhrie, D. P. (2001). Influence of nonenzymatic glycation on biomechanical properties of cortical bone. Bone 28, 195–201. 103. Bätge, B., Winter, C., Notbohm, H., Acil, Y., Brinckmann, J., and Muller, P. K. (1997). Glycosylation of human bone collagen in relation to lysylhydroxylation and fibril diameter. J. Biochem. (Tokyo) 122, 109–115. 104. Fledelius, C., Johnsen, A. H., Cloos, P. A. C., Bonde, M., and Qvist, P. (1997). Characterization of urinary degradation products derived from type I collagen. Identification of a beta-isomerized Asp-Gly sequence within the C-terminal telopeptide (α1) region. J. Biol. Chem. 272, 9755–9763. 105. Brady, J. D., Ju, J., and Robins, S. P. (1999). Isoaspartyl bond formation within n-terminal sequences of collagen type I: implications for their use as markers of collagen degradation. Clin. Sci. 96, 209–215. 106. Garnero, P., Cloos, P., Sornay-Rendu, E., Qvist, P., and Delmas, P. D. (2002). Type I collagen racemization and isomerization and the risk of fracture in postmenopausal women: the OFELY prospective study. J. Bone Miner. Res. 17, 826–833.

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Chapter 3

Vitamin K Dependent Proteins of Bone and Cartilage Caren M. Gundberg Ph.D.

Department of Orthopaedics and Rehabilitation, Yale University School of Medicine, New Haven, Connecticut 06510

Satoru K. Nishimoto Ph.D.

Department of Molecular Sciences, The University of Tennessee College of Medicine, Memphis, Tennesse 38163

V. Gas6 VI. Vitamin K/Warfarin References

I. Abstract II. Introduction III. Osteocalcin IV. Matrix Gla Protein

I. ABSTRACT

and cellular functions. The formation of gammacarboxyglutamic acid (Gla) occurs via a unique posttranslational modification of specific peptide-bound glutamate residues in selective proteins. This reaction, which is required for the biological activities of all the vitamin K-dependent proteins, is inhibited by warfarin (Fig. 1). Beside the vitamin K-dependent coagulation factors, prothrombin, and factors VII, IX, X, Gla-containing proteins include gamma-carboxylase, Protein C and Protein S anticoagulation factors and Gas6 which is involved in cell proliferation. Gla-containing proteins of unknown function include Protein Z, a plasma glycoprotein, and Proline Rich Gla Protein 1, Proline Rich Gla Protein 2, Transmembrane Gla Protein 3, and Transmembrane Gla Protein 4 [1–5]. The last four proteins were identified as transmembrane Gla Proteins and are widely expressed in many tissue types [6, 7]. In addition, either carboxylase activity or vitamin K-dependent formation of Gla residues have been observed in a wide variety of other tissues, including bone, kidney, intestine, placenta, pancreas, skin,

The importance of vitamin K for the normal functioning of blood coagulation factors is well known. However, this vitamin is also involved in the physiological activation of various proteins that are not involved in hemostasis. Here we focus on the biochemistry and physiology of osteocalcin and Matrix Gla Protein (MGP), which are abundant in bone and cartilage, respectively. While osteocalcin biosynthesis is restricted to bone and dentin, MGP is synthesized in a variety of tissues and cell types. Two other vitamin K-dependent proteins, Gas6, which is involved in the regulation of cell proliferation and Protein S, an inhibitor of coagulation, have been shown to have a potential role in bone metabolism.

II. INTRODUCTION The family of vitamin K-dependent gammacarboxylated proteins are important in a variety of tissue Dynamics of Bone and Cartilage Metabolism

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Copyright © 2006 by Academic Press. All rights of reproduction in any form reserved.

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Figure 1 Gamma-carboxyglutamic acid (Gla). Vitamin K and CO2 are required for the carboxylation of specific glutamic acid residues in osteocalcin. This reaction is inhibited by warfarin. The adjacent carboxyl groups of Gla are binding sites for Ca2+.

spleen, lung, heart, blood vessels, spinal cord, and testis. Accumulation of Gla-containing proteins in sites of pathological ectopic calcification have been observed [2–4]. Bone and cartilage Gla proteins are osteocalcin and Matrix Gla Protein (MGP), which are small extracellular Ca2+-binding proteins. Osteocalcin is exclusive to bone and dentin. MGP is found in highest concentrations in calcified cartilage, but MGP occurs in a broad variety of tissues [8–10]. MGP plays an important role in regulating cartilage and vascular calcification [11–14]. Osteocalcin and MGP have been grouped in the same gene family because they share significant sequence similarity. In addition two other vitamin K-dependent proteins, Gas6 and Protein S, have been identified in bone [15, 16].

III. OSTEOCALCIN A. Structure and Biosynthesis Osteocalcin is a small protein (49 amino acids) synthesized by mature osteoblasts, odontoblasts, and hypertrophic chondrocytes. Osteocalcin is characterized by the presence of three residues of Gla and in all species the Gla-containing portion of the molecule is strongly

conserved [17, 18]. In human bone, however, the first potential Gla residue at position 17 is only partially carboxylated (55–89%) while residues 21 and 24 are greater than 90% gamma-carboxylated, likely due to partial carboxylation of the protein during synthesis [19]. In all species, the protein is synthesized as an 11-kDa molecule consisting of a 23 residue hydrophobic signal peptide, a 26 residue propeptide, and the 49 residue mature protein. After the hydrophobic region is cleaved by a signal peptidase, pro-osteocalcin is gamma-carboxylated. The pro-region contains a gamma-carboxylation recognition site homologous to corresponding regions in the vitamin K-dependent clotting factors (Fig. 2). Arg at position -1 and Phe at -16 are strictly conserved and appear to be critical for the binding of the vitamin K-dependent carboxylase enzyme to its substrates [19–22]. However, recent studies show that the carboxylase has a 2 × 105-fold weaker affinity for the osteocalcin propeptide compared to most of the other propeptide factors and an attached propeptide is not required for carboxylation of osteocalcin. Rather, osteocalcin contains a high-affinity carboxylase recognition site within its primary sequence [23]. After carboxylation, the propeptide is removed and the mature protein is secreted [24]. Combined chemical, immunochemical, and spectral investigations have provided a model of osteocalcin structure [25]. A detailed three-dimensional structure for osteocalcin has been determined by 1H 2D NMR and X-ray crystallography (Fig. 3). Apo-osteocalcin is relatively unstructured except for a disulfide bond between Cys23–Cys29. Calcium addition induces a folded structure containing several helical regions, a C-terminal hydrophobic core and an unstructured N-terminus from residues 1–15. The structured region spans residues 16–49 and contains three regions of helical secondary structure. All three Gla residues are found in the first helical region and a disulfide bond between Cys23 and Cys29 separates this and the second helical region (residues 27–35).

Figure 2 Comparison of the primary sequences of human osteocalcin (bottom) and Human MGP (top). Bold symbols indicate conserved regions; E’ indicate γ-carboxyglutamic acid; italics indicate phosphoserines. Processing of the 84 amino acid protein yields the 1–77 form isolated from human bone. A γ-carboxylation recognition site is homologous to corresponding regions in the vitamin K-dependent clotting factors. Arg (at position −1 in OC and +30 in MGP) and Phe (at −16 in OC and +15 in MGP) are strictly conserved and appear to be critical for the binding of the vitamin K-dependent carboxylase enzyme to its substrates. The ala-gly substitution at −10 osteocalcin results in reduced affinity for the vitamin K-carboxylase enzyme.

Chapter 3 Vitamin K Dependent Proteins of Bone and Cartilage

Figure 3 Native osteocalcin contain several helical regions, a C-terminal hydrophobic core and an unstructured N-terminus from residues 1–15. The structured region spans residues 16–49 and contains three regions of helical secondary structure. All three Gla residues are found in the first helical region and a disulfide bond between Cys23 and Cys29 separates this and the second helical region (residues 27–35). Residues 41–44 comprise the third helical region. Hydrogen bonds within these three regions serve to stabilize the helical regions forming a tight globular structure.

Residues 41–44 comprise the third helical region. Hydrogen bonds within these three regions serve to stabilize the helical regions forming a tight globular structure. The Gla residues which coordinate calcium are on the same face of the helical region and are surface exposed. These can interact with the intercalcium spacings in the hydroxyapatite lattice. The C-terminus extends outward and would be accessible to neighboring cells as well as endogenous proteinases [26, 27]. This structure is consistent with many reports of osteocalcin C-terminal peptides having chemotactic activity to osteoclast precursor cells [28–30]. Likewise, consistent with an unstructured N-terminus, the serine proteases, cathepsin D, L, and H degrade osteocalcin at the 7–8 bond [31].

B. Function Although osteocalcin is one of the most abundant noncollagenous proteins in bone, the biological function of osteocalcin has not been precisely defined. Because of its specific interaction with hydroxyapatite, osteocalcin was thought to affect the growth or maturation of Ca2+-phosphate mineral phases. Osteocalcin first appears coincident with the onset of mineralization and an increase in synthesis of the protein occurs in concert with hydroxyapatite deposition during skeletal growth [32, 33]. Immunolocalization studies show that the protein is distributed throughout the mineralized regions of bone matrix, dentin, and calcified cartilage [34–36] . However, an accumulation of evidence indicates that osteocalcin is not related to events which allow mineral deposition to occur but rather that it participates in the regulation of mineralization or bone turnover. Early studies showed that the

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protein could regulate growth of hydroxyapatite crystals in solution [37]. In vitro studies demonstrate that osteocalcin is a marker of late osteoblast differentiation [38–40]. When osteoblasts are grown in culture they produce a dense collagenous extracellular matrix [41, 42]. Alkaline phosphatase activity and MGP expression increase post-proliferatively coincident with the onset of mineralization. As the extracellular matrix accumulates and mineralizes, synthesis of osteocalcin and other calcium-binding proteins, e.g. osteopontin and bone sialoprotein, is initiated. Alkaline phosphatase activity subsequently decreases, but osteocalcin synthesis remains high throughout the life of the culture [43, 44]. In cultures in which mineralization is delayed, expression of osteocalcin is also delayed [38]. Osteocalcin-null mice are characterized by a progressive increase in bone mass, leading to bone of better biomechanical quality by 6–9 months. These mice exhibit an accelerated rate of bone formation without changes in osteoclast or osteoblast number [45]. The gradual appearance of this phenotype is consistent with the fact that osteocalcin content of bone is low in newborn rats (less than 4% of the adult level) and increases gradually with age [46]. No changes in mineral content of the bones of osteocalcindepleted mice were detectable by von Kossa staining or histomorphometry. However, a more sensitive assay of mineralization, Fourier transform infrared microspectroscopy, revealed differences in the size and perfection of the crystallite [47]. In wild-type animals the crystals were larger and more perfect in the cortical bone than in trabecular bone. In contrast, in the osteocalcin knockout animals, the crystal size and perfection were the same in both the trabecular and cortical bone and resembled that of the wild-type trabecular bone. These findings are consistent with impaired mineral maturation in the osteocalcin-deficient bone and implies the presence of newer (less remodeled) mineral. Similar findings were observed when ovariectomized wild-type and knockout mice were compared. Based on these findings, it appears that osteocalcin plays some as yet undefined role in regulating bone mineral turnover. Other in vivo and in vitro studies support a role for osteocalcin in bone remodeling. First, disrupted collagen fibrillogenesis in the cloned mouse calvarial cell line MCT3T3-E1 results in increased turnover of the collagenous matrix, a decrease in alkaline phosphatase but a fivefold increase in osteocalcin biosynthesis [48]. Second, the pattern of osteocalcin distribution in human osteons changes with gender and age, and localized reductions of osteocalcin in the extracellular matrix are associated with reduced cortical remodeling [49]. Several earlier studies have suggested that osteocalcin is involved in

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recruitment and activation of bone resorbing cells. The protein is a chemoattractant for peripheral mononuclear cells and osteoclast-like cells of giant cell tumors [28–30]. Incorporation of osteocalcin into synthetic hydroxyapatite particles placed onto chick chorioallantoic membrane enhanced osteoclast formation [50]. Subcutaneously implanted bone particles which were osteocalcin-deficient show a decrease in both progenitor cell recruitment and resorption of the bone particles [51]. It is possible that osteocalcin may act in combination with other hydroxyapatite binding proteins as a signal for bone remodeling. Osteopontin, which potentiates osteoclast adhesion to mineral surfaces, forms a complex with osteocalcin in vitro [52, 53]. Altered expression of both osteocalcin and osteopontin have been reported in rat models of osteopetrosis in which osteoclastic resorption is impaired. Further studies in animals depleted of combinations of matrix proteins should provide clues to the function of osteocalcin and other bone-specific proteins.

C. Gene Structure and Regulation The human osteocalcin gene is a single-copy gene located at the distal long arm of chromosome 1 [54]. The structure and sequence of the gene in various species are similar [55] but multiple copies of the gene occur in the rat and mouse [56, 57]. In the rat, one or more copies of the gene have been found, depending upon the strain. The mouse osteocalcin cluster contains three genes, two of which encode osteocalcin and are expressed only in bone (OG1 and OG2), while a third gene, the osteocalcinrelated gene (ORG) is found in low levels in brain, lung, and kidney, but not in bone. ORG is expressed at day 10 in embryonic development, earlier than OG1 and OG2, which are expressed at day 17, the beginning of osteogenesis. The coding sequence of ORG carries five amino acid substitutions, one at the propeptide cleavage site. ORG also contains an additional exon that is not translated and a 3-kb insertion separates the ORG coding sequence from a bone-specific promoter. The insertion sequence has the structure of a typical retrovirus, an attribute which leads to the down-regulation of transcription, possibly explaining the low levels of expression of ORG in nonosseous tissues [57]. Analysis of the OG2 promoter revealed that a 147-bp fragment contains two osteoblast cis-acting elements, OSE1 and OSE2. The OSE2-binding protein is Runx2, a master regulator of osteoblast differentiation [58]. The OSE1-binding protein is ATF4, also known as cAMPresponse element protein 2, which belongs to the subfamily of cAMP-response element-binding protein/ATF

basic leucine zipper proteins [59]. ATF4, like Runx2 and Osterix, another known osteoblast-specific transcription factor, can induce osteoblast-specific gene expression in nonosteoblastic cells and all three appear to be important factors involved in the regulation of temporal expression of osteoblastic genes [60]. Osteocalcin expression is regulated by various hormones and growth factors [reviewed in reference 61]. A specific vitamin D regulatory element has been identified in the osteocalcin promoter [62–65]. In most species 1,25(OH)2D up-regulates osteocalcin biosynthesis; however, in the mouse osteocalcin synthesis is suppressed by 1,25(OH)2D [66, 67]. Stimulation of osteocalcin by 1,25(OH)2D in chicks is increased when cultures are derived from 12-dayold embryos, but decreased in 17-day-old embryos [68]. There is a significant nucleotide difference in the VDRE of the mouse gene and the lack of inducibility of mouse osteocalcin is likely attributable to the inability of the VDR to bind and transactivate the mouse promoter [69]. This vitamin D response element is also the mediator of positive regulation in response to retinoic acid [70]. Separate promoter regions are responsible for repression by glucocorticoid and stimulation by TGFβ. On the other hand, cAMP influences osteocalcin gene expression by modulating mRNA stability [71]. Some effectors, e.g. γ-interferon, IGF I or II, TNFα, and some metal ions indirectly regulate osteocalcin through vitamin D responsiveness [72]. Osteocalcin mRNA levels are also altered by estrogen, thyroxine, calcitonin, prostaglandin E2, or BMPs, but the specific mechanisms have not been defined and may be indirect [61]. Osteocalcin mRNA has also been detected in bone marrow megakaryocytes and peripheral blood platelets, liver, lung, and brain by reverse transcription-polymerase chain reaction but translation into protein has not been verified [73, 74]. Like the mouse ORG gene, the mRNA levels observed in these tissues are three orders of magnitude lower than found in bone. Furthermore, they are not vitamin D-responsive, suggesting that control of expression is under a nonskeletal promoter. To date, bone and dentine are the only known sites of production of osteocalcin.

D. Catabolism While osteocalcin is primarily deposited in the extracellular matrix of bone, a small amount enters the blood. The circulating levels of osteocalcin are taken as a specific marker of osteoblastic activity [75; see also Chapter 34]. The catabolic fate of osteocalcin is of interest because of the use of circulating osteocalcin as a marker of bone formation in metabolic bone disease. Various fragments

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Chapter 3 Vitamin K Dependent Proteins of Bone and Cartilage

of osteocalcin are known to circulate and these have been detected by antibodies originally made against the intact molecule. It is thought that the majority of circulating osteocalcin comprises the intact molecule and a large N-terminal mid-molecule fragment [76, 77]. Although the large N-terminal mid-molecule fragment has not been directly sequenced, it is thought to encompass residues 1–43. This is based on the fact that osteocalcin contains a tryptic site at residue 43, and the two antibodies used to characterize this fragment are specific for residues 5–13 and for the 25–37 region. It has been suggested that the large N-terminal midmolecule fragment is generated by proteolysis in the circulation or during sample processing and storage. This is primarily based on the findings that: (1) the fragment is detected immediately after blood sampling; and (2) osteocalcin levels decrease with incubation at room temperature when measured by conventional RIA or by intact assays, but values are stable with an assay that recognizes both the intact and large N-terminal mid-molecule fragment [76, 78]. However, the large N-terminal mid-molecule fragment was also detected in conditioned media from human osteoblast-like cells [76]. Whether this fragment was derived from proteolysis in the media or by intracellular processing has not been established. Studies by Taylor et al. [79] suggest that a unique fragment found in sera from patients with Paget’s disease may have been derived from altered osteoblastic synthesis. It is possible that osteoblastic degradation of osteocalcin may serve as a mechanism to regulate its own concentration. Recent data indicate that there are in fact other smaller (<30 residues) immunoreactive species of osteocalcin in the serum and urine in addition to the larger mid molecule fragment [80, 81]. In urine, fragments encompassing residues 14–28 and 6–31 have been isolated and identified by mass spectrometry and N-terminal sequencing [82–84]. The breakdown of osteocalcin may also occur during osteoclastic dissolution of bone. Both intact osteocalcin and mid-molecular fragments were liberated from resorption lacunae when osteoclasts were cultured on bovine bone and the concentration of osteocalcin (and fragments) in the culture media was highly correlated with a marker of bone resorption [85, 86]. In two recent studies, measurements of urinary mid-molecule fragments showed a higher correlation with resorption markers than with formation markers and were predictive of vertebral fractures [82, 87]. It is likely that these small fragments are the product of osteoclastic enzymes [88]. Cathepsins D, L, and H will degrade human osteocalcin to the 1–41, 4–43, 8–43, 4–49, and 8–49 fragments, and these can be recognized by many polyclonal antibodies [89]. Cathepsin K degrades

osteocalcin to the 8–33 fragment (unpublished observations). In addition, plasmin cleaves osteocalcin to the 1–43 and 44–49 peptides both when the protein is in solution and when it is bound to hydroxyapatite, resulting in the detachment of the peptides from the crystal surface [90]. This protease, which is associated with membranes of osteoblast-like cells, has a potential role in bone resorption by activating collagenase. Whether the generation of osteocalcin fragments during bone resorption is related to its potential biological function in regulating bone turnover is unknown.

IV. MATRIX Gla PROTEIN A. Structure and Biosynthesis MGP was the second vitamin K-dependent protein to be discovered in bone. MGP is likely to be the ancestral template for osteocalcin, as sharks contain MGP but not osteocalcin, which makes its first appearance with the bony fishes [91]. There are several similarities between osteocalcin and MGP:(1) there is 20% homology between osteocalcin and the carboxy-terminal region of MGP (Fig. 2); (2) although MGP is γ-carboxylated at five sites, two of the Gla residues are adjacent to a disulfide bond as in osteocalcin, potentially facilitating interaction of MGP with hydroxyapatite; (3) like osteocalcin, MGP contains a 19-amino acid N-terminal hydrophobic transmembrane secretion signal which is cleaved during translocation into the rough endoplasmic reticulum [92]. A difference between MGP and osteocalcin is that MGP does not undergo amino-terminal proteolytic processing of a propeptide. Rather the consensus sequence for γ-carboxylation resides in the N-terminus of the mature protein. Because MGP retains the recognition sequence, decarboxylated MGP is an excellent in vitro substrate of γ-carboxylase [93]. MGP is phosphorylated on serines 3, 6, and 9 in the N-terminus (Fig. 4). These serines are in a consensus sequence of Ser-X-Glu/Ser(P), also found in milk caseins,

Figure 4

Diagram of MGP interactions. The N-terminus is an anionic domain due to the high density of negative charges from phosphoserines and Gla, and appears necessary to target MGP for secretion. The Gla-containing domain from 35–54 contains four of the five Gla residues and binds BMP-2. A region near the physiological C-terminal from 61–77 (human) can bind to the matrix proteins vitronectin and fibronectin.

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salivary proteins and some regulatory peptides (IL-6, proenkephalin A, growth factor-binding protein 1) [94]. It is not known whether MGP is a substrate for acid and alkaline phosphatases, or whether the properties and function of MGP are altered by dephosphorylation, similar to that observed in osteopontin when its degree of phosphorylation is altered [95]. The phosphoserines are implicated in the correct targeting of MGP for secretion from the cell. Overexpressed nonphosphorylated MGP localizes to the cytoplasm of VSMC but the phosphorylated form of MGP is secreted whether or not it contains Gla [96]. Proteolytic processing of MGP occurs at the C-terminus. Cleavage of the signal peptide leaves an 84-amino acid long protein [92]. However, MGP is isolated from human bone only as the 77-amino acid protein with a carboxyterminal phenylalanine, whereas 1–79 and 1–83 forms are isolated from bovine bone. No MGP protein corresponding to the 1–84 sequence has ever been detected [97]. Intracellular processing of an 84-amino acid MGP by the action of a carboxypeptidase would yield the 1–83 protein; whereas the 1–77 and 1–79 forms could derive from sequential proteolysis via a trypsin-like activity followed Table I.

B. Tissue Localization MGP is synthesized by many kinds of cells derived from normal tissue from bone, cartilage, blood vessels, lung, kidney, heart, and eye [99–106]. Multiple cancers express MGP, e.g. renal cell, prostate, breast, ovarian, glioma, and testicular germ-cell tumors [107–109]. In vivo, MGP mRNA has been found in most tissues examined with the highest levels in kidney, heart, lung, and cartilage (Table I). Many of the early observations of Gla-containing proteins in nonhepatic tissues may in fact now be ascribed to MGP. Once secreted, MGP can anchor to the extracellular mineralized matrix of cartilage and bone. The protein occurs normally only in bone, cartilage,

In vivo MGP Content, Localization, and mRNA Expression in Development, Pathological Calcification, or Cancer

Process Development Chondrogenesis

Circulatory system

Kidney development

Lung morphogenesis

Pathologic calcifications Growth plate Arterial wall Atherosclerosis

Fracture callus

Calcifying pilomatricoma Cancer and apoptosis Cancer

Apoptosis

by removal of an end-terminal arginine [90]. Proteolytic processing of the C-terminal may be a mechanism to control extracellular deposition of MGP, as we have shown that a peptide encompassing amino acids 61–77 of the human sequence binds to the ECM proteins fibronectin and vitronectin [98 and unpublished data] (Fig. 4).

Observation mRNA in mouse resting, proliferative, late hypertrophic Chondrocytes of growth plate, limb bud prechondrocytes Newborn monkey protein more widely distributed than adult No change in rat protein levels from 6–30 months age Protein in monkey vascular smooth muscle mRNA and protein in human artery vascular smooth muscle mRNA in mouse aortic valves and media of arteries mRNA and protein in weanling rat kidney mRNA in mouse kidney medulla mRNA and protein in rat kidney tubules peak after birth Then mRNA levels decrease after 1 month mRNA and protein in weanling rat lung mRNA in developing trachea, differentiating tubules, later in cartilaginous components of bronchi mRNA levels in rat lung decrease rapidly after 1 month MGP null mice exhibit disorganized chondrocyte columns, abnormal calcification, short stature and osteopenia MGP-null mice had chondrocyte-like cells in arteries, matrix vesicles and calcification of all elastic and muscular arteries mRNA in human lesion, highest in macrophages and smooth muscle cells, and lumen-lining endothelial cells Protein in the matrix around lipid-rich, calcified areas mRNA increases with progression of rabbit lesion mRNA in fibroblast-like and chondrocytes of early rat callus proliferating and hypertrophic chondrocytes and trabecular surface osteoblasts of 9-day-old rat callus Protein seen in monkey osteoclasts mRNA enhanced in pilomatricoma tissues mRNA in malignant human breast cells mRNA in human breast cancers mRNA in urogenital malignancies mRNA induced by rat prostate apoptosis

Chapter 3 Vitamin K Dependent Proteins of Bone and Cartilage

and calcified cartilage. MGP is particularly abundant in calcified costal cartilage, in which it accounts for 35–40% of the total protein [91]. MGP content in calcified cartilage and bone matrix remains constant throughout adult life [110]. The MGP in mature rat bone (0.4 mg/g) is about 10% of the osteocalcin level on a molar basis [46]. Small amounts of MGP are present in normal adult arteries, but MGP is present in many soft tissue organs in the neonatal period, and in juvenile animals [101, 110, 111]. It is not known if the MGP in soft tissues is γ-carboxylated, phosphorylated, and C-terminally proteolytically processed as is the MGP in mature bone or cartilage [94, 97]. In situ hybridization analysis shows that MGP mRNA is present during embryonic development as early as day 10.5, prior to the onset of skeletal mineralization. Its expression is restricted to the epithelial–mesenchymal interface, principally in lung and limb buds. By day 12.5 MGP is predominantly expressed in cartilage and blood vessels. In cells of the chondrocytic lineage, expression occurs both in areas that will become ossified and in areas that will remain cartilaginous, such as the trachea and bronchi. Later, MGP is restricted to cartilaginous regions such as bronchi and trachea; alveolar cells do not express the protein. MGP is found in the media of arteries and in the aortic valve. Low levels are also found in kidney medulla [11, 102]. Skeletal expression of MGP occurs in resting, proliferative, and late hypertrophic chondrocytes but not in early hypertrophic chondrocytes or osteoblasts. Primary cultures of osteoblasts have been shown to synthesize and secrete MGP. During osteoblast differentiation in vitro, MGP expression is maximal during maturation of the extracellular collagenous matrix which follows the down-regulation of cellular proliferation [112]. Similarly, in osteosarcoma cell lines MGP synthesis is often lower if differentiated osteoblast markers such as osteocalcin are high [113, 114]. MGP is synthesized by arterial smooth muscle cells in adult human, adult cynomolgus monkey, and in developing mice [11, 115, 116]. In humans, the protein is deposited by smooth muscle cells near calcified areas of atherosclerotic plaque in association with lipid-rich matrices and in the media of arteries and atheromatous intima [115]. MGP synthesis increases with progression of atherosclerosis in a rabbit model [117].

C. Regulation MGP synthesis is stimulated by developmental and pathological events. Thus, many hormones, vitamins, and growth factors are able to regulate synthesis of MGP.

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Control of the gene is influenced by multiple signals that include vitamin K availability, and hormone or morphogenic factors such as retinoic acid (RA), 1,25-dihydroxyvitamin D3, EGF, bFGF, and TGF-β. Few substances have a universal effect on MGP synthesis due in part to the multiple cell types that make MGP. The human MGP gene is present as a single copy on the short arm of chromosome 12 (12p). The gene is 3937 bp long from the transcriptional start site (major 5′ cap site) to the AATAAA polyadenylation signal. Four exons contribute the 615 nucleotides of the MGP mRNA and 103 amino acids comprise the initial translation product [92]. Between mouse and man the intron–exon structure is highly conserved, as is the 5′ flanking nucleotide sequence. However, while the human gene contains a putative upstream regulatory element for vitamin D (VDRE), retinoic acid (RARE), and CAT box binding proteins (CCAAT), these are not present in the mouse gene [102]. The MGP-stimulatory response of skeletal cells to 1,25-dihydroxyvitamin D3 is species- and tissue-dependent. 1,25-Dihydroxyvitamin D3 stimulates MGP synthesis in normal rat osteoblasts and chondrocytes [112] and in human and rat osteosarcoma cells [113, 118, 119]. It does not stimulate MGP synthesis in normal human adult or fetal osteoblasts, fetal chondrocytes, or newborn fibroblasts [119]. Retinoic acid is a major regulator of MGP synthesis. It stimulates synthesis in skeletal cells [118–120] but inhibits synthesis in kidney epithelial cells [121]. The presence of estrogen receptor can alter the response of breast cancer cells to retinoic acid. In MCF-7 human breast cancer cells with estrogen receptors, RA inhibits synthesis [121, 122], whereas RA treatment of breast cancer cells either lacking or with low levels of the estrogen receptor results in an increase in MGP gene expression [122]. Epithelial growth factor (EGF) and basic fibroblast growth factor inhibit MGP gene expression, while plateletderived growth factor has no effect. TGF-β1 stimulates MGP synthesis [123, 124]. In preosteoblasts the effect is specific for TGF-β1, as the closely related BMP-4 does not stimulate MGP synthesis [123]. Large increases in MGP synthesis are found in postconfluent cell culture [112]. MGP levels decline 50– 100-fold when rat kidney cells are subcultured 1:30 [125]. This induction may be due to interaction of the cells with the extracellular matrix. Rat preosteoblasts, differentiating skeletal cells, and kidney epithelial cells increase MGP synthesis in the presence of ascorbic acid after the deposition of a competent collagenous matrix [99, 112, 118]. Conversely, EGF down-regulates the expression of MGP in post-confluent cells at a time when collagen message is reduced [125]. It has been suggested that MGP may

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play some role in cell–cell interactions. However early studies suggesting interaction with integrins have not been substantiated [126, 127]. Rather, a fibronectin-like contaminant in some MGP preparations was probably responsible for cell attachment activity attributed to MGP. It is tempting to speculate that MGP expression is linked to apoptosis of certain cell lineages. MGP is differentially expressed after 1,25-dihydroxyvitamin D3 treatment in C6.9 rat glioma cells which undergo a cell death program when exposed to the hormone [128]. MGP synthesis is increased during apoptosis of prostate cells after androgen withdrawal [129]. It is also increased in many human cancers and it is possible that MGP reflects apoptosis in these tumor cells [130, 131]. It is not clear what role, if any, MGP has in the conversion of normal cells to cancers, but the tumors found to express MGP may reflect upon the lineage of cells that transformed into tumor cells. Indeed a potential role in apoptosis might explain the differential MGP expression in late hypertrophic chondrocytes in the growth plate, but not in early hypertrophic chondrocytes. Perhaps the inability to synthesize MGP in growth plates of MGP-null mice leads to disruption of apoptosis and contributes to the disorganization of the normal columnar orientation of the cells.

D. Function of MGP Studies of MGP function have focused on its ability to inhibit calcification, chondrogenesis, and osteogenesis. MGP binds and regulates BMP-2 activity. MGP also binds hydroxyapatite and is a component of a fetuincalcium-phosphate transport complex in serum. Loss of MGP gene expression leads to Keutel syndrome, characterized by calcification of cartilage and arterial stenosis and calcification [12, 13]. Thus, MGP functions as a regulator of cartilage development and inhibits mineralization. In mice with a non-functional MGP gene (MGPnull) excessive mineralization of arteries and cartilage occurs, showing MGP normally functions to inhibit mineralization in these tissues [11, 14]. Several phenotypic abnormalities were observed, which are consistent with the pattern of expression of MGP. First, growth plate cartilage from MGP-null mice exhibited disorganized columns of chondrocytes with calcification extending into the zone of proliferating chondrocytes. This resulted in the inability to appropriately replace the cartilage with normal bone matrix and short stature, osteopenia, and fractures ensued in these animals. The second phenotypic change in the MGP-null mice was the calcification of all elastic and muscular arteries. By 2–3 weeks after birth, the aortic media was calcified

with disruption of normal artery wall tissue structure. Death resulted within 2 months of birth from rupture of the aorta which contained apatite similar to that found in atherosclerotic lesions. However, no atherosclerotic plaques were observed in these animals. The calcified areas contained hypertrophic chondrocytes with typical cartilage matrix containing matrix vesicles, type II collagen, and proteoglycan, recapitulating the initial stages of skeletal calcification [11]. The presence of chondrocytes may also represent an attempt to repair structural failures in the calcified artery wall in response to compressive forces, similar to those that are known to encourage chondrocyte-like cells in healing tendon and bone. Arterial calcification is a common complication associated with atherosclerosis. Cultures of calcifying vascular smooth muscles cells undergo osteoblast-like differentiation [132–134]. cAMP provokes the in vitro calcification of these cells and increases MGP synthesis as well as that of other early markers of osteoblast differentiation [135]. It appears that the increase in MGP in human vascular calcification may serve to regulate deposition of mineral and ectopic formation of skeletal tissues. No gross abnormalities were observed in embryonic and newborn MGP-null mice, despite the fact that the protein is expressed in affected tissues early during development. MGP is likely to be part of a regulatory system for BMPs which have sufficient alternative or redundant inhibitors such as sclerostin, noggin, and chordin. The necessity for alternative inhibitors of skeletogenesis and mineralization during development may derive from the fact that there is little placental transfer of vitamin K to the embryo. In the fetus and newborn, vitamin K is maintained at levels less than necessary to achieve full γ-carboxylation of vitamin K-dependent proteins (see below). This begs the question, however, of the significance of early expression of the protein and suggests that MGP may function in cell differentiation or proliferation/ survival processes through interactions with extracellular matrix (ECM) as well as BMPs. MGP binds and regulates the function of BMP-2 [136–139]. Indeed, MGP was originally purified as a contaminant of BMP purifications from bone extracts [140]. MGP regulation of BMP-2 may be important for regulating chondrogenesis and osteogenesis. For example, ectopic cartilage forms in arteries of MGP-deficient animals [11, 14, 102]. The results imply that dysregulation of BMP-2 results in ectopic cartilage and bone. Arterial calcification in warfarin-treated animals supports the necessity for Gla residues in order to inhibit calcification [141]. This has been confirmed by the inability of MGP to bind BMP-2 and inhibit mineralization unless it contains Gla [139]. Additional evidence supports the necessity

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of Gla residues for its calcification inhibitory activity. Murshed et al. also showed that MGP expressed by osteoblasts inhibited intramembraneous bone mineralization in MGP-null mice, but mutant MGP that was Gla deficient did not inhibit bone mineralization [142]. In vivo only locally synthesized MGP and not serum-derived MGP could rescue the arterial calcification in MGP-null mice. MGP synthesized by vascular smooth muscle cells rescued arterial calcification but not the cartilage mineralization phenotype. MGP synthesized in liver cells could not rescue the MGP calcification phenotype but serum containing high levels of MGP inhibited in vitro nodule calcification by osteoblasts. Osteocalcin could not rescue the MGP-null phenotype, nor could it inhibit intramembraneous bone mineralization in vivo [142]. Thus, MGP function is not duplicated by osteocalcin, and requires local synthesis of normally γ-carboxylated protein to inhibit calcification. Figure 4 shows the region of MGP with BMP-2 binding activity. The specificity of MGP for BMPs other than BMP-2 is not known, but it has been postulated that MGP inhibits BMP-4 due to its effects on branching morphogenesis in the lung [143]. Accumulating evidence implicates MGP as a developmental regulator of BMP activity. In addition, MGP in adult tissues is likely to regulate BMP activity to maintain adult tissue function. It will be necessary in future to determine the specificity of MGP for binding and inhibiting other BMP homodimers and heterodimers. It is likely that MGP will be able to bind more than one BMP type. Other BMP ligands like sclerostin, the SOST gene product, limits bone formation by inhibiting BMP-6, BMP-7, and possibly BMP-2 and BMP-4 [144]. Sclerostin can also bind and inhibit noggin, another BMP antagonist [145]. Apart from its interaction with BMP-2, MGP has properties that suggest it has multiple functions in the regulation of skeletogenesis and mineralization. MGP is a component of a serum complex of the glycoprotein fetuin with calcium and phosphate [146]. The inverse relationship between arterial calcification and serum fetuin concentrations suggests that the transport system is necessary for cardiovascular maintenance [147]. MGP binds to hydroxyapatite and to extracellular matrix proteins. Hydroxyapatite binding by MGP is regulated by the presence of calcium, phosphate, and magnesium. Hydroxyapatite binding is augmented by the presence of millimolar calcium [148]. This property is similar to the calcium-dependent affinity of osteocalcin for hydroxyapaptite that is related to conformational changes in osteocalcin. But so far there have been no structural studies on MGP, due, in part, to its poor solubility in physiological buffers.

Phospho-serines at the MGP N-terminal end appear to be essential for the normal secretion of MGP. Overexpression experiments in vascular smooth muscle cells revealed that phosphorylated MGP is secreted in the microsomal fraction of cells whether Gla replete or Gla deficient [96]. Nonphosphorylated MGP localizes to the cytoplasm of vascular smooth muscle cells whether or not they contain Gla. Figure 4 shows the region containing phosphoserines that are proposed to be important for intracellular trafficking. MGP interacts with extracellular matrix proteins. Price et al. predicted that fibronectin was an MGP-binding protein because a fibronectin-sized integrin-binding protein co-purified with MGP [127]. Preliminary studies confirm that MGP binds to fibronectin, vitronectin, and fibrinogen [98]. Binding to these proteins could localize and anchor MGP to specific areas in extracellular matrices and could serve to modulate the interaction of MGP with BMPs, the fetuin-calcium-phosphate complex, or hydroxyapatite. MGP also inhibits cell attachment to vitronectin, implying that MGP may act as an antiadhesive protein like the matricellular protein SPARC (Osteonectin) [149]. Figure 4 shows the region of MGP that has vitronectin/fibronectin-binding activity.

V. Gas6 A recently described growth factor, Gas6, is the latest member of the family of vitamin K-dependent proteins to be found in bone. Gas6 (the product of growth arrest specific gene 6), is a polypeptide growth factor that is up-regulated in response to serum starvation and downregulated during growth induction in cultured cells [150]. Gas6 displays significant sequence similarity to Protein S (43% amino acid homology), a vitamin K-dependent protein and coregulator of the blood coagulation cascade. Both proteins contain a Gla domain, a thrombin-sensitive segment, an EGF-like domain, and a sex steroid-binding domain (SHB). Protein S is expressed primarily in the liver while Gas6 is widely expressed with the highest levels in lung, intestine, heart, kidney, and hematopoietic tissue [150]. A Gas6 splice variant has been identified and has a similar distribution pattern as Gas6 [151]. Gas6 and Protein S act as ligands for the AUS class of tyrosine kinase receptors (RTK). The members of this receptor subfamily include Axl (also called Ufo and Ark), Tyro3 (also known as Rse, Brt, Tif, and Sky) and Mer (also called cEyk). These RTKs share about 35% homology and a conserved general domain structure, suggesting that they are recognized by related ligands. These three related RTKs display differential patterns of expression.

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Tyro3 is most prominently expressed in the adult nervous system, although it is also expressed in kidney, ovary, testis, differentiating embryonic stem cells, and a number of hematopoietic cell lines. Axl and Mer are also expressed in the nervous system but are more widely expressed than Tyro3 in peripheral tissues [152, 153]. The SHB domain of Gas6 is responsible for its binding to tyrosine kinase receptors. The Gla domain is important for receptor activation by the calcium-dependent binding of Gla residues to the phosphatidyl serines of the membrane [154, 155]. Although the exact physiological function for Gas6 and its receptors remains to be discovered, several studies show that they may be important for growth regulation. Gas6 has mitogenic activity for Schwann cells and NIH3T3 cells [156, 157]. Bellosta et al. found that Gas6 increased cell survival by protecting NIH3T3 cells from apoptosis induced by serum starvation [158]. Interestingly, there are several reports that Gas6 is important in vascular smooth muscles cells. Gas6 has Gla-dependent growth potentiating activity in vascular smooth muscle cells which may work in conjunction with G protein receptor-mediated signaling pathways [159–161]. It also induces receptormediated chemotactic activity of smooth muscles cells [162]. Recently Gas6 and Axl were found in the synovial fluid and tissue from patients with rheumatoid arthritis and osteoarthritis which suggests a role for this interaction in the maintenance of vasculature in RA [163]. Relevant to bone, a recent study by Nakamura et al. showed that Protein S and Gas6 were direct regulators of osteoclasts [15]. Gas6 induced the rapid phosphorylation of endogenous Tyro3 receptors on osteoclasts derived from 10-day-old rabbits. Osteoclast activity, as assessed by pit area, was increased in a dose-dependent manner by both proteins, but differentiation of osteoclast precursors was not affected. In addition, the authors reported that bone marrow stromal cells express Gas6 and, furthermore, that its expression is regulated by parathyroid hormone and 1,25(OH)2 vitamin D3. These reports are interesting in light of the previous observation that Protein S is secreted and synthesized by osteoblasts, suggesting that Protein S (and perhaps Gas6) provides a link between the osteoblast and the osteoclast [16]. Gas6 has also been detected in human cartilage and chondrocytes. Furthermore, both Gas6 and conditioned medium from chondrocyte cultures stimulated Axl tyrosine phosphorylation in chondrocytes [164].

VI. VITAMIN K/WARFARIN The presence of Gla-containing proteins in bone suggests that vitamin K may play a role in skeletal health.

Several studies have directly assessed vitamin K status in healthy individuals and those with osteoporosis. Low dietary phylloquinone (vitamin K1) intakes or low circulating vitamin K levels have been associated with increased hip fracture risk, particularly among postmenopausal women [165–167]. However, the associations between dietary phylloquinone intake and bone mineral density (BMD) are equivocal. Among elderly men and women participating in the Framingham Heart Study, low dietary phylloquinone intakes were not associated with low BMD at either the hip or spine [166]. In contrast, among the younger participants in the Framingham Offspring Study, dietary phylloquinone intakes were associated with BMD at the hip and spine in pre- and postmenopausal women, but not men [168]. Studies assessing the direct effect of warfarin therapy on bone density in humans have been conflicting. Two studies showed no effect of warfarin at the spine and hip [169, 170] while another two studies showed modest reduction at these sites [171, 172]. In a large multicenter study there was no difference in spine or total hip BMD or fracture rate in 150 Caucasian women 65 years old or older using warfarin compared to non-users [173]. The degree of undercarboxylated osteocalcin (ucOC) in serum is the most sensitive measure of vitamin K nutritional status [174]. In prospective studies, ucOC has been associated with an increased risk of hip fracture in elderly institutionalized and free-living men and women [175–178]. Elevated serum ucOC has also been associated with low BMD in postmenopausal women [179]; similarly, low plasma phylloquinone concentrations have been associated with low BMD at the spine [180]. The phenotype of the osteocalcin knockout mouse precludes a direct role for osteocalcin per se in any potential negative effects of vitamin K deficiency on bone. Unlike the liver-derived clotting factors, osteocalcin is not maximally carboxylated in humans even when subjects receive vitamin K at the RDI. Supplementation with 420 µg/day of phylloquinone results in a 41% increase in carboxylation of osteocalcin [181]. It has recently been shown that a patient with warfarin sensitivity had an alanine to threonine mutation at residue −10 in the propeptide region of factor IX. This alanine residue is invariantly found in all vitamin K dependent proteins except osteocalcin, which has a glycine in the homologous position. Both amino acid substitutions (ala-thr and ala-gly) resulted in reduced affinity for the vitamin K-carboxylase enzyme in vitro [5]. It is interesting to speculate that the osteoblast may have relatively greater vitamin K requirements than the liver to impede deposition of osteocalcin. Thus, naturally occurring undercarboxylation of osteocalcin may represent a selective advantage for the protection of bone.

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Nothing is known about the normal carboxylation status of the other vitamin K-dependent proteins involved with bone and cartilage. It is not known if MGP is maximally carboxylated in all tissues in which it is expressed and at all developmental stages, but the majority of MGP made in atherosclerotic plaque is Gla-deficient when measured by specific antibodies [139]. The administration of vitamin K antagonists may produce adverse side effects by inducing and stimulating cardiovascular calcification [182]. Any effects of vitamin K deficiency or warfarin administration may be regulated in a cell-specific way and the mechanism for control dependent upon the function of the gene product. Warfarin increased levels of MGP mRNA in cultured ROS 25/1 osteosarcoma cells and in cultured rat pneumocytes but had no effect in prechondrocytes and fetal calvarial bone organ culture MGP mRNA levels were unchanged [183, 184]. Furthermore, warfarin had opposing effects on osteocalcin and MGP mRNA levels in the same cells [183]. Rats chronically treated with warfarin show premature closure of the epiphyseal growth plate and a denser cortical bone shaft [185], likely reflecting the effect of the drug on expression of both MGP and osteocalcin. Furthermore, carboxylation may play differing roles in the intracellular fate of the various vitamin K-dependent proteins. Acarboxyprothrombin is retained by rat hepatocytes, but secreted to variable degrees by human and bovine cells [186]. On the other hand, the presence of Gla residues is not absolutely required for production of the mature circulating osteocalcin species [187]. Recent studies suggest that extrahepatic vitamin K-dependent proteins respond to vitamin K depletion or repletion differently than do the liver-derived vitamin K-dependent proteins and probably reflects the diverse biological functions of these proteins [188]. The differing requirements for vitamin K in different tissues is illustrated by recent studies which indicate that regulation of placental transfer of vitamin K may be tightly regulated to allow for appropriate signaling pathways in embryogenesis [189]. Modern tools in molecular biology are leading to the discovery of additional vitamin K-dependent proteins. Four novel proteins, proline-rich Gla proteins (PRGP1 and PRGP2) and transmembrane Gla proteins (TMG3 and TMG4), have recently been identified which are single-pass transmembrane proteins expressed in multiple tissues in fetal and adult life [6, 7]. It is highly likely that there are other vitamin K-dependent proteins which are as yet unknown. Potential involvement of Glacontaining proteins with signal transduction pathways and/or calcium homeostasis may be responsible for any underlying bone pathology resulting from vitamin K deficiency.

References 1. Suttie, J.W., ed. (1987). ‘Current Advances in Vitamin K Research.’ Elsevier, New York. 2. Levy, R. J., Gundberg, C. M., and Scheinman, R. (1983). The presence of the bone specific vitamin K dependent protein, osteocalcin, in calcified atheroscleratic plaque and mineralized heart valves. Atherosclerosis 46, 49–56. 3. Lian, J. B., Skinner, M., Glimcher, M. J., and Gallop, P. M. (1976). The presence of gamma-carboxyglutamic acid in the proteins associated with ectopic calcification. Biochem. Biophys. Res. Commun. 73, 349–355. 4. Nakagawa, Y., Abram, V., Kezdy, F. J., Kaiser, E. T., and Coe, F. L. (1983). Purification and characterization of the principal inhibitor of calcium oxalate monohydrate crystal growth in human urine. J. Biol. Chem. 258, 12594–12600. 5. Berkner, L., and Pudota, B. N. (1998). Vitamin K-dependent carboxylation of the carboxylase. Proc. Natl Acad. Sci. USA 95, 466–471. 6. Kulman, J. D., Harris, J. E., Haldeman, B. A., and Davie, E. W. (1997). Primary structure and tissue distribution of two novel proline-rich gamma-carboxyglutamic acid proteins. Proc. Natl Acad. Sci. USA 94, 9058–9062. 7. Kulman, J. D., Harris, J. E., Xie, L., and Davie, E. W. (2001). Identification of two novel transmembrane gamma-carboxyglutamic acid proteins expressed broadly in fetal and adult tissues. Proc. Natl Acad. Sci. USA 98, 1370–1375. 8. Price, P. A., Urist, M. R., and Otawara, Y. (1983). Matrix Gla protein, a new gamma-carboxyglutamic acid-containing protein which is associated with the organic matrix of bone. Biochem. Biophys. Res. Commun. 117, 765–771. 9. Price, P. A., and Williamson, M. K. (1985). Primary structure of bovine matrix Gla protein, a new vitamin K dependent bone protein. J. Biol. Chem. 260, 14971–14975. 10. Cancela, M. L., Ohresser, M. C. P., Reia, J. P., Viegas, C. S. B., Williamson, M. K., and Price, P. A. (2001). Matrix Gla Protein in Xenopus laevis: Molecular cloning, tissue distribution, and evolutionary considerations. J. Bone. 9, 1611–1622. 11. Luo, G., Ducy, P., McKee, M. D., Pinero, J. P., Loyer, E., Behringer, R. R., and Karsenty, G. (1997). Spontaneous calcification of arteries and cartilage in mice lacking Matrix Gla protein. Nature 386, 78–81. 12. Munroe, P. B., Olgunturk, R. O., Fryns, J.-P., Van Maldergem, L., Ziereisen, F., Yuksel, B., Gardiner, R. M., and Chung, E. (1999). Mutations in the gene encoding the human MGP cause Keutel syndrome. Nat. Gen. 21, 142–144. 13. Meier, M., Weng, L.P., Alexandrakis, E., Ruschoff, J., and Goeckenjan, G. (2001). Tracheobronchial stenosis in Keutel Syndrome. Eur. Respir. J. 17, 566–569. 14. El-Maadawy, S., Kaartinen, M. T., Schinke, T., Murshed, M., Karsenty, G., and McKee, M. D. (2003). Cartilage formation and calcification in arteries of mice lacking matrix Gla protein. Connect. Tissue Res. 44, 272–278. 15. Nakamura, Y. S., Hakeda, Y., Takakura, N., Kameda, T., Hamaguchi, I., Miyamoto, T., Kakudo, S., Nakano, T., Kumegawa, M., and Suda, T. (1998). Tyro 3 receptor tyrosine kinase and its ligand, gas6, stimulate the function of osteoclasts. Stem Cells (Japan) 16, 229–238. 16. Maillard, C., Berruyer, M., Serre, C. M., Dechavanne, M., and Delmas, P. D. (1992). Protein-S, a vitamin K-dependent protein, is a bone matrix component synthesized and secreted by osteoblasts. Endocrinology 130, 1599–1604. 17. Nishimoto, S. K., Araki, N., Robinson, F. D., and Waite, J. H. (1992). Discovery of bone gamma-carboxyglutamic acid protein in mineralized scales. The abundance and structure of Lepomis macrochirus bone gamma-carboxyglutamic acid protein. J. Chem. 267, 11600–11605.

66 18. Nishimoto, S. K., Waite, J. H., Nishimoto, M., and Kriwacki, R. W. (2003) Structure, activity, and distribution of fish osteocalcin. J. Chem. 278, 11843–11848. 19. Cairns, J. R., and Price, P. A. (1994). Direct demonstration that the vitamin K-dependent bone Gla protein is incompletely g-carboxylated in humans. J. Bone Min. Res. 9, 1989–1997. 20. Pan, L. C., and Price, P. A. (1985). The propeptide of rat bone g-carboxyglutamic acid protein shares homology with other vitamin K-dependent protein precursors. Proc. Natl Acad. Sci. USA 82, 6109. 21. Price, P. A., Fraser, J. D., and Metz-Virca, G. (1987). Molecular cloning of matrix Gla protein: implications for substrate recognition by the vitamin K-dependent γ-carboxylase. Proc. Natl Acad. Sci. USA 84, 8335–8339. 22. Jorgensen, M. J., Cantor, A. B., Furie, B. C., Brown, C. L., Shoemaker, C. B., and Furie, B. (1987). Recognition site directing vitamin K-dependent gamma-carboxylation resides on the propeptide of Factor IX. Cell 48, 185–191. 23. Houben, R. J., Jin, D., Stafford, D. W., Proost, P., Ebberink, R. H., Vermeer, C., and Soute, B. A. (1999). Osteocalcin binds tightly to the gamma-glutamylcarboxylase at a site distinct from that of the other known vitamin K-dependent proteins. Bioch. J. 341, 265–269. 24. Gundberg, C. M., and Clough, M. E. (1992). The osteocalcin propeptide is not secreted in vivo or in vitro. J. Bone Min. Res. 7, 73–80. 25. Hauschka, P. V., and Carr, S. A. (1982). Calcium-dependent alpha-helical structure in osteocalcin. Biochemistry 21, 2538–2547. 26. Dowd, T. L., Rosen, J. F., Li, L., and Gundberg, C. M. (2003). The three-dimensional structure of bovine calcium ion-bound osteocalcin using 1H NMR spectroscopy. Biochemistry 42, 7769–7779. 27. Hoang, Q. Q., Sichert, F., Howard, A. J., and Yang, D. S. C. (2003). Bone recognition mechanism of porcine osteocalcin from crystal structure. Nature 425, 977–980. 28. Malone, J. D., Teitelbaum, S. L., Griffin, G. L., Senior, R. M., and Kahn, A. J. (1982). Recruitment of osteoblast precursors by purified bone matrix constituents. J. Cell Biol. 92, 227–230. 29. Lian, J. B., Dunn, K., and Key, L. L. (1986). In vitro degradation of bone particles by human monocytes is decreased with the depletion of the vitamin K-dependent bone protein from the matrix. Endocrinology 118, 1636–1642. 30. Chenu, C., Colucci, S., Grano, M., Zigrino, P., Barattolo, R., Zambonin, G., Baldini, N., Vergnaud, P., Delmas, P. D., and Zallone, A. Z. (1994). Osteocalcin induces chemotaxis, secretion of matrix proteins, and calcium-mediated intracellular signaling in human osteoclast-like cells. J. Cell Biol. 127, 1149–1158. 31. Baumgrass, R., Williamson, M. K., and Price, P. A. (1997). Identification of peptide fragments generated by digestion of bovine and human osteocalcin with the lysosomal proteinases cathepsin B, D, L, H, and S. J. Bone Min. Res. 12, 447–455. 32. Hauschka, P. V., and Reid, M. L. (1978). Timed appearance of a calcium-binding protein containing g-carboxyglutamic acid in developing chick bone. Dev. Biol. 65, 426. 33. Lian, J. B., Roufosse, A. H., and Reit, B. (1982). Concentrations of osteocalcin and phosphoprotein as a function of mineral content and age in cortical bone. Calcif. Tissue Int. 34, S82–S87. 34. Mark, M. P., Prince, C. W., Gay, S., Austin, R. L., Bhown, M., Finkelman, R. D., and Butler, W. T. (1987). A comparative immunocytochemical study on the subcellular distributions of 44 kDa bone phosphoprotein and bone gammacarboxyglumatic acid (Gla)-containing protein in osteoblasts. J. Bone Min. Res. 2, 337–346. 35. Boivin, G., Morel, G., Lian, J. B., Dubois, P. M., and Meunier, P. J. (1990). Localization of endogenous osteocalcin in neonatal rat bone and its absence in articular cartilage. Influence of warfarin treatment on the localization of osteocalcin. Virchows Archiv. A. Path. Anat. Histopathol. 417, 505–512.

Caren M. Gundberg and Satoru K. Nishimoto 36. McKee, M. D., and Nanci, A. (1993). Ultrastructural, cytochemical and immunocytochemical studies of bone and its interfaces. Cells Mater. 3, 219–243. 37. Hauschka, P. V., and Gallop, P. M. (1977). Purification and calciumbinding properties of osteocalcin, the gamma-carboxyglutamatecontaining protein of bone. In R. Wasserman, et al (eds): Calcium Binding Proteins and Calcium Function. Elsevier/North-Holland, Amsterdam, p. 338. 38. Owen, T. A., Aronow, M., Shalhoub, V., Barone, L. M., Wilming, L., Tassinari, M. S., Kennedy, M. B., Pockwinse, S., Lian, J. B., and Stein, G. S. (1990). Progressive development of the rat osteoblast phenotype in vitro: reciprocal relationships in expression of genes associated with osteoblast proliferation and differentiation during formation of the bone extracellular matrix. J. Cell Physiol. 143, 420. 39. Harris, S., Enger, R., Riggs, B., and Spelsberg, T. (1995). Development and characterization of a conditionally immortalized human fetal osteoblastic cell line. J. Bone Min. Res. 10, 178–184. 40. Pockwinse, S. M., Lawrence, J. B., Singer, R. H., Stein, J. L., Lian, J. B., and Stein, G. S. (1993). Gene expression at single cell resolution associated with development of the bone cell phenotype: ultrastructural and in situ hybridization analysis. Bone 14, 347–352. 41. Gerstenfeld, L. C., Chipman, S. D., Glowacki, J., and Lian, J. B. (1987). Expression of differentiated function by mineralizing cultures of chicken osteoblasts. Dev. Bio. 122, 49–60. 42. Gerstenfeld, L., Lian, J., Gotoh, Y., Lee, D., Landis, W., McKee, M., Nanci, A., and Glimcher, M. (1989). Use of cultured embryonic chicken osteoblasts as a model of cellular differentiation and bone mineralization. Conn. Tissue Res. 21, 215–222. 43. Zhou, H., Choong, P., McCarthy, R., Chou, S., Martin, T., and Ng, K. (1994). In situ hybridization to show sequential expression of osteoblast gene markers during bone formation in vivo. J. Bone Min. Res. 9, 1489–1499. 44. Stein, G. S., Lian, J. B., and Owen, T. A. (1990). Relationship of cell growth to the regulation of tissue-specific gene expression during osteoblast differentiation. FASEB J. 4, 3111–3123. 45. Ducy, P., Desbois, C., Boyce, B., Pinero, G., Story, B., Dunstan, C., Smith, E., Bonadio, J., Goldstein, S., Gundberg, C., Bradley, A., and Karsenty, G. (1996). Increased bone formation in osteocalcin-deficient mice. Nature 382, 448–452. 46. Otawara, Y., and Price, P. A. (1986). Developmental appearance of matrix GLA protein during calcification in the rat. J. Biol. Chem. 261, 10828–10832. 47. Boskey, A. L., Gadaleta, S., Gundberg, C., Doty, S. B., Ducy, P., and Karsenty, G. (1998). FT-IR microspectroscopic analysis of bones osteocalcin-deficient mice provides insight into the function of osteocalcin. Bone 23, 187–196. 48. Wenstrup, R. J., Fowlkes, J. L., Witte, D. P., and Florer, J. B. (1996). Discordant expression of osteoblast markers in MC3T3-E1 cells that synthesize a high turnover matrix. J. Biol. Chem. 271, 10271–10276. 49. Ingram, R. T., Park, Y-K., Clarke, B. L., and Fitzpatrick, L. A. (1994). Age- and gender-related changes in the distribution of osteocalcin in the extracellular matrix of normal male and female bone. J. Clin. Invest. 93, 989–997. 50. Webber, D., Osdoby, P., Hauschka P., and Krukowski, M. (1990). Correlation of an osteoclast antigen and ruffled border on giant cells formed in response to resorbable substrates. J. Bone Min. Res. 5, 401–410. 51. Lian, J. B., Tassinari, M., and Glowacki, J. (1984). Resorption of implanted bone prepared from normal and warfarin-treated rats. J. Clin. Invest. 73, 1223–1226. 52. Reinholt, F. P., Hultenby, K., Oldberg, A., and Heinegard, D. (1990). Osteopontin – a possible anchor of osteoclasts to bone. Proc. Natl Acad. Sci. USA 87, 4473–4475.

Chapter 3 Vitamin K Dependent Proteins of Bone and Cartilage 53. Ritter, N. M., Farach-Carson, M. C., and Butler, W. T. (1992). Evidence for the formation of a complex between osteopontin and osteocalcin. J. Bone and Min. Res. 7, 877–885. 54. Puchacz, E., Lian, J. B., and Stein, G. S. (1989). Chromosomal localization of the human osteocalcin gene. Endocrinology 124, 2648–2650. 55. Yoon, K., Rutledge, S. J. C., Buenage, R. F., and Rodan, G. A. (1988). Characterization of the rat osteocalcin gene: Stimulation of promoter activity by 1, 25-Dihydroxyvitamin D3. Biochemistry 27, 8521–8526. 56. Rahman, S., Oberdorf, A., Montecino, M., Tanhauser, S. M., Lian, J. B., Stein, G. S., Laipis, P. J., and Stein, J. L. (1993). Multiple copies of the bone-specific osteocalcin gene in mouse and rat. Endocrinology 133(6), 3050–3053. 57. Bedbois, C., Hogue, D. A., and Karsenty, G. (1994). The mouse osteocalcin gene cluster contains three genes with two separate spatial and temporal patterns of expression. J. Biol. Chem. 269, 1183–1190. 58. Ducy, P., and Karsenty, G. (1995) Two distinct osteoblast-specific cis-acting elements control expression of a mouse osteocalcin gene. Mol. Cell. Biol. 15, 1858–1869. 59. Schinke, T., and Karsenty, G. (1999). Characterization of Osf1, an osteoblast-specific transcription factor binding to a critical cis-acting element in the mouse Osteocalcin promoters. J. Biol. Chem. 274, 30182–30189. 60. Yang, X., and Karsenty, G. (2004). ATF4, the osteoblast accumulation of which is determined post-translationally, can induce osteoblastspecific gene expression in non-osteoblastic cells. J. Biol. Chem. 279, 47109–47114. 61. Stein, G. S., Lian, J. B., van Wijnen, A. J., and Stein, J. L. (1997). The osteocalcin gene: a model for multiple parameters of skeletal-specific transcriptional control. Mol. Biol. Rep. (Belgium) 24, 185–196. 62. Morrison, N. A., Shine, J., Fragonas, J. C., Verkest, V., McMenemy, L., and Eisman, J.A. (1989). 1,25-dihydroxyvitamin D-responsive element and glucocorticoid repression in the osteocalcin gene. Science 246, 1158–1161. 63. Demay, M. B., Gerardi, J. M., DeLuca, H. F., and Kronenberg, H. M. (1990). DNA sequences in the rat osteocalcin gene that bind the 1,25-dihydroxyvitamin D3 receptor and confer responsive to 1,25dihydroxyvitamin D3. Proc. Natl Acad. Sci. USA 87, 369–373. 64. Markose, E. R., Stein, J. L., Stein, G. S., and Lian, J. B. (1990). Vitamin D-mediated modifications in protein-DNA interactions at two promoter elements of the osteocalcin gene. Proc. Natl Acad. Sci. USA 87, 1701–1705. 65. Kerner, S. A., Scott, R. A., and Pike, J. W. (1989). Sequence elements in the human osteocalcin gene confer basal activation and inducible response to hormonal vitamin D3. Proc. Natl Acad. Sci. USA 86, 4455–4459. 66. Lian, J. B., Shalhoub, V., Aslam, F., Frenkel, B., Green, J., Hamrah, M., Stein, G. S., and Stein, J. L. (1997). Species-specific glucocorticoid and 1,25 dihydroxyvitamin D responsiveness in mouse MC3T3-E1 osteoblasts: dexamethasone inhibits osteoblast differentiation and vitamin D down-regulates osteocalcin gene expression. Endocrinology 138: 2117–2127. 67. Carpenter, T. O., Moltz, K. C., Ellis, B., Andreoli, M., McCarthy, T. L., Centrella, M., Bryan, D., and Gundberg, C. (1998). Osteocalcin production in primary osteoblast cultures derived from normal and Hyp mice. Endocrinology 139, 35–43. 68. Gerstenfeld, L. C., Zurakowski, D., Schaffer, J. L., Nichols, D. P., Toma, C. D., Broess, M., Bruder, S. P., and Caplan, A. L. (1996). Variable hormone responsiveness of osteoblast populations isolated at different stages of embryogenesis and its relationship to the osteogenic lineage. Endocrinology 137, 3957–3968. 69. Zhang, R., Ducy, P., and Karsenty, G. (1997). 1,25-Dihydroxyvitamin D3 inhibits osteocalcin expression in mouse through an indirect mechanism. J. Biol. Chem. 272, 110–116.

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70. Nishimoto, S. K., Salka, C., and Nimni, M. E. (1987). Retinoic acid and glucocorticoids enhance the effect of 1,25-dihydroxyvitamin D3 on bone gamma-carboxyglutamic acid protein synthesis by rat osteosarcoma cells. J. Bone Min. Res. 2, 571–577. 71. Noda, M., Yoon, K., and Rodan, G. A. (1988). Cyclic AMP-mediated stabilization of osteocalcin mRNAa in rat osteoblast-like cell treated with parathyroid hormone. J. Biol. Chem. 263, 18574–18577. 72. Egrise, D., Martin, D., Neve, P., Verhas, M., and Schoutens, A. (1990). Effects and interactions of 17B-estradiol, T3 and 1,25 (OH)2D3 on cultured osteoblasts from mature rats. Bone Min. 11, 273–283. 73. Thiede, M. A., Smock, S. L., Petersen, D. N., Grasser, W. A., Thompson, D. D., and Nishimoto, S. K. (1994). Presence of messenger ribonucleic acid encoding osteocalcin, a marker of bone turnover, in bone marrow megakaryocytes and peripheral blood platelets. Endocrinology 135(3), 929–937. 74. Fleet, J. C., and Hock, J. M. (1994). Identification of osteocalcin mRNA in non-osteoid tissue of rats and humans by reverse transcription-polymerase chain reaction. J. Bone Min. Res. 9, 1565–1573. 75. Calvo, M. S., Eyre, D., and Gundberg, C. M. (1996). Molecular basis and clinical application of biological markers of bone turnover. Endocrine Rev. 17, 333–368. 76. Garnero, P., Grimaux, M., Seguin, P., and Delmas, P. D. (1994). Characterization of immunoreactive forms of human osteocalcin generated in vivo and in vitro. J. Bone Min. Res. 9, 255–264. 77. Rosenquist, C., Quist, P., Bjarnason, N., and Christiansen, C. (1995). Measurement of a more stable region of osteocalcin in serum by ELISA with two monoclonal antibodies. Clin. Chem. 41, 1439–1445. 78. Blumsohn, A., Hannon, R. A., and Eastell, R. (1995). Apparent instability of osteocalcin in serum as measured with different commercially available immunoassays. Clin. Chem. 41, 318–320. 79. Taylor, A. K., Linkhart, S., Mohan, S., Christenson, R. A., Singer, F. R., and Baylink, D. J. (1990). Multiple osteocalcin fragments in human urine and serum as detected by a midmolecule osteocalcin radioimmunoassay. J. Clin. Endocrinol. Metab. 70, 467–472. 80. Chen, J. T., Hosoda, K., Hasumi, K., Ogata, E., and Shiraki, M. (1996). Serum N-terminal osteocalcin is a good indicator for estimating responders to hormone replacement therapy in postmenopausal women. J. Bone Min. Res. 11, 1784–1792. 81. Gorai, L., Hosoda, K., Taguchi, Y., Chacki, O., Nakavama, M., Yoh, K., Yamaii, T., and Minaguchi, H. (1997). A heterogeneity in serum osteocalcin N-terminal fragments in Paget’s disease: A comparison with other biochemical indices in pre- and post-menopause. J. Bone Min. Res. 12, T678. 82. Srivastava, A. K., Mohan, S., Singer, F. R., and Baylink, D. J. (2002). A urine midmolecule osteocalcin assay shows higher discriminatory power than a serum midmolecule osteocalcin assay during short-term alendronate treatment of osteoporotic patients. Bone 31, 62–69. 83. Matikainen, T., Kakonen, S. M., Pettersson, K., Karp, M., Lovgren, T., Vaananen, H. K., and Hellman, J. (1999). Demonstration of the predominant urine osteocalcin fragments detectable by two-site immunoassays. J. Bone Min. Res. 14, 431–438. 84. Ivaska, K. K., Hellman, J., Likojarvi, J., Kakonen, S. M., Gerdhem, P., Akesson, K., Obrant, K. J., Pettersson, K., and Vaananen, H. K. (2003). Identification of nevel proteolytic forms of osteocalcin in human urine. Biochem. Biophys. Res. Comm. 306, 973–980. 85. Salo, J., Lehenkari, P., Mulari, M., Metsikkö, K., and Väänänen, H. K. (1997). Removal of osteoclast bone resorption products by transcytosis. Science 276, 270–273. 86. Ivaska, K. K., Hentunen, T. A., Vääräniemi, J., Ylipahkala, H., Pettersson, K., and Väänänen, H. K. (2004). Release of intact and fragmented osteocalcin molecules from bone matrix during bone resorption in vitro. J. Biol. Chem. 279, 18361–18369. 87. Gerdhem, P., Ivaska, K. K., Alatalo, S. L., Halleen, J. M., Hellman, J., Isaksoon, A., Pettersson, K., Vaananen, H. K., Alesson, K., and

68

88.

89.

90. 91.

92.

93.

94.

95.

96.

97.

98.

99.

100.

101.

102.

103.

104.

105.

Caren M. Gundberg and Satoru K. Nishimoto Obrant, K. J. (2004). Biochemical markers of bone metabolism and prediction of fracture in elderly women. J. Bone Min. Res. 19, 386–393. Page, A. E., Hayman, A. R., Andersson, L. M. B., Chambers, T. J., and Warburton, M. J. (1993). Degradation of bone matrix proteins by osteoclast cathepsins. Internat. J. Biochem. 25, 545–550. Baumgrass, R., Williamson, M. K., and Price, P. A. (1997). Identification of peptide fragments generated by digestion of bovine and human osteocalcin with the lysosomal proteinases cathepsin B, D, L, H, and S. J. Bone Min. Res. 12, 447–455. Novak, J. F., Hayes, J. D., and Nishimoto, S. K. (1997). Plasmin-mediated proteolysis of osteocalcin. J. Bone Min. Res. 12, 1035–1042. Rice, J. S., Williamson, M. K., and Price, P. A. (1994). Isolation and sequence of the vitamin K-dependent Matrix Gla Protein from the calcified cartilage of the soupfin shark. J. Bone Min. Res. 9, 567–576. Cancela, M. L., Hsieh, C., Francke, U., and Price, P. A. (1990). Molecular structure, chromosome assignment, and promoter organization of the human matrix Gla protein gene. J. Biol. Chem. 265, 15040–15048. Engelke, J. E., Hale, J. E., Suttie, J. W., and Price, P. A. (1991). Vitamin K-dependent carboxylase: utilization of decarboxylated bone Gla protein and matrix Gla protein as substrates. Biochim. Biophys. Acta 1078, 31–34. Price, P. A., Rice, J. S., and Williamson, M. K. (1994). Conserved phosphorylation of serines in the Ser-X-Glu/Ser(P) sequences of the vitamin K-dependent matrix Gla protein from shark, lamb, rat, cow and human. Pro. Sci. 3, 822–830. Nemir, M., DeVouge, M. W., and Mukherjee, B. B. (1989). Normal rat kidney cells secrete both phosphorylated and nonphosphorylated forms of osteopontin showing different physiological properties. J. Biol. Chem. 264, 18202–18208. Wajih, N., Borras, T., Xue, W., Hutson, S. M., and Wallin, R. (2004). Processing and transport of Matrix Gla Protein (MGP) and Bone Morphogenetic Protein-2 (BMP-2) in cultured human vascular smooth muscle cells: Evidence for an uptake mechanism for serum fetuin. J. Biol. Chem. 279, 25276–25283. Hale, J. E., Williamson, M. K., and Price, P. A. (1991). Carboxylterminal proteolytic processing of matrix Gla protein. J. Biol. Chem. 266, 21145–21149. Nishimoto, S. K., and Nishimoto, M. (2003). Matrix Gla protein inhibits cell attachment to vitronectin, binds vitronectin, and co-localizes with vitronectin in vivo. Mol. Biol. Cell 14, 338A. Zhao, J., Araki, N., and Nishimoto, S. K. (1995). Quantification of matrix Gla protein by competitive polymerase chain reaction using glyceraldehyde-3-phosphate dehydrogenase as an internal control. Gene 155, 159–165. Zhao, J., and Nishimoto, S. K. (1995). An RNA-competitive polymerase chain reaction method for human matrix Gla protein mRNA measurement. Anal. Biochem. 228, 162–164. Zhao, J., and Nishimoto, S. K. (1996). Matrix Gla protein gene expression is elevated during postnatal development. Matrix Biology 15, 131–140. Luo, G., D’Souza, R., Hogue, D., and Karsenty, G. (1995). The Matrix Gla protein gene is a marker of the chondrogenesis lineage during mouse development. J. Bone Min. Res. 10, 325–334. Fraser, J. D., and Price, P. A. (1988). Lung, heart, and kidney express high levels of mRNA for the vitamin K-dependent protein matrix Gla protein. J. Biol. Chem. 263, 11033–11036. Sakai, R., Kinouchi, T., Kawamoto, S., Dana, M. R., Hamamoto, T., Tsuru, T., Okubo, K., and Yamagami, S. (2002). Construction of human corneal endothelial cDNA library and identification of novel active genes. Invest. Ophthalmol. Vis. Sci. 43, 1749–1756. Ahmed, F., Brown, K. M., Stephan, D. A., Morrison, J. C., Johnson, E. C., and Tomarev, S. I. (2004). Microarray analysis of changes in mRNA levels in the rat retina after experimental elevation of intraocular pressure. Invest. Ophthalmol. Vis. Sci. 45, 1247–1258.

106. Young, T. L., Scavello, G. S., Paluru, P. C., Choi, J. D., Rappaport, E. F., and Rada, J. A. (2004). Microarray analysis of gene expression in human donor sclera. Mol. Vis. 10, 163–176. 107. Levedakou, E. N., Strohmeyer, T. G., Effert, P. J., and Liu, E. T. (1992). Expression of the matrix Gla protein in urogenital malignancies. Int. J. Cancer 52, 534–537. 108. Hough, C. D., Cho, K. R., Zonderman, A. B., Schwartz, D. R., and Morin, P. J. (2001). Coordinately up-regulated genes in ovarian cancer. Cancer Res. 61, 3869–3876. 109. van den Boom, J., Wolter, M., Kuick, R., Misek, D. E., Youkilis, A. S., Wechsler, D. S., Sommer, C., Reifenberger, G., and Hanash, S. M. (2003). Characterization of gene expression profiles associated with glioma progression using oligonucleotide-based microarray analysis and real-time reverse transcription-polymerase chain reaction. Am. J. Pathol. 163, 1033–1043. 110. Nishimoto, S. K., Robinson, F. D., and Snyder, D. L. (1993). Effect of aging and dietary restriction on matrix Gla protein and other components of rat tracheal cartilage. Matrix 13, 373–380. 111. Spronk, H. M. H., Soute, B. A. M., Schurgers, L. J., Cleutjens, J. P. M., Thijssen, H. H. W., DeMey, J. G. R., and Vermeer, C. (2001). Matrix Gla protein accumulates at the border of regions of calcification and normal tissue in the media of the arterial vessel wall. Biochem. Biophys. Res. Commun. 289, 485–490. 112. Barone, L. M., Owen, T. A., Tassinari, M. S., Bortell, R., Stein, G. S., and Lian, J. B. (1991). Developmental expression and hormonal regulation of the rat matrix Gla protein (MGP) gene in chondrogenesis and osteogenesis. J. Cellular Biochem. 46, 351–365. 113. Fraser, J. D., Otawara, Y., and Price, P. A. (1988). 1,25 dihydroxyvitamin D3 stimulates the synthesis of matrix Gla protein by osteosarcoma cells. J. Biol. Chem. 263, 911–916. 114. Nishimoto, S. K., and Price, P. A. (1980). Secretion of the vitamin K-dependent protein of bone by rat osteosarcoma cells. Evidence for an intracellular precursor. J. Biol. Chem. 255, 6579–6583. 115. Carlson, C. S., Tulli, H. M., Jayo, M. J., Loeser, R. F., Tracy, R. P., Mann, K. G., and Adams, M. R. (1993). Immunolocalization of noncollagenous bone matrix proteins in lumbar vertebrae from intact and surgically menopausal cynomolgus monkeys. J. Bone Min. Res. 8, 71–81. 116. Shanahan, C. M., Cary, N. R. B., Metcalf, J. C., and Weissberg, P. L. (1994). High expression of genes for calcification-regulating proteins in human atherosclerotic plaques. J. Clin. Invest. 93, 2393–2402. 117. Sohma, Y., Suzuki, T., Sasano, H., Nagura, H., Nose, M., and Yamamoto, T. (1994). Expression of mRNA for matrix Gla protein during progression of atherosclerosis in aortae of watanabe heritable Hyperlipidemic rabbits. J. Biochem. 116, 747–751. 118. Choong, P. F. M., Martin, T. J., and Ng, K. W. (1993). Effects of ascorbic acid, calcitriol, and retinoic acid on the differentiation of preosteoblasts. J. Orthop. Res. 11, 638–647. 119. Cancela, M. L., and Price, P. A. (1992). Retinoic acid induces matrix Gla protein gene expression in human cells. Endocrinology 130, 102–108. 120. Cancela, M. L., Williamson, M. K., and Price, P. A. (1993). Retinoic acid increases MGP in rat plasma. Nut. Res. 13, 87–91. 121. Kirfel, J., Kelter, M., Cancela, M. L., Price, P. A., and Schule, R. (1997). Identification of a novel negative retinoic acid responsive element in the promoter of the human matrix Gla protein gene. Proc. Natl Acad. Sci. USA 94, 2227–2232. 122. Sheikh, M. S., Shao, Z-M., Chen, J-C., and Fontana, J. A. (1993). Differential regulation of matrix Gla protein (MGP) gene expression by retinoic acid and estrogen in human breast carcinoma cells. Molec. Cell Endocrinol. 92, 153–160. 123. Zhou, H., Hammonds, R. G., Findlay, D. M., Martin, T. J., and Ng, K. W. (1993). Differential effects of transforming growth factor-1 and bone morphogenetic protein 4 on gene expression and differentiated function of preosteoblasts. J. Cell Physiol. 155, 112–119.

Chapter 3 Vitamin K Dependent Proteins of Bone and Cartilage 124. Zhao, J., and Warburton, D. (1997). Matrix Gla protein gene expression is induced by transforming growth factor-b in embryonic lung culture. Am. J. Physiol. 273, L282–L287. 125. Cancela, M. L., Hu, B., and Price, P. A. (1997). Effect of cell density and growth factors on matrix Gla protein expression by normal rat kidney cells. J. Cell Physiol. 171, 125–134. 126. Loeser, R. F., and Wallin, R. (1992). Cell adhesion to matrix Gla protein and its inhibition by an Arg-Gly-Asp-containing peptide. J. Biol. Chem. 267, 9459–9462. 127. Cancela, M. L., Williamson, M. K., and Price, P. A. (1994). The putative RGD-dependent cell adhesion activity of Matrix Gla Protein is due to higher molecular weight contaminants. J. Biol. Chem. 269, 12185–12189. 128. Baudet, C., Perret, E., Delpech, B., Kaghad, M., Brachet, P., Wion, D., and Caput, D. (1998). Differentially expressed genes in C6.9 glioma cells during vitamin D-induced cell death program. Cell Death Differentiation 5, 116–125. 129. Briehl, M. M., and Miesfeld, R. L. (1991). Isolation and characterization of transcripts induced by androgen withdrawal and apoptotic cell death in the rat ventral prostate. Mol. Endocrin. 5, 1381–1388. 130. Chen, L., O’Bryan, J. P., Smith, H. S., and Liu, E. (1990). Overexpression of matrix Gla protein mRNA in malignant human breast cells: isolation by differential cDNA hybridization. Oncogene 5, 1391–1395. 131. Hirota, S., Ito, A., Nagoshi, J., Takeda, M., Kurata, A., Takatsuka, Y., Kohri, K., Nomura, S., and Kitamura, Y. (1995). Expression of bone matrix protein messenger ribonucleic acids in human breast cancers. Lab. Invest. 72, 64–69. 132. Tintut, Y., Parhami, F., Bostrom, K., Jackson, S. M., and Demer, L. L. (1998). cAMP stimulates osteoblast-like differentiation of calcifying vascular cells, potential signaling pathway for vascular calcification. J. Biol. Chem. 273, 7547–7553. 133. Proudfoot, D., Skepper, J. N., Shanahan, C. M., and Weissberg, P. L. (1998). Calcification of human vascular cells in vitro is correlated with high levels of matrix Gla protein and low levels of osteopontin expression. Arterioscler. Thromb. Vasc. Biol. 18(3), 379–388. 134. Bostrom, K., Watson, K. E., Stanford, W. P., and Demer, L. L. (1995). Atherosclerotic calcification: relation to developmental osteogenesis. Am. J. Cardiol. 75, 88–91. 135. Giachelli, C. M., Bae, N., Almeida, M., Denhardt, D. T., Alpers, C. E., and Schwartz, S. M. (1993). Osteopontin is elevated during neointimal formation in rat arteries and is a novel component of human atherosclerotic plaques. J. Clin. Invest. 92, 1686–1696. 136. Bostrom, K., Tsao, D., Shen, S., Wang, Y., and Demer, L. L. (2001). Matrix Gla Protein modulates differentiation induced by Bone Morphogenetic Protein-2 in C3H10T1/2 cells. J. Biol. Chem. 176, 14044–14052. 137. Zebboudj, A. F., Imura, M., and Bostrom, K. (2002). Matrix Gla Protein, a regulatory protein for Bone Morphogenetic Protein-2. J. Biol. Chem. 277, 4388–4394. 138. Wallin, R., Cain, D., Hutson, S. M., Sane, D. C., and Loeser, R. (2000). Modulation of the binding of Matrix Gla Protein (MGP) to bone morphogenetic protein-2 (BMP-2). Thromb. Haemost. 84, 1039–1044. 139. Sweat, A., Sane, C. C., Hustson, C. M., and Wallin, R. (2003). Matrix Gla Protein (MGP) and bone morphogenetic protein-2 in aortic calcified lesions of aging rats. J. Throm. Haemost. 1, 178–185. 140. Price, P. A., Urist, M. R., and Otawara, Y. (1983). Matrix Gla protein, a new Gla-containing protein which is associated with the organic matrix of bone. Biochem. Biophys. Res. Commun. 117, 765–771. 141. Price, P. A., Faus, S. S., and Williamson, M. K. (2000). Warfarin-induced artery calcification is accelerated by growth and vitamin D. Arterioscler. Thromb. Vasc. Biol. 20, 317–327. 142. Murshed, M., Schinke, T., McKee, M. D., and Karsenty, G. (2004). Extracellular matrix mineralization is regulated locally; different roles of two gla-containining proteins. J. Cell Biol. 165, 625–630.

69

143. Gilbert, K. A., and Rannels, S. R. (2004). Matrix GLA Protein modulates branching morphogenesis in fetal rat lung. Am. J. Lung Cell Mol. Physiol. 286, L1179–L1187. 144. Avsian-Kretchmer, O., and Hsueh, A. J. W. (2004). Comparative genomic analysis of the eight-membered ring cystine knot-containing bone morphogenetic protein antagonists. Mol. Endocrinol. 18, 1–12. 145. Winkler, D. G., Yu, C., Geoghegan, J. C., Ojala, E. W., Skonier, J. E., Shpektor, D., Sutherland, M. K., and Latham, J. A. (2004). Noggin and sclerostin bone morphogenetic protein antagonists form a mutually inhibitory complex. J. Biol. Chem. 279, 36293–36298. 146. Price, P. A., Thomas, G. R., Pardini, A. W., Figueira, W. F., Caputo, J. M., and Williamson, M. K. (2002). Discovery of a high molecular weight complex of calcium, phosphate, fetuin, and matrix Gla protein in the serum of etidronate-treated rats. J. Biol. Chem. 277, 3926–3934. 147. Ketteler, M., Borgartz, P., Westenfeld, R., Wildberger, J. E., Mahnken, A. H., Bohm, R., Metzer, T., Wanner, C., Jahnen-Dechent, W., and Floege, J. (2003). Association of low fetuin-A (AHSG) concentrations in serum with cardiovascular mortality in patients on dialysis: a cross-sectional study. Lancet 361, 827–833. 148. Roy, M. E., and Nishimoto, S. K. (2002). Matrix Gla Protein binding to hydroxyapatite is dependent on the ionic environment: Calcium enhances binding affinity but phosphate and magnesium decrease affinity. Bone 31, 296–302. 149. Bornstein, P., and Sage, E.H. (2002). Matricellular proteins: extracellular modulators of cell function. Curr. Opin. Cell Biol. 14, 608–616. 150. Manfioletti, G., Brancolini, C., Avanzi, G., and Schneider, C. (1993). The protein encoded by a growth arrest arrest-specific gene (gas6) is a new member of the vitamin K-dependent proteins related to protein S, a negative coregulator in the blood coagulation cascade. Mol. Cell. Biol. 13, 4976–4985. 151. Marcandalli, P., Gostissa, M., Varnum, B., Goruppi, S., and Schneider, C. (1997). Identification and tissue expression of a splice variant for the growth arrest-specific gene gas6. FEBS Lett. 415, 56–58. 152. Stitt, T. N., Conn, G., Gore, M., Lai, C., Bruno, J., Radziejewski, C., Mattsson, K., Fisher, J., Gles, D. R., Jones, P. F., Masiakowski, P., Ryan, T. E., Tobkes, N. J., Chen, D. H., DiStefano, P. S., Long, G. L., Basilico, C., Goldfarb, M. P., Lemke, G., Glass, D. J., and Yancopoulos, G. D. (1995). The anticoagulation factor protein S and its relative, gas6, are ligands for the tyro 3/axl family of receptor tyrosine kinases. Cell 80, 661–670. 153. Crosier, P. S., Lewis, P. M., and Hall, L. R. (1994). Isolation of a receptor tyrosine kinase (DTK) from embryonic stem cells: structure, genetic mapping and analysis of expression. Growth Factors 11, 125–136. 154. Mark, M. R., Chen, J., Hammonds, R. G., Sadick, M., and Godowsk, P. J. (1996). Characterization of gas6, a member of the superfamily of G domain-containing proteins, as a ligand for Rse and Axl. J. Biol. Chem. 271, 9785–9789. 155. Joseph, D. R. (1997). Sequence and functional relationships between androgen-binding protein/sex hormone-binding globulin and its homologs protein S, gas6, laminin, and agrin. Steroids 62, 578–588. 156. Li, R., Chen, J., Hammonds, G., Phillips, H., Armanini, M., Wood, P., Bunge, R., Godowski, P. J., Sliwkowski, M. X., and Mather, J. P. (1996). Identification of gas6 as a growth factor for human schwann cells. J. Neuroscience 16, 2012–2019. 157. Goruppi, S., Ruaro, E., Varnum, B., and Schneider, C. (1997). Requirement of phosphatidylinositol 3-kinase-dependent pathway and Src for gas6-Axl mitogenic and survival activities in HIH 3T3 fibroblasts. Mol. Cell Biol. 17, 4442–4453. 158. Bellosta, P., Zhang, Q., Goff, S. P., and Basilico, C. (1997). Signaling through the ARK tyrosine kinase receptor protects from apoptosis in the absence of growth stimulation. Oncogene 15, 2387–2397. 159. Nakano, T., Higashino, K., Kikuchi, N., Kishino, J., Nomura, K., Fujita, H., Ohara, O., and Arita, H. (1995). Vascular smooth muscle

70

160.

161.

162.

163.

164.

165.

166.

167.

168.

169.

170.

171.

172. 173.

Caren M. Gundberg and Satoru K. Nishimoto cell-derived, Gla-containing growth-potentiating factor for Ca2+mobilizing growth factors. J. Biol. Chem. 270, 5702–5705. Nakano, T., Ishimoto, Y., Kishino, J., Umeda, M., Inoue, K., Nagata, K., Ohashi, K., Mizuno, K., and Arita, H. (1997). Cell adhesion to phosphatidylserine mediated by a product of growth arrest-specific gene 6. J. Biol. Chem. 272, 29411–29414. Nakano, T., Kawamoto, K., Kishino, J., Nomura, K., Higashino, K., and Arita, H. (1997). Requirement of gamma-carboxyglutamice acid residue for the biological activity of gas6: contribution of endogenous gas6 to the proliferation of vascular smooth muscle cells. Biochem. J. 323, 387–392. Fridell, Y. W. C., Villa, J., Attar, E., and Liu, E. T. (1998). Gas6 induces axl-mediated chemotaxis of vascular smooth muscle cells. J. Biol. Chem. 273, 7123–7126. O’Donnell, K., Harkes, I. C., Dougherty, L., and Wicks, I. P. (1999). Expression of receptor tyrosine kinase AXL and its ligand Gas6 in rheumatoid arthritis: evidence for a novel endothelial cell survival pathway. Am. J. Pathol. 154, 1171–1180. Loesser, R. F., Varnum, B. C., Carlson, C. S., Goldring, M. B., Liu, E. T., Sadiev, S., Kute, T. E., and Wallin, R. (1997). Human chondrocytes expression of growth-arrest-specific gene and the tyrosine kinase receptor axl: potential role in autocrine signaling in cartilage. Arthritis Rheum. 40, 1455–1465. Hodges, S. J., Akesson, K., Vergnaud, P., Obrant, K., and Delmas, P. D. (1993). Circulating levels of vitamin K1 and K2 decreased in elderly women with hip fracture. J. Bone Min. Res. 8, 1241–1245. Feskanich, D., Weber, P., Willett, W. C., Rockett, H., Booth, S. L., and Colditz, G. A. (1999). Vitamin K intake and hip fractures in women: a prospective study. Am. J. Clin. Nutr. 69, 74–79. Booth, S. L., Tucker, K. L., Chen, H., Hannan, M. T., Gagnon, D. R., Cupples, L. A., Wilson, P. W., Ordovas, J., Schaefer, E. J., Dawson-Hughes, B., and Kiel, D. P. (2000). Dietary vitamin K intakes are associated with hip fracture but not with bone mineral density in elderly men and women. Am. J. Clin. Nutr. 71, 1201–1208. Booth, S., Broe, K., Gagnon, D., Tucker, K. L., Hannan, M. T., McLean, R. R., Dawson-Hughes, B., Wilson, P. W., Cupples, L. A., and Kiel, D. P. (2003). Vitamin K intake and bone mineral density in women and men. Am. J. Clin. Nutr. 77, 512–516. Rosen, H. N., Maitland, L. A., Suttie, J. W., Manning, W. J., Glynn, R. J., and Greenspan, S. L. (1993). Vitamin K and maintenance of skeletal integrity in adults. Amer. J. Med. 94, 62–68. Lafforque, P., Daver, L., Monites, J. R., Chagnaud, C., deBoissezon, M. C., and Aquaviva, P. C. (1997). Bone mineral density in patients given oral vitamin K antagonists. Revu Du Rhumatisme 64, 249–254. Philip, W. J. U., Martin, J. C., Richardson, J. M., Reid, D. M., Webster, J., and Douglas, A. S. (1995). Decreased axial and peripheral bone density in patients taking long-term warfarin. QJ Med. 88, 635–840. Monreal, M., Olive, A., and Lafozz, E. (1991). Heparins, coumarin, and bone density. Lancet 338, 706. Jamal, S., Milani, G., Lipschutz, R., Stone, R., Browner, W. S., and Cummings, S. R. (1997). The therapeutic use of Warfarin does not influence post-menopausal osteoporosis. J. Bone Min. Res. 12(S1), T512.

174. Sokoll, L. J., and Sadowski, J. A. (1996). Comparison of biochemical indexes for assessing vitamin K nutritional status in a healthy adult population. Am. J. Clin. Nutr. 63, 566–573. 175. Szulc, P., Arlot, M., Chapuy, M. C., Duboeuf, F., Meunier, P. J., and Delmas, P. D. (1994). Serum undercarboxylated ostepcalcin correlates with hip bone mineral density in elderly women. J. Bone Miner. Res. 9, 1591–1595. 176. Szulc, P., Chapuy, M. C., Meunier, P. J., and Delmas, P. D. (1993). Serum undercarboxylated osteocalcin is a marker of the risk of hip fracture in elderly women. J. Clin. Invest. 91, 1769–1774. 177. Szulc, P., Chapuy, M. C., Meunier, P. J., and Delmas, P. D. (1996). Serum undercarboxylated osteocalcin is a marker of the risk of hip fracture: a three year follow-up study. Bone 18, 487–588. 178 Vergnaud, P., Garnero, P., Meunier, P. J., Breart, G., Kamihagi, K., and Delmas, P. D. (1997). Undercarboxylated osteocalcin measured with a specific immunoassay predicts hip fracture in elderly women: the EPIDOS Study. J. Clin. Endocrinol. Metab. 82, 719–724. 179. Luukinen, H., Kakonen, S. M., Pettersson, K., Koski, K., Laippala, P., Lovgren, T., Kivela, S. L., and Vaananen, H. K. (2000). Strong prediction of fractures among older adults by the ratio of carboxylated to total serum osteocalcin. J. Bone Miner. Res. 15, 2473–2478. 180. Kanai, T., Takagi, T., Masuhiro, K., Nakamura, M., Iwata, M., and Saji, F. (1997). Serum vitamin K level and bone mineral density in post-menopausal women. Int. J. Gynaecol. Obstet. 56, 25–30. 181. Sokoll, L. J., Booth, S. L., O’Brien, M. E., Davidson, K. W., Tsaioun, K. I., and Sadowski, J. A. (1997). Changes in serum osteocalcin, plasma phylloquinone, and urinary gamma-carboxyglutamic acid in response to altered intakes of dietary phylloquinone in human subjects. Am. J. Clin. Nutr. 65, 779–784. 182. Schurgers, L. J., Aebert, H., Vermeer, C., Bueltmann, B., Janzen, J. (2004). Oral anticoagulant treatment: friend or foe in cardiovascular disease? Blood 104, 3231–3232. 183. Barone, L. M., Aronow, M. A., Tassinari, M. T., Conlon, D., Canalis, E., Stein, G. S., and Lian, J. B. (1994). Differential effects of warfarin on mRNA levels of developmentallly regulated vitamin K dependent proteins, osteocalcin and matrix Gla protein in vitro. J. Cellular Physiol. 160, 255–264. 184. Rannels, S. R., Cancela, M. L., Wolpert, E. B., and Price, P. A. (1993). Matrix Gla protein mRNA expression in cultured type II pneumocytes. Am. J. Physiol. 265, L270–L278. 185. Price, P. A., and Williamson, M. K. (1981). Effects of warfarin on bone: Studies on the vitamin K-dependent protein of rat bone. J. Biol. Chem. 256, 12754–12759. 186. Wu, W., Bancroft, J. D., and Suttie, J. W. (1996). Differential effects of warfarin on the intracellular processing of vitamin K-dependent proteins. Thrombosis Haemostasis 76, 46–52. 187. Nishimoto, S. K., and Price, P. A. (1985). The vitamin K-dependent bone protein is accumulated within cultured osteosarcoma cells in the presence of the vitamin K antagonist warfarin. J. Biol. Chem. 260, 2832–2836. 188. Booth, S. L., and Suttie, J. W. (1998). Dietary intake and adequacy of vitamin K. J. Nutr. 128, 785–788. 189. Saxena, S. P., Israels, E. D., and Israels, L. G. (2001). Novel vitamin K-dependent pathways regulating cell survival. Apoptosis 6, 57–68.

Chapter 4

Noncollagenous Proteins; Glycoproteins and Related Proteins Dick Heinegård, Pilar Lorenzo, Tore Saxne

I. Introduction II. Cartilage Extracellular Matrix III. Bone, Extracellular Matrix

IV. Concluding Remarks References

I. INTRODUCTION

extracellular matrix breakdown, it is important to know the organization of the molecules with regard to key elements in interactions. Also, in repair it is important that synthesis is coordinated such that a balance between newly produced molecules is optimal for their assembly into matrix structures. As a background to the understanding of normal matrix biology and pathology in cartilage and bone, this chapter will describe key extracellular macromolecules and what is known about their functional roles. It should be kept in mind that the properties of cartilage and bone are quite different, both with regard to mechanics and with regard to composition. However, at the interface between the cartilage and the bone, it is imperative that molecules can interact between the two tissue structures to provide a tightly interwoven zone that is not sheared off, even by the high stress forces generated in the use of the joints.

Over recent years, development in areas of cartilage and bone biology has provided expanding knowledge on a number of molecular constituents and in some cases also an understanding of functional properties, both with regard to bone and cartilage. Thus, we can view both tissues as composite materials with an organic major extracellular matrix, which provides the tissue properties. In bone this organic matrix actually guides the deposits of mineral, essential for the tissue properties. The processes in the tissue are, however, governed by the cells that have to recognize events like fatigue, with resulting material insufficiency and also have to respond to altered requirements such as modified tissue load. Thus, the cells appear to sense alterations in the tissue via a variety of receptors, often integrins or special cell-surface proteoglycans. To understand degradation of the tissue and consequences of Dynamics of Bone and Cartilage Metabolism

Departments of Experimental Medical Science and Clinical Science, BMC plan C12, SE-22184, Lund, Sweden

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II. CARTILAGE EXTRACELLULAR MATRIX Cartilage matrix, schematically illustrated in Figure 1, consists of fibrillar networks, primarily of collagen II [1–3], collagen VI, and highly negatively charged molecules of aggrecan [4, 5] providing fixed-charged density and therefore an osmotic environment that creates a swelling pressure [4–7]. Also there are a number of noncollagenous glycoproteins that apparently contribute to the regulation of tissue assembly and properties. In some cases these constitute small proteoglycans.

underlying bone. In fact, the charge density of aggrecan is so extreme that there is a very high swelling pressure in the tissue [6, 7] which is resisted by the collagen network as discussed below. The tissue can thus be viewed as a composite of aggrecan containing more than 100 negatively charged chondroitin sulfate side-chains, each with some 50 carboxyl and 50 sulfate groups. The other component then is the fibrillar network consisting of collagen and attached crossbridging and linking noncollagenous as well as collagenous matrix proteins.

B. The Collagen Network A. Aggrecan As discussed in the chapter on proteoglycans aggrecan has a major role in providing fixed-charge groups creating an osmotic environment in the cartilage, thus immobilizing water and restricting water flow. This is of essence for the function of aggrecan in taking up and distributing load over the cartilage tissue and further onto the

The primary functional property of the major network in cartilage based on collagen II is to provide tensional stability. To achieve this, the tissue contains collagen fibers apparently linked together by glycoproteins/proteoglycans. Such linking molecules may function to extend the collagen fibrillar network throughout the tissue. They may also serve to regulate fibrillogenesis by preventing accretion

Figure 1 Illustration of components in the cartilage extracellular matrix. The different molecular organization of territorial (close to cells) and interterritorial (distant from cells) matrix is depicted. Major constituents are the networks based on collagen II and VI, respectively, where collagen fibers contain numerous bound molecules that have roles in regulating assembly and maintaining function of the network. Interactions at the surface of the chondrocyte are likely to have roles in providing cells with information on matrix properties. MAT1,3: Matrilin 1 and/or 3. PRELP: proline, arginine-rich end leucine-rich repeat protein. HS-PG: heparan sulfate proteoglycan. KS: keratan sulfate. CS: chondroitin sulfate. COMP: catilage oligomeric matrix protein. CILP: cartilage intermediate layer protein. HA: hyaluronic acid. CD44: cluster differentiation 44, hyaluronan receptor.

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of new collagen molecules to a pre-existing or forming fibril. Indeed, inactivation of the genes for two of these collagen binding proteins (decorin and lumican) leads to animals having thicker and irregular collagen fibers in several tissues [8, 9]. However, the collagen fibers themselves contain two types of molecules, i.e. collagen II and a few percent of collagen XI, that are closely related and that form the actual fibers [10]. Collagen IX, which represents a more complex collagen consisting of classical triple-helical domains interrupted by globular domains, is bound along the surface of the fibrils [10, 11]. This collagen is largely covalently cross-linked to the collagen II fibers [12]. Of particular interest is that collagen IX may contain a glycosaminoglycan side-chain bound to one of the nontriple-helical domains. Also, the N-terminal part of the α1-chain [13] contains a large globular, rather cationic domain. It has been shown that this extends out from the collagen fibril and would provide a site for interactions with other anionic molecules in the matrix. It is becoming clear that there are collagen fiber subpopulations with different molecules bound at their surface. Indeed recent data indicate that there are fibers that lack collagen IX and others that lack decorin [14], the latter is discussed below. There is a distinct and apparently separate collagen fibrillar system in cartilage made up of collagen VI molecules. The character of these molecules is quite distinct, with globular domains capping the triple-helical structures [15, 16]. They form thinner fibrils that occur predominantly closer to the cells [17]. The function of this network is not clear, but in the tissue it interacts both with the collagen type II fibrillar system and with aggrecan [18]. It may

Figure 2

thus have a role in organizing and connecting the various networks in the tissue.

C. Collagen Associated Molecules There are a number of noncollagenous molecules bound to the collagen fibrils in the tissue. These include decorin, fibromodulin, and lumican, all binding to fibers of collagen [19, 20], but in general not showing a great deal of specificity for a particular type of collagen. These molecules, as is discussed below, have structures of the protein core, suitable for binding to other matrix components. A related molecule, biglycan, appears not to primarily bind collagen II but binds tightly to the N-terminal domain of collagen VI [21]. Decorin, fibromodulin, and lumican are members of a larger family of leucine-rich proteins with homologous structures (see Fig. 2; [22, 23]). These proteins have a central region of leucine-rich repeats, each of some 25 amino acid residues, but somewhat variable in length. There are 10 or 11 such repeats in the proteins present in the extracellular matrix, with an additional three molecules containing a shorter repeat of only six repeats (opticin, osteoglycin and epiphycan/PG-Lb). The repeat region is capped by loop structures containing disulfide bridges. In the N-terminal end there is usually an extension carrying either the dermatan sulfate or chondroitin sulfate side-chains in the case of decorin and biglycan (see [22]) or a domain rich in tyrosine sulfate in the case of fibromodulin [24] with up to nine sulfate residues [25], lumican [26] with at least two and possibly four sulfates [25],

Illustration of LRR-rich proteins in the extracellular matrix.

74 and osteoadherin [27] with up to six sulfates [25]. Keratocan is also likely to contain an N-terminal domain with tyrosine sulfate [28]. Asporin differs in containing a variably long polyaspartate sequence [29]. In PRELP the N-terminal end is rich in proline and arginine [30]. Structural details of decorin representing one subclass of this family of molecules have been elucidated by X-ray crystallography [31]. The molecule contains a central region of 12 repeats, varying in length from 21 to 30 amino acids. Each repeat forms a hairpin loop structure and the whole region is curved to produce a concave surface with the β-sheet structures of each repeat. The outer convex surface shows a less conserved structure. A surprising finding was that the decorin molecules in the crystals appeared as dimers where the two molecules are arranged in an antiparallel fashion with the concave surfaces facing each other. The dimerization domain includes the N-terminal cysteine loop cap structure and extends through some two-thirds of the whole molecule, thus forming a large interactive region. This dimerization, also verified by other techniques [32], has important implications for interactions of this group of molecules with other matrix constituents and previous concepts implicating the concave β-sheet structure, as the one involved in heterotypic interactions may have to be modified. Most of the LRR proteins contain an anionic N-terminal part as discussed above. In the case of PRELP, this extension is basic and can bind heparin and heparan sulfate [33]. PRELP can also bind collagen with high affinity [34]. The protein can actually bind to collagen at the same time as it binds to the heparan sulfate side chains of perlecan [34]. It appears that PRELP has a role in crossbridging tissue structures. One of its locations juxtapositioned to certain basement membranes, e.g. in skin [34], provides clues to this function. Another possible role of PRELP is at the cell surface interacting with heparan sulfate in, for example, syndecan. This interaction may function in regulating cell signals. Indeed it has been reported that PRELP can mediate cell binding [35], possibly involving heparan sulfate. The one known exception among the LRR proteins which contains no N-terminal extension [36] is the integrinbinding chondroadherin [37]. Some of the LRR proteins occur as proteoglycans depending on tissue and species. These are represented by fibromodulin, lumican, osteoadherin, and possibly keratocan, that all may contain one or two keratan sulfate chains, where the particular site in other cases is an N-glycosidically linked oligosaccharide chain [38, 39]. In the C-terminal end, particularly, osteoadherin contains a long extension with a high abundance of acidic amino acids [27] and two tyrosine sulfate residues [25]. Thus, the

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members of this family of extracellular matrix leucinerich repeat proteins include decorin, biglycan, fibromodulin, lumican, keratocan, PRELP, chondroadherin, and osteoadherin, as well as three members with fewer repeats, i.e. opticin [40], osteoglycin, and epiphycan (see [22]), as is illustrated in Figure 2. These highly anionic structures indeed appear to form a second functional domain. Thus, as is illustrated in Figure 1, the protein core of these molecules appears to be bound to one collagen fiber, such that the negatively charged side-chain may participate in interactions with the neighboring fibril, either directly or via positively charged domains, e.g. in the form of the cationic NC4 domain of the α1-chains of collagen IX bound to these fibers. Consequently, it is likely that these interactions will provide a tight, multiple-site linkage between adjacent collagen fibers stabilizing the network and the tissue. The recently observed dimerization may provide for additional crossbridging capabilities. The extensive networking creates a highly tensile-resistant structure, which is likely to be of fundamental importance for the cartilage, since the highly negatively charged domain of the aggrecan molecules creates a swelling pressure that needs to be counteracted, in this case by a stable, stiff fibrillar network. It is conceivable that at some stage in the processes of joint disease, the stability of this network is impaired by proteolytic cleavage of the participating noncollagenous proteins. This may lead to the swelling of the tissue, as is often seen in osteoarthritis. Indeed by IL-1 stimulation of cartilage in explant culture, a fragmentation of fibromodulin by MMP-13 is induced just to the N-terminal side of the first disulfide loop leading to release of almost the entire tyrosine sulfate domain [41]. Another functional property has been ascribed to some members of the LRR family of proteins/proteoglycans. Biglycan, decorin, fibromodulin [42], and more recently also asporin [43] have been shown to be able to bind TGF-β in vitro via their core proteins. The physiological relevance of these interactions is presently not known, although one can speculate on a role in growth, remodeling, and repair by releasing TGF-β. Among the LRR matrix proteins, chondroadherin and osteoadherin have been shown to bind cells. This occurs via the integrins α2β1 for chondroadherin [37] and αvβ3 for osteoadherin [44]. Interestingly, cells binding to chondroadherin do not show spreading and cell division [35, 37] an effect commonly observed when cells bind via this integrin to, for example, collagen. Thus, the molecule may have a role in regulating cell growth in the cartilage matrix. In support, its localization is predominantly close to cells in the pericellular environment [45].

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Asporin is a more recently discovered member of the LRR protein family [29, 46]. The protein was partially purified from human articular cartilage and meniscus [29] and is expressed in a variety of tissues with higher levels in articular cartilage, heart, aorta, uterus and liver [29]. The protein has an extended, variable polyaspartate sequence of up to 15 uninterrupted such residues in the N-terminal extension. Although there are no firm data on function, asporin appears to have the ability to bind to collagen. The production of the protein is up-regulated in osteoarthritis [47]. In a recent publication it was shown that asporin with 14 aspartates N-terminally is overrepresented in Japanese individuals with hip osteoarthritis compared with unaffected individuals, for example, the molecule with 13 aspartates [43]. It is suggested that asporin may modulate TGF-β function which contributes to the development of osteoarthritis [43].

D. Other Cartilage Extracellular Matrix Constituents 1. COMP

A prominent component in cartilage is COMP (Cartilage Oligomeric Matrix Protein). This protein is a homopentamer of five subunits, each with a molecular mass of 87 000 daltons [48–51]. These are joined via a coiled-coil domain near the N-terminal [52] and the interaction is further stabilized by disulfide bonds. The protein has extensive homology to the thrombospondins, but shows a more restricted tissue distribution to cartilage [48] and tendon [53, 54]. It has been shown that COMP interacts with triple helical collagen with a KD of 10 −9 in a Zn2+-dependent manner [55]. Each of the five subunits contains one binding site in the C-terminal globular domain, thus potentially providing the protein with five interaction sites. Therefore, COMP may have a role in stabilizing the collagen network and/or in promoting the collagen fibril assembly. It has furthermore been proposed that COMP may bind chondrocytes [56]. However, the protein is predominantly present in the interterritorial compartment in adult articular cartilage at a distance from the cells [57] and thus a primary role in this tissue appears not to be cell binding. In contrast, the protein is found more abundant close to the cells in the proliferative region of the growth plate [57], where it thus may have a role in cell interactions. Interestingly, there are calcium-binding repeats in COMP (see [55]). These appear to have a role in interactions, since the protein can be extracted from tissues using EDTA [49]. Furthermore, mutations in this domain cause growth abnormalities in pseudo-achondroplasia and

multiple epiphyseal dysplasia [58–60]. These patients have a phenotype which becomes apparent only after birth, and show severe dysplasia and early onset osteoarthritis. One feature of the patients is lax joints, perhaps an effect of a possible function of COMP in the tendons and ligaments. The consequences of this defective Ca2+-binding are not apparent, although it appears that the collagen binding does not depend on Ca2+. On the other hand the chondrocytes show major lamellar deposits containing COMP and also collagen IX [61]. It may be that one consequence of the COMP retention in the cells is that other key matrix proteins are also retained, creating a major molecular defect in the matrix. Despite severe growth disturbances when COMP has been mutated, the null mouse shows no detectable phenotype [62]. It is possible that one or several related molecules compensate for the lack of COMP. Accordingly, COMP is a member of the thrombospondin family, which contains five members, where COMP is number 5. Thrombospondin 1 and 2 contain three subunits somewhat larger than that of COMP, while thrombospondins 3, 4, and 5 contain five subunits. Both thrombospondin 1 and COMP have been identified in cartilage [56], but little is known of their function in the tissue. COMP is also detected in certain other tissues, particularly those experiencing high-pressure load, e.g. certain parts of tendon [54]. There are small amounts of the protein in the synovial capsule, but since the level is about 1:100 of that in synovial fluid [63], it is likely that this protein represents a small proportion, released from cartilage to synovial fluid and then diffusing into the capsule tissue. The synthesis of COMP in the tissue is up-regulated in the early stages of osteoarthritis [47] indicating a role for the protein in repair attempts. 2. Matrilins

Cartilage matrix protein (CMP, Matrilin-1) [64] is the first member of the matrilin family with four known members [65]. These proteins contain von Willebrand Factor A domains (vWFA), in the case of matrilin-1 [66] two such domains in each of the three identical subunits with a molecular mass of around 50 000 Da. The three subunits are held together via a coiled-coil domain. The matrilins also contain one (Matrilin-1) or several (four in Matrilin-3) EGF repeat domains [65, 66]. Matrilin-3 contains only one vWFA domain [67]. Matrilins-1 and -3 are generally restricted to cartilage [67, 68] while Matrilin-2 has a more general distribution. Interestingly, Matrilin-1 has an even further restricted distribution in that it is not found in mechanically loaded tissues like articular cartilage and the intervertebral disk,

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while it is particularly prominent in the tracheal cartilage of older individuals [68]. The protein is present in the more immature cartilage of the femoral head during earlier phases of development and can be seen in the early bone anlagen [69]. However, at the time of the formation of articular cartilage, the protein is no longer found in this part of the tissue [68]. Its role in the cartilage is not clear, but its two vWFA homology domains (only one in Matrilin-3) may mediate the ability of the protein to bind to collagen [18, 70–72]. Furthermore, Matrilin-1 was initially isolated because of its apparent ability to bind to aggrecan [73]. In subsequent studies it has been shown that the protein appears very tightly bound to distinct regions of aggrecan, possibly via a covalent bond [74]. Matrilin-1 can be isolated from cartilage as a mixed polymer that contains subunits of Matrilin-1 as well as -3 [18, 75]. The functional significance of this heterocomplex is not understood. As discussed below, matrilins also interact with biglycan and decorin, important in the collagen VI network interactions [18]. 3. The Collagen VI Network

An important filamentous network present in many weight-bearing tissues is the one containing collagen VI as the basic unit. The unit secreted from the cell is a tetramer, where each molecule (subunit) contains a trimer of three different alpha chains. The central part of the unit is a triple helical region, with globular structures at each end. The N-terminal extension of the alpha 3 chain contains ten vWFA domains forming the major part of this globule of the subunit, while each of the three chains contains two vWFA domains in their C-terminal extension. These subunits are arranged in pairs in an antiparallel fashion such that the N-terminal domains form both ends of the building block [76]. These are forming long filaments and orthogonal structures by interactions via the N-terminal domains in both end-to-end and side-to-side interactions [77], schematically outlined in Figure 1. The formation of this fibril assembly is catalyzed in vitro particularly by biglycan and less efficiently by decorin. This important effect depends on interactions between structures in the N-terminus of the collagen VI and the proteoglycan core protein [21, 77], as well as on its glycosaminoglycan side chains [77]. The proteoglycan is retained with the completed filament [77]. Indeed, collagen VI filamentous structures isolated from tissue actually demonstrated an assembly where biglycan/decorin binds to the collagen VI N-terminal globular domain and the proteoglycan in turn binds matrilin 1, 2, or 3 [18]. The matrilin in turn can bind collagen or aggrecan, probably via another of the subunits, as is illustrated in Figure 1 [18]. These interactions may

have a central role in networking the various structural elements in cartilage and thereby in forming an assembly suitable for sustaining high load. The fact that collagen VI assemblies can be isolated from tissue where the biglycan/matrilin linker binds procollagen, apparently of type II [18], implies a putative role of collagen VI in collagen II fibril assembly. 4. CILP

This cartilage protein [78] is coded by a gene that is transcribed into a large message that includes two proteins. The major N-terminal portion is the 92 kDa protein CILP and the C-terminal portion represents a homolog to porcine nucleotide pyrophospho-hydrolase [79]. It appears that the precursor protein is cleaved in conjunction with its secretion from the cells to form the two proteins [79]. At this time there is no defined function of the CILP, a protein that shares little homology with previously described proteins. Its distribution within the articular cartilage, however, suggests specific functions. Indeed, the CILP is not found equally throughout the cartilage and shows a restricted distribution to the middle–deep portion of the cartilage [78]. This indicates that this part of the tissue has some distinct functional requirements, where the presence of CILP contributes to the unique matrix properties important for meeting this challenge. Furthermore, CILP is markedly increased in both early and late osteoarthritis [47], perhaps an indication of an important role of the protein in repair/remodeling.

E. Other Proteins in Cartilage There are a number of other proteins in cartilage where the information on functional roles is scarce. These include a 39-kDa protein (GP-39), also referred to as YKL-40, found predominantly in the more superficial parts of articular cartilage [80]. This protein is also found in several other tissues. A novel proteoglycan, containing both keratan sulfate and chondroitin sulfate chains is present primarily in the superficial part of the articular cartilage and also produced in, for example, the synovial capsule [81]. This protein has had several names, but recent data indicate that it represents lubricin, a protein originally identified as the major lubricating factor in synovial fluid [82]. Recent work [83] has identified lubricin as the protein, also referred to as CACP, which is mutated in camptodactyly-arthropathycoxa vara-pericarditis syndrome. One function of the protein appears to relate to lubrication at soft tissue interfaces. Fibronectin is present in cartilage and is interestingly up-regulated in early and late osteoarthritis [47, 84].

Chapter 4 Noncollagenous Proteins

The form present in cartilage is distinguished from that present in serum by being differently spliced [85, 86]. Fibronectin appears to have important roles in tissue formation and building. Fibulins 1 and 2 are dimeric, multidomain proteins [87], present in many tissues including cartilage, where fibulin-1 is restricted to the embryonic stages. These proteins interact tightly with the C-terminal lectin homology domain of aggrecan and versican [88, 89] and appear to be able to catalyze crossbridging and network formation also via this end of the proteoglycans thereby enhancing the network formed via the joining of several such molecules via interactions by their N-terminal, G1 domain to hyaluronan [90]. Similarly, a member of the tenascin family present in growing cartilage interacts with this C-terminal lectin homology domain of aggrecan. As a consequence of its six identical subunits it may link several proteoglycan molecules [91]. WARP contains a vWFA domain and is present in cartilage [92]. It appears to represent an oligomeric protein, but details on its function are still lacking.

F. Concluding Remarks The identification of an increasing number of matrix constituents and their characterization has resulted in new tools that can be used in the study of the normal biology of articular cartilage, and also provide means for studies of the pathobiology of the tissue in joint disease. An important future development should be to increase our understanding of the function of the individual proteins in order to correlate alterations in abundance with losses of components of functions in the cartilage. Furthermore, this could potentially also provide a background for future attempts to develop procedures to selectively stimulate their synthesis and thus promote any repair process.

III. BONE, EXTRACELLULAR MATRIX A major feature of bone is its continuous remodeling. Therefore, the molecular constituents of bone will be described in relation to this process. It is particularly important since load causes fatigue and often altered requirements on the tissue architecture. Thus, there is a characteristic bone remodeling cycle, schematically depicted in Figure 3, with one purpose – to remove old, less functional bone and replace this with fresh bone. Overall this remodeling cycle has two phases. In the initial stage bone breakdown is executed by the osteoclasts, which adhere to the bone surface and create a function of a secondary lysosome having an acid pH to dissolve the

77 hydroxyapatite mineral. This structure underneath the cells contains a variety of lysosomal enzymes with capacity to attack, degrade and dissolve the organic matrix (Fig. 3, panels 1–5). The second, rebuilding, phase depends on the recruitment of osteoblasts that lay down an osteoid, which in a tightly regulated fashion becomes mineralized (Fig. 3, panels 6–8). Critical issues for understanding the normal regulation and balance of the various events of this cycle and how this becomes unbalanced in disease include understanding mechanisms for osteoclast recruitment, how these cells become attached to the surface of the mineralized bone and how their activity is regulated. Other central topics concern how osteoblasts or precursor cells are selectively recruited to the location where bone was previously removed by the osteoclasts. How do the osteoblast lay down an initially nonmineralized matrix, osteoid, and what initiates its calcification? Current data on osteoclastic bone breakdown clearly indicate a role also for the osteoblast in this phase. Thus, it appears that the osteoblast or lining cell is stimulated, most likely by local factors, probably including mechanical stress, to secrete proteases with ensuing removal of the thin layer of osteoid that covers the mineralized bone. It is likely that concomitant with this process, which exposes the mineralized bone surface, either cell-binding macromolecules are secreted and bound to this surface or previously deposited such molecules become exposed. Thus, from studies of osteoclast attachment, it is clear that these cells can bind to osteopontin in vitro via their αvβ3 integrin [93]. In support for a role also in vivo, studies of osteoclasts in bone by immuno-electron-microscopy have shown that the αvβ3 integrin is localized in the clear zone [94], i.e. the site where the cells are attached to the underlying mineralized surface. In further support, the osteopontin protein is selectively localized facing this clear zone part of the osteoclast [94]. Other studies [95, 96] have similarly shown location of the α v β 3 integrin to the clear zone area. Corroborating work has shown that osteoclasts in bone of an osteopetrotic rat (ia/ia) do not develop any significant ruffled border resorption region and the surface facing the bone remains as a large clear zone area, especially without development of a resorptive ruffled border region. The α v β 3 integrin is localized to this entire area and osteopontin is found much enriched in the corresponding part of the opposing mineralized bone [97]. The defect appears to block the secretory vesicles from emptying their contents into the extracellular matrix such that they become enriched on the cytosolic side of the plasma membrane. In summary, overwhelming data demonstrate that the only ligand that is localized at the attachment site,

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Figure 3

Schematic illustrations of the bone remodeling cycle. Breakdown is illustrated in panels 1-5 and rebuilding in panels 6-8. Matrix constituents having apparent roles in the various events are indicated. TRAP: tartrate-resistant phosphatase. OPN: osteopontin. PTH: parathyroid hormone. Vit. D3: 1,25-dihydroxyvitamin D3. Panels 1–5 represent the catabolic phase and panels 6–8 represent the rebuilding phase.

osteopontin, will bind to the αvβ3 integrin concentrated in the clear zone attachment area of the osteoclast plasma membrane. Further information on the key role of integrin α v β 3 and its ligand is the fact that bone turnover can be interfered with by adding RGD-integrin-binding peptides [98, 99]. Also, it has been shown that osteopontin can be dephosphorylated by the tartrate-resistant acid phosphatase [100] secreted into the underlying extracellular area by osteoclasts [100, 101]. Such dephosphorylated osteopontin does not promote binding of osteoclasts in vitro [100], potentially representing a release mechanism for these cells. The role of osteoblasts is indicated by the fact that osteoblasts surrounding active osteoclasts have been shown to contain mRNA for osteopontin [102, 103]. It can thus be hypothesized that the cells actually lay down the osteopontin on the mineralized matrix and when the protein becomes exposed it promotes recruitment of the osteoclasts.

Interestingly, more recent data on OPN-null mice have provided additional information. Thus, although these mice are grossly normal and not osteopetrotic [104], they do not develop the rapid osteoporosis upon ovarectomy that is normally seen indicating that the ability to break down bone is compromised [105]. Studies of a different OPN-null mouse reveal that bone does contain more abundant osteoclasts, but the cells develop very little resorptive ruffled border region. It appears that the cells bind to the occasional integrinbinding protein found throughout bone, but in the absence of a high density of osteopontin that can mediate tight attachment, either formation of the seal zone is inappropriate or signaling to accomplish formation of the secondary lysosome is inadequate. The results indicate that the initiation of the bone remodeling cycle is maintained but its propagation is much reduced (Franzén, Reinholt, Önnerfjord, and Heinegård, unpublished data).

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Once the osteoclast has left the site of resorption, the next step is the recruitment of osteoblasts/osteoblast precursors. These cells must find an adhesion molecule at the surface of the “eroded” bone. However, an alternative may be that they secrete their own adhesion molecule, which in turn binds to some structure at the surface of the cells that they are targeted to. It is interesting that a bone surface that previously provided adhesion of the osteoclast, now instead will bind an osteoblast. There are a number of candidate adhesion molecules. Collagen I is abundantly present in the bone and can certainly bind cells via α1β1, α2β1, α3β1 as well as α10β1 and α11β1 integrins [106–109]. Fibronectin, although not particularly abundant at this site [102], would also have a potential for supporting cell binding via a number of integrins [110]. Other molecules that occur at this site in the bone and that are known to bind the integrins are of course osteopontin and BSP (bone sialoprotein) [93, 100] as well as osteoadherin [44]. All these proteins primarily bind cells via αvβ3 integrin, although osteopontin has been shown to bind to other integrins, e.g. α9β1. The osteoblast has the capacity to secrete these proteins [27, 111, 112]. Another alternative is that the particular protein remains present at the eroded bone surface. BSP is a potentially interesting candidate, since it has been shown to be particularly enriched in areas of bone formation [103]. This includes seams and the bone–cartilage interface in the growth plate [103]. Osteoadherin, a bone-specific protein, appears to have the capacity to bind cells [44] via their αvβ3 integrin. Studies of the distribution of this bone protein in the tissue shows its presence throughout the bone, albeit more enriched at the osteochondral junction [113]. In situ hybridization shows that the protein is expressed by osteoblasts at the surface of trabecular bone [114]. It should be emphasized that to date there are no data to indicate whether one of these proteins actually has a primary role in the recruitment of osteoblasts or whether there are yet other proteins that have this primary role. One protein with potential roles in regulating cellular activities, albeit not binding the cells, is osteonectin (or SPARC) [115]. This protein is present in many connective tissues and has several names, e.g. its original name osteonectin [115], BM-40 [116], and SPARC [117]. It is particularly abundant in bone. Studies in vitro have shown roles in modulating cell division and cell migration [118]. Another protein, the multimeric thrombospondin-1, also present in bone [119], has similar activities and may also actually bind cells ([120], for references see [121, 122]). The more detailed role of thrombospondin-1 in bone has to be further elucidated. In recent work, another member of this protein family, thrombospondin-2, was shown to

have a significant role in bone homeostasis [123]. Thus, mice where the thrombospondin-2 gene was inactivated showed alterations in, e.g. the bone mineral density [123]. The next event in the new bone formation is that the osteoblasts lay down an osteoid, nonmineralized matrix, where the major constituent is collagen I, that forms fibers. These fibers contain molecules bound at their surface, i.e. fibromodulin, biglycan, and decorin, as outlined above. It is particularly interesting that both decorin and fibromodulin bind to the gap region [124, 125], which appears to have a role in mineral deposition. Indeed, it is known from work by several groups (for references see [126–128]) that collagen itself can actually support mineralization. Although it is not clear what elicits this mineralization process one may hypothesize that removal of molecules occupying sites in the gap zone may expose previously blocked functions in the collagen. Thus, it is possible that an important event preceding mineralization is removal of, e.g. decorin and fibromodulin. Indeed, in some preliminary experiments we have found (Hultenby, Reinholt, and Heinegård, unpublished data) using immunolocalization at the ultrastructural level, that fibromodulin, while present in the osteoid on the collagen fibers is not found in the mineralized bone. This loss of the protein from the tissue may represent an event preceding the mineralization, but may alternatively represent a secondary phenomenon to changes in the conformation of the collagen fibers by the forming mineral deposits. A molecule that is unique to bone is osteocalcin [129, 130]. This is one of the few proteins that contains a specific post-translational modification in the form of γ-carboxyglutamic acid residues [129]. These residues provide the protein with particular negatively charged clusters which may promote binding to mineral deposits. Indeed, mice in which the gene for osteocalcin has been inactivated show an increased bone formation [131]. Bone also contains numerous other molecules without defined functional properties. Many of these, including α 2 HS [132, 133], are not synthetic products of the bone cells and are not further discussed here.

IV. CONCLUDING REMARKS The increased knowledge on matrix constituents of bone and cartilage has provided an opportunity to develop assays for an increasing number of these molecules. Their continued study is likely to provide important insight into the understanding of the biology of the tissues and also to provide means for the study of events in tissues in normal remodeling as well as in pathological events in disease. This technology also offers opportunities for repeated

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analyses in the monitoring of evolving disease processes and in characterizing the fragments produced. Thus, over recent years the so-called marker technology has developed. This aims at the measurement of, for example, fragments of matrix constituents released from their confinement in tissues as a result of a catabolic process. Quantitative assessment of such fragments has been shown to provide important information on early events leading to cartilage destruction in osteoarthritis [134, 135] as well as in rheumatoid arthritis [136–138] and experimental arthritis [139]. Future work can be anticipated to provide important tools for the diagnosis of disease in the musculoskeletal system, for monitoring the progress of disease and for discerning mechanistic aspects.

References 1. Eyre, D. R. (1991). The collagens of articular cartilage. Sem. Arthirits Rheumatism 21, 2–11. 2. Mayne, R., Van Der Rest, M., Bruckner, P., and Schmid, T. M. (1996). The collagens of cartilage (types II, IX, X and XI) and the type IXrelated collagens of other tissues (types XII and XIV). In “Extracellular Matrix. A Practical Approach” (M. A. Haralson and J. R. Hassell, Eds.), IRL Press, Oxford University Press, Oxford, pp. 33–97. 3. Van Der Rest, M., and Garrone, R. (1991). Collagen family of proteins. FASEB J. 5, 2814–2823. 4. Hardingham, T. E., and Fosang, A. J. (1992). Proteoglycans: many forms and many functions. FASEB J. 6, 861–870. 5. Heinegård, D., Aspberg, A., Franzén, A., and Lorenzo, P. (2002). Glycosylated matrix proteins. In “Connective Tissue and Its Heritable Disorders” 2nd edition (P. M. Royce, and B. Steinmann, Eds.), Wiley-Liss, New York, pp. 271–291. 6. Mow, V., and Setton, L. (1998). Mechanical properties of normal and osteoarthritic articular cartilage. In “Osteoarthritis” (K. D. Brandt, M. Doherty, and L. S. Lohmander, Eds.), Oxford University Press Inc., New York, pp. 108–122. 7. Maroudas, A. (1979). Physicochemical properties of articular cartilage. In “Adult Articular Cartilage” (M. A. R. Freeman, Ed.), Pitman Medical, Turnbridge Wells, pp. 215–290. 8. Chakravarti, S., Magnuson, T., Lass, J., Jepsen, K., LaMantia, C., and Carroll, H. (1998). Lumican regulates collagen fibril assembly: skin fragility and corneal opacity in the absence of lumican. J. Cell Biol. 141, 1277–1286. 9. Danielson, K., Baribault, H., Holmes, D., Graham, H., Kadler, K., and Iozzo, R. (1997). Targeted disruption of decorin leads to abnormal collagen fibril morphology and skin fragility. J. Cell Biol. 136, 729–743. 10. Mendler, M., Eich-Bender, S. G., Vaughan, L., Winterhalter, K. H., and Bruckner, P. (1989). Cartilage contains mixed fibrils of collagen types II, IX, and XI. J. Cell Biol. 108, 191–197. 11. Eyre, D., Wu, J.-J., and Woods, P. (1992). Cartilage specific collagens. Structural studies. In “Articular Cartilage and Osteoarthritis” (K. E. Kuettner, R. Schleyerbach, J. G. Peyron, and V. C. Hascall, Eds.), Raven Press, New York. 12. Wu, J. J., Woods, P. E., and Eyre, D. R. (1992). Identification of crosslinking sites in bovine cartilage type IX collagen reveals an antiparallel type II-type IX molecular relationship and type IX to type IX bonding. J. Biol. Chem. 267, 23007–23014.

13. Muragaki, Y., Nishimura, I., Henney, A., Ninomiya, Y., and Olsen, B. R. (1990). The α1(IX) collagen gene gives rise to two different transcripts in both mouse embryonic and human fetal RNA. Proc. Natl Acad. Sci. 87, 2400–2404. 14. Hägg, R., Bruckner, P., and Hedbom, E. (1998). Cartilage fibrils of mammals are biochemically heterogeneous: differential distribution of decorin and collagen IX. J. Cell Biol. 142, 285–294. 15. Ayad, S., Marriott, A., Morgan, K., and Grant, M. E. (1989). Bovine cartilage types VI and IX collagens. Characterization of their forms in vivo. Biochem. J. 262, 753–761. 16. Chu, M.-L., Zhang, R.-Z., Pan, T.-C., Stokes, D., Conway, D., Kuo, H.-J., Glanville, R., Mayer, U., Mann, K., Deutzmann, R., and Timpl, R. (1990). Mosaic structure of globular domains in the human type VI collagen α3 chain: similarity to von Willebrand factor, fibronectin, actin, salivary proteins and aprotinin type protease inhibitors. EMBO J. 9:2, 385–393. 17. Kielty, C. M., Whittaker, S. P., Grant, M. E., and Shuttleworth, C. A. (1992). Type VI collagen microfibrils: evidence for a structural association with hyaluronan. J. Cell Biol. 118, No. 4, 979–990. 18. Wiberg, C., Klatt, A. R., Wagener, R., Paulsson, M., Bateman, J. F., Heinegård, D., and Mörgelin, M. (2003). Complexes of matrilin-1 and biglycan or decorin connect collagen VI microfibrils to both collagen II and aggrecan. J. Biol. Chem. 278, 37698–37704. 19. Hedbom, E., and Heinegård, D. (1993). Binding of fibromodulin and decorin to separate sites on fibrillar collagens. J. Biol. Chem. 268, 27307–27312. 20. Rada, J., Cornuet, P., and Hassell, J. (1993). Regulation of corneal collagen fibrillogenesis in vitro by corneal proteoglycan (lumican and decorin) core proteins. Exp. Eye Res. 56, 635–648. 21. Wiberg, C., Hedbom, E., Khairullina, A., Lamande, S. R., Oldberg, Å., Timpl, R., Mörgelin, M., and Heinegård, D. (2001). Biglycan and decorin bind close to the n-terminal region of the collagen VI triple helix. J. Biol. Chem. 276, 18947–18952. 22. Hocking, A., Shinomura, T., and McQuillan, D. (1998). Leucine-rich repeat glycoproteins of the extracellular matrix. Matrix Biol. 17, 1–19. 23. Iozzo, R. V. (1999). The biology of the small leucine-rich proteoglycans. Functional network of interactive proteins. J. Biol. Chem. 274, 18843–18846. 24. Antonsson, P., Heinegård, D., and Oldberg, Å. (1991). Posttranslational modifications of fibromodulin. J. Biol. Chem. 266, 16859–16861. 25. Önnerfjord, P., Heathfield, T. F., and Heinegård, D. (2004). Identification of tyrosine sulfation in extracellular leucine-rich repeat proteins using mass spectrometry. J. Biol. Chem. 279, 26–33. 26. Blochberger, T., Vergnes, J.-P., Hempel, J., and Hassell, J. (1992). cDNA to chick lumican (corneal keratan sulfate proteoglycan) reveals homology to the small interstitial proteoglycan gene family and expression in muscle and intestine. J. Biol. Chem. 267, 347–352. 27. Sommarin, Y., Wendel, M., Shen, Z., Hellman, U., and Heinegård, D. (1998). Osteoadherin, a cell-binding keratan sulfate proteoglycan in bone, belongs to the family of leucine-rich repeat proteins of the extracellular matrix. J. Biol. Chem. 273, 16723–16729. 28. Corpuz, L. M., Funderburgh, J. L., Funderburgh, M. L., Bottomley, G. S., Prakash, S., and Conrad, G. W. (1996). Molecular cloning and tissue distribution of keratocan. Bovine corneal keratan sulfate proteoglycan 37A. J. Biol.Chem. 271, 9759–9763. 29. Lorenzo, P., Aspberg, A., Önnerfjord, P., Bayliss, M., Neame, P., and Heinegård, D. (2001). Identification and characterization of asporin-a – novel member of the leucine rich repeat protein family closely related to decorin and biglycan. J. Biol. Chem. 276, 12201–12211. 30. Bengtsson, E., Neame, P. J., Heinegård, D., and Sommarin, Y. (1995). The primary structure of a basic leucine-rich repeat protein, PRELP, found in connective tissues. J. Biol. Chem. 270, 25639–25644. 31. Scott, P. G., McEwan, P. A., Dodd, C. M., Bergmann, E. M., Bishop, P. N., and Bella, J. (2004). Crystal structure of the dimeric protein core

81

Chapter 4 Noncollagenous Proteins

32.

33.

34.

35.

36.

37.

38.

39.

40.

41.

42.

43.

44.

45.

46.

47.

48.

of decorin, the archetypal small leucine-rich repeat proteoglycan. Proc. Natl. Acad. Sci. USA 101, 15633–15638. Scott, P. G., Grossmann, J. G., Dodd, C. M., Sheehan, J. K., and Bishop, P. N. (2003). Light and X-ray scattering show decorin to be a dimer in solution. J. Biol. Chem. 278, 18353–18359. Bengtsson, E., Aspberg, A., Heinegård, D., Sommarin, Y., and Spillmann, D. (2000). The amino-terminal part of PRELP binds to heparin and heparan sulfate. J. Biol. Chem. 275, 40695–40702. Bengtsson, E., Mörgelin, M., Sasaki, T., Timpl, R., Heinegård, D., and Aspberg, A. (2002). The leucine rich repeat protein PRELP binds perlecan and collagens and may function as a basement membrane anchor. J. Biol. Chem. 277, 15061–15068. Sommarin, Y., Larsson, T., and Heinegård, D. (1989). Chondrocytematrix interactions. Attachment to proteins isolated from cartilage. Exp. Cell Res. 184, 181–192. Neame, P. J., Sommarin, Y., Boynton, R. E., and Heinegård, D. (1994). The structure of a 38-kDa leucine-rich protein (chondroadherin) isolated from bovine cartilage. J. Biol. Chem. 269, 21547–21554. Camper, L., Heinegård, D., and Lundgren-Åkerlund, E. (1997). Integrin alpha2beta1 is a receptor for the cartilage matrix protein chondroadherin. J. Cell Biol. 138, 1159–1167. Oldberg, Å., Antonsson, P., Lindblom, K., and Heinegård, D. (1989). A collagen-binding 59-kd protein (fibromodulin) is structurally related to the small interstitial proteoglycans PG-S1 and PG-S2 (decorin). EMBO J. 8, 2601–2604. Plaas, A., Neame, P., Nivens, C., and Reiss, L. (1990). Identification of the keratan sulfate attachment sites on bovine fibromodulin. J. Biol. Chem. 265:33, 20634–20640. Reardon, A. J., Le Goff, M., Briggs, M. D., McLeod, D., Sheehan, J. K., Thornton, D. J., and Bishop, P. N. (2000). Identification in vitreous and molecular cloning of opticin, a novel member of the family of leucinerich repeat proteins of the extracellular matrix. J. Biol. Chem. 275, 2123–2129. Heathfield, T. F., Önnerfjord, P., Dahlberg, L., and Heinegård, D. (2004). Cleavage of fibromodulin in cartilage explants involves removal of the N-terminal tyrosine sulfate-rich region by proteolysis at a site that is sensitive to matrix metalloproteinase-13. J. Biol. Chem. 279, 6286–6295. Hildebrand, A., Romaris, M., Rasmussen, L. M., Heinegård, D., Twardzik, D. R., Border, W. A., and Ruoslahti, E. (1994). Interaction of the small interstitial proteoglycans biglycan, decorin and fibromodulin with transforming growth factor beta. Biochem. J. 302 (Pt 2), 527–534. Kizawa, H., Kou, I., Iida, A., Sudo, A., Miyamoto, Y., Fukuda, A., Mabuchi, A., Kotani, A., Kawakami, A., Yamamoto, S., Uchida, A., Nakamura, K., Notoya, K., Nakamura, Y., and Ikegawa, S. (2005). An aspartic acid repeat polymorphism in asporin inhibits chondrogenesis and increases susceptibility to osteoarthritis. Nat. Genet. 37, 138–144. Wendel, M., Sommarin, Y., and Heinegård, D. (1998). Bone matrix proteins: isolation and characterization of a novel cell-binding keratan sulfate proteoglycan (osteoadherin) from bovine bone. J. Cell Biol. 141, 839–847. Shen, Z., Gantcheva, S., Månsson, B., Heinegård, D., and Sommarin, Y. (1998). Chondroadherin expression changes in skeletal development. Biochem. J. 330 (Pt 1), 549–557. Henry, S. P., Takanosu, M., Boyd, T. C., Mayne, P. M., Eberspaecher, H., Zhou, W., de Crombrugghe, B., Hook, M., and Mayne, R. (2001). Expression pattern and gene characterization of asporin. a newly discovered member of the leucine-rich repeat protein family. J. Biol. Chem. 276, 12212–12221. Lorenzo, P., Bayliss, M. T., and Heinegård, D. (2004). Altered patterns and synthesis of extracellular matrix macromolecules in early osteoarthritis. Matrix Biol. 23, 381–391. Hedbom, E., Antonsson, P., Hjerpe, A., Aeschlimann, D., Paulsson, M., Rosa-Pimentel, E., Sommarin, Y., Wendel, M., Oldberg, Å., and

49.

50.

51.

52.

53.

54.

55.

56.

57.

58.

59.

60.

61.

62.

63.

Heinegård, D. (1992). Cartilage matrix proteins. An acidic oligomeric protein (COMP) detected only in cartilage. J. Biol. Chem. 267, 6132–6136. Mörgelin, M., Heinegård, D., Engel, J., and Paulsson, M. (1992). Electron microscopy of native cartilage oligomeric matrix protein purified from the Swarm rat chondrosarcoma reveals a five-armed structure. J. Biol. Chem. 267, 6137–6141. Oldberg, Å., Antonsson, P., Lindblom, K., and Heinegård, D. (1992). COMP (cartilage oligomeric matrix protein) is structurally related to the thrombospondins. J. Biol. Chem. 267, 22346–22350. Zaia, J., Boynton, R. E., McIntosh, A., Marshak, D. R., Olsson, H., Heinegård, D., and Barry, F. P. (1997). Post-translational modifications in cartilage oligomeric matrix protein. Characterization of the N-linked oligosaccharides by matrix-assisted laser desorption ionization time-of-flight mass spectrometry. J. Biol. Chem. 272, 14120–14126. Malashkevich, V. N., Kammerer, R. A., Efimov, V. P., Schulthess, T., and Engel, J. (1996). The crystal structure of a five-stranded coiled coil in COMP: A prototype ion channel? Science 274, 761–765. DiCesare, P., Hauser, N., Lehman, D., Pasumarti, S., and Paulsson, M. (1994). Cartilage oligomeric matrix protein (COMP) is an abundant component of tendon. FEBS Lett. 354, 237–240. Smith, R. K., Zunino, L., Webbon, P., and Heinegård, D. (1997). The distribution of cartilage oligomeric matrix protein (COMP) in tendon and its variation with tendon site, age and load. Matrix Biol. 16, 255–271. Rosenberg, K., Olsson, H., Mörgelin, M., and Heinegård, D. (1998). Cartilage oligomeric matrix protein shows high affinity zinc-dependent interaction with triple helical collagen. J. Biol. Chem. 273, 20397–20403. DiCesare, P. E., Morgelin, M., Mann, K., and Paulsson, M. (1994). Cartilage oligomeric matrix protein and thrombospondin 1. Purification from articular cartilage, electron microscopic structure, and chondrocyte binding. Eur. J. Biochem. 223, 927–937. Shen, Z., Heinegård, D., and Sommarin, Y. (1995). Distribution and expression of cartilage oligomeric matrix protein and bone sialoprotein show marked changes during rat femoral head development. Matrix Biol. 14, 773–781. Briggs, M. D., Hoffman, S. M. G., King, L. M., Olsen, A. S., Mohrenweiser, H., Leroy, J. G., Mortier, G. R., Rimoin, D. L., Lachman, R. S., Gaines, E. S., Cekleniak, J. A., Knowlton, R. G., and Cohn, D. H. (1995). Pseudoachondroplasia and multiple epiphyseal dysplasia due to mutations in the cartilage oligomeric matrix protein gene. Nat. Gen. 10, 330–336. Deere, M., Sanford, T., Ferguson, H. L., Daniels, K., and Hecht, J. T. (1998). Identification of twelve mutations in cartilage oligomeric matrix protein (COMP) in patients with pseudoachondroplasia. Am. J. Med. Gen. 80, 510–513. Hecht, J. T., Nelson, L. D., Crowder, E., Wang, Y., Elder, F. F. B., Harrison, W. R., Francomano, C. A., Prange, C. K., Lennon, G. G., Deere, M., and Lawler, J. (1995). Mutations in exon 17B of cartilage oligomeric matrix protein (COMP) cause pseudoachondroplasia. Nat. Gen. 10, 325–329. Maddox, B. K., Keene, D. R., Sakai, L. Y., Charbonneau, N. L., Morris, N. P., Ridgway, C. C., Boswell, B. A., Sussman, M. D., Horton, W. A., Bächinger, H. P., and Hecht, J. T. (1997). The fate of cartilage oligomeric matrix protein is determined by the cell type in the case of a novel mutation in pseudoachondroplasia. J. Biol. Chem. 272, 30993–30997. Svensson, L., Aszodi, A., Heinegård, D, Hunziker, E., Fässler, R., and Oldberg, Å. (2002). Cartilage oligomeric matrix protein-deficient mice have normal skeletal development. Mol. Cell Biol. 12, 4366–4371. Skiöldebrand, E., Lorenzo, P., Zunino, L., Rucklidge, G. J., Sandgren, B., Carlsten, J., and Ekman, S. (2001). Concentration of collagen, aggrecan and cartilage oligomeric matrix protein (COMP) in synovial fluid from equine middle carpal joints. Equine Vet. J. 33, 394–402.

82 64. Paulsson, M., and Heinegård, D. (1981). Purification and structural characterization of a cartilage matrix protein. Biochem. J. 197, 367–375. 65. Belluoccio, D., Schenker, T., Baici, A., and Paulsson, M. (1998). Characterization of human Matrilin-3. Genomics 53, 391–394. 66. Argraves, W. S., Deák, F., Sparks, K. J., Kiss, I., and Goetinck, P. F. (1987). Structural features of cartilage matrix protein deduced from cDNA. Proc. Natl Acad. Sci. 84, 464–468. 67. Wagener, R., Kobbe, B., and Paulsson, M. (1997). Primary structure of matrilin-3, a new member of a family of extracellular matrix proteins related to cartilage matrix protein (matrilin-1) and von Willebrand factor. FEBS Lett. 413, 129–134. 68. Paulsson, M., and Heinegård, D. (1982). Radioimmunoassay of the 148-kilodalton cartilage protein. Distribution of the protein among bovine tissues. Biochem. J. 207, 207–213. 69. Franzén, A., Heinegård, D., and Solursh, M. (1987). Evidence for sequential appearance of cartilage matrix proteins in developing mouse limbs and in cultures of mouse mesenchymal cells. Differentiation 36, 199–210. 70. Deak, F., Wagener, R., Kiss, I., and Paulsson, M. (1999). The matrilins: a novel family of oligomeric extracellular matrix proteins. Matrix Biol. 18, 55–64. 71. Tondravi, M. M., Winterbottom, N., Haudenschild, D. R., and Goetinck, P. F. (1993). Cartilage matrix protein binds to collagen and plays a role in collagen fibrillogenesis. Prog. Clin. Biol. Res. 383B, 515–522. 72. Winterbottom, N., Tondravi, M. M., Harrington, T. L., Klier, F. G., Vertel, B. M., and Goetinck, P. F. (1992). Cartilage matrix protein is a component of the collagen fibril of cartilage. Developmental Dynamics 193, 266–276. 73. Paulsson, M., and Heinegård, D. (1979). Matrix proteins bound to associatively prepared proteoglycans from bovine cartilage. Biochem. J. 183, 539–545. 74. Hauser, N., Paulsson, M., Heinegård, D., and Mörgelin, M. (1996). Interaction of cartilage matrix protein with aggrecan. Increased covalent cross-linking with tissue maturation. J. Biol. Chem. 271, 32247–32252. 75. Wu, J. J., and Eyre, D. R. (1998). Matrilin-3 forms disulfide-linked oligomers with matrilin-1 in bovine epiphyseal cartilage. J. Biol. Chem. 273, 17433–17438. 76. Lamande, S. R., Sigalas, E., Pan, T. C., Chu, M. L., Dziadek, M., Timpl, R., and Bateman, J. F. (1998). The role of the alpha3(VI) chain in collagen VI assembly. Expression of an alpha3(VI) chain lacking N-terminal modules N10–N7 restores collagen VI assembly, secretion, and matrix deposition in an alpha3(VI)-deficient cell line. J. Biol. Chem. 273, 7423–7430. 77. Wiberg, C., Heinegård, D, Wenglén, C., Timpl, R., and Mörgelin, M. (2002). Biglycan organizes collagen VI into hexagonal-like networks resembling tissue structures. J. Biol. Chem. 51, 49120–49126. 78. Lorenzo, P., Bayliss, M. T., and Heinegård, D. (1998). A novel cartilage protein (CILP) present in the mid-zone of human articular cartilage increases with age. J. Biol. Chem. 273, 23463–23468. 79. Lorenzo, P., Neame, P., Sommarin, Y., and Heinegård, D. (1998). Cloning and deduced amino acid sequence of a novel cartilage protein (CILP) identifies a proform including a nucleotide pyrophosphohydrolase. J. Biol. Chem. 273, 23469–23475. 80. Volck, B., Ostergaard, K., Johansen, J. S., Garbarsch, C., and Price, P. A. (1999). The distribution of YKL-40 in osteoarthritic and normal human articular cartilage. Scand. J. Rheumatol. 28, 171–179. 81. Schumacher, B. L., Block, J. A., Schmid, T. M., Aydelotte, M. B., and Kuettner, K. E. (1994). A novel proteoglycan synthesized and secreted by chondrocytes of the superficial zone of articular cartilage. Arch. Biochem. Biophy. 311, 144–152. 82. Swann, D. A., Silver, F. H., Slayter, H. S., Stafford, W., and Shore, E. (1985). The molecular structure and lubricating activity of lubricin isolated from bovine and human synovial fluids. Biochem. J. 225, 195–201. 83. Marcelino, J., Carpten, J. D., Suwairi, W. M., Gutierrez, O. M., Schwartz, S., Robbins, C., Sood, R., Makalowska, I., Baxevanis, A.,

Dick Heinegård ET AL.

84.

85.

86.

87.

88.

89.

90.

91.

92.

93.

94.

95.

96.

97.

98.

99.

Johnstone, B., Laxer, R. M., Zemel, L., Kim, C. A., Herd, J. K., Ihle, J., Williams, C., Johnson, M., Raman, V., Alonso, L. G., Brunoni, D., Gerstein, A., Papadopoulos, N., Bahabri, S. A., Trent, J. M., and Warman, M. L. (1999). CACP, encoding a secreted proteoglycan, is mutated in camptodactyly-arthropathy-coxa vara-pericarditis syndrome. Nat. Genet. 23, 319–322. Burton-Wurster, N., and Lust, G. (1985). Deposition of fibronectin in articular cartilage of canine osteoarthritic joints. Am. J. Vet. Res. 46, 2542–2545. Zang, D.-W., Burton-Wurster, N., and Lust, G. (1994). Antibody specific for extra domain B of fibronectin demonstrates elevated levels of both extra domain B(+) and B(-) fibronectin in osteoarthritic canine cartilage. Matrix Biol. 14, 623–633. Burton-Wurster, N., and Lust, G. (1989). Molecular and immunologic differences in canine fibronectins from articular cartilage and plasma. Arch. Biochem. Biophys. 269:1, 32–45. Pan, T.-C., Sasaki, T., Zhang, R.-Z., Fässler, R., Timpl, R., and Chu, M.-L. (1993). Structure and expression of fibulin-2, a novel extracellular matrix protein with multiple EGF-like repeats and consensus motifs for calcium binding. J. Cell Biol. 123, 1269–1277. Aspberg, A., Adam, S., Kostka, G., Timpl, R., and Heinegård, D. (1999). Fibulin-1 is a ligand for the C-type lectin domains of aggrecan and versican. J. Biol. Chem. 274, 20444–20449. Isogai, Z., Aspberg, A., Keene, D. R., Ono, R. N., Reinhardt, D. P., and Sakai, L. Y. (2001). Versican interacts with fibrillin-1 and links extracellular microfibrils to other connective tissue networks. J. Biol. Chem. 277, 4565–4572. Mörgelin, M., Paulsson, M., Malmström, A., and Heinegård, D. (1989). Shared and distinct structural features of interstitial proteoglycans from different bovine tissues revealed by electron microscopy. J. Biol. Chem. 264, 12080–12090. Olin, A. I., Mörgelin, M., Sasaki, T., Timpl, R., Heinegård, D., and Aspberg, A. (2001). The proteoglycans aggrecan and versican form networks with fibulin-2 through their lectin domain binding. J. Biol. Chem. 276, 1253–1261. Fitzgerald, J., Tay, T. S., and Bateman, J. F. (2002). WARP is a new member of the von Willebrand factor A-domain superfamily of extracellular matrix proteins. FEBS Lett. 517, 61–66. Flores, M., Norgård, M., Heinegård, D., Reinholt, F., and Andersson, G. (1992). RGD-directed attachment of isolated rat osteoclasts to osteopontin, bone sialoprotein, and fibronectin. Exp. Cell Res. 201, 526–530. Reinholt, F., Hultenby, K., Oldberg, Å., and Heinegård, D. (1990). Osteopontin – a possible anchor of osteoclasts to bone. Proc. Natl Acad. Sci. USA 87, 4473–4475. Zambonin-Zallone, A., Teti, A., Gaboli, M., and Marchisio, P. C. (1989). Beta 3 subunit of vitronectin receptor is present in osteoclast adhesion structures and not in other monocyte-macrophage derived cells. Conn. Tissue Res. 20, 143–149. Miyauchi, A., Alvarez, J., Greenfield, E. M., Teti, A., Grano, M., Colucci, S., Zambonin-Zallone, A., Ross, F. P., Teitelbaum, S. L., and Cheresh, D. (1991). Recognition of osteopontin and related peptides by an alpha v beta 3 integrin stimulates immediate cell signals in osteoclasts. J. Biol. Chem. 266, 20369–20374. Hultenby, K., Reinholt, F. P., Heinegård, D., Andersson, G., and Marks, S. C., Jr. (1995). Osteopontin: a ligand for the alpha v beta 3 integrin of the osteoclast clear zone in osteopetrotic (ia/ia) rats. Ann. N.Y. Acad. Sci. 760, 315–318. Horton, M. A., Taylor, M. L., Arnett, T. R., and Helfrich, M. H. (1991). Arg-Gly-Asp (RGD) peptides and the anti-vitronectin receptor antibody 23C6 inhibit dentine resorption and cell spreading by osteoclasts. Exp. Cell Res. 195, 368–375. Sato, M., Sardana, M. K., Grasser, W. A., Garsky, V. M., Murray, J. M., and Gould, R. J. (1990). Echistatin is a potent inhibitor of bone resorption in culture. J. Cell Biol. 111, 1713–1723.

Chapter 4 Noncollagenous Proteins 100. Ek-Rylander, B., Flores, M., Wendel, M., Heinegård, D., and Andersson, G. (1994). Dephosphorylation of osteopontin and bone sialoprotein by osteoclastic tartrate-resistant acid phosphatase. Modulation of osteoclast adhesion in vitro. J. Biol. Chem. 269, 14853–14856. 101. Reinholt, F., Mengarelli-Widholm, S., Ek-Rylander, B., and Andersson, G. (1990). Ultrastructural localization of a tartrate-resistant acid ATPase in bone. J. Bone Min. Res. 5:10, 1055–1061. 102. Hultenby, K., Reinholt, F. P., Oldberg, Å., and Heinegård, D. (1991). Ultrastructural immunolocalization of osteopontin in metaphyseal and cortical bone. Matrix 11, 206–213. 103. Hultenby, K., Reinholt, F. P., Norgård, M., Oldberg, Å., Wendel, M., and Heinegård, D. (1994). Distribution and synthesis of bone sialoprotein in metaphyseal bone of young rats show a distinctly different pattern from that of osteopontin. Eur. J. Cell Biol. 63, 230–239. 104. Rittling, S. R., Matsumoto, H. N., McKee, M. D., Nanci, A., An, X. R., Novick, K. E., Kowalski, A. J., Noda, M., and Denhardt, D. T. (1998). Mice lacking osteopontin show normal development and bone structure but display altered osteoclast formation in vitro. J. Bone Miner. Res. 13, 1101–1111. 105. Yoshitake, H., Rittling, S. R., Denhardt, D. T., and Noda, M. (1999). Osteopontin-deficient mice are resistant to ovariectomy-induced bone resorption [published erratum appears in Proc. Natl Acad. Sci. USA 1999 Sep 14;96(19):10944]. Proc. Natl Acad. Sci. USA 96, 8156–8160. 106. Gullberg, D., Gehlsen, K. R., Turner, D. C., Åhlén, K., Zijenah, L. S., Barnes, M. J., and Rubin, K. (1992). Analysis of α1β1, α2β1 and α3β1 integrins in cell–collagen interactions: identification of conformation dependent α1β1 binding sites in collagen type I. EMBO J. 11, 3865–3873. 107. Camper, L., Hellman, U., and Lundgren-Akerlund, E. (1998). Isolation, cloning, and sequence analysis of the integrin subunit alpha10, a beta1-associated collagen binding integrin expressed on chondrocytes. J. Biol. Chem. 273, 20383–20389. 108. Bengtsson, T., Camper, L., Schneller, M., and Lundgren-Åkerlund, E. (2001). Characterization of the mouse integrin subunit alpha10 gene and comparison with its human homologue. Genomic structure, chromosomal localization and identification of splice variants. Matrix Biol. 20, 565–576. 109. Tiger, C. F., Fougerousse, F., Grundstrom, G., Velling, T., and Gullberg, D. (2001). alpha11beta1 integrin is a receptor for interstitial collagens involved in cell migration and collagen reorganization on mesenchymal nonmuscle cells. Dev. Biol. 237, 116–129. 110. Charo, I. F., Nannizzi, L., Smith, J. W., and Cheresh, D. A. (1990). The vitronectin receptor αvβ3 binds fibronectin and acts in concert with α5β1 in promoting cellular attachment and spreading on fibronectin. J. Cell Biol. 111:6, 2795–2800. 111. Oldberg, Å., Franzén, A., and Heinegård, D. (1986). Cloning and sequence analysis of rat bone sialoprotein (osteopontin) cDNA reveals an Arg-Gly-Asp cell-binding sequence. Proc. Natl Acad. Sci. USA 83, 8819–8823. 112. Oldberg, Å., Franzén, A., and Heinegård, D. (1988). The primary structure of a cell-binding bone sialoprotein. J. Biol. Chem. 263, 19430–19432. 113. Ramstad, V. E., Franzén, A., Heinegård, D., Wendel, M., and Reinholt, F. P. (2003). Ultrastructural distribution of osteoadherin in rat bone shows a pattern similar to that of bone sialoprotein. Calcif. Tissue Int. 72, 57–64. 114. Shen, Z., Gantcheva, S., Sommarin, Y., and Heinegård, D. (1999). Tissue distribution of a novel cell binding protein, osteoadherin, in the rat. Matrix Biol. 18, 533–542. 115. Termine, J. D., Kleinman, H. K., Whitson, S. W., Conn, K. M., McGarvey, M. L., and Martin, G. R. (1981). Osteonectin, a bonespecific protein linking mineral to collagen. Cell 26, 99–105.

83 116. Lankat-Buttgereit, B., Mann, K., Deutzmann, R., Timpl, R., and Krieg, T. (1988). Cloning and complete amino acid sequences of human and murine basement membrane protein BM-40 (SPARC, osteonectin). FEBS Lett. 236, 352–356. 117. Mason, I. J., Taylor, A., Williams, J. G., Sage, H., and Hogan, B. L. M. (1986). Evidence from molecular cloning that SPARC, a major product of mouse embryo parietal endoderm, is related to an endothelial cell ‘culture shock’ glycoprotein of Mr 43 000. EMBO J. 5:7, 1465–1472. 118. Sage, H., Vernon, R. B., Funk, S. E., Everitt, E. A., and Angello, J. (1989). SPARC, a secreted protein associated with cellular proliferation, inhibits cell spreading in vitro and exhibits Ca+2-dependent binding to the extracellular matrix. J. Cell Biol. 109, 341–356. 119. Robey, P. G., Young, M. F., Fisher, L. W., and McClain, T. D. (1989). Thrombospondin is an osteoblast-derived component of mineralized extracellular matrix. J. Cell Biol. 108, 719–727. 120. Lawler, J., Weinstein, R., and Hynes, R. O. (1988). Cell attachment to thrombospondin: the role of ARG-GLY-ASP, calcium, and integrin receptors. J. Cell Biol. 107:6, 2351–2361. 121. Bornstein, P. (1992). Thrombospondins: structure and regulation of expression. FASEB J. 6, 3290–3299. 122. Lawler, J., Duquette, M., Urry, L., McHenry, K., and Smith, T. F. (1993). The evolution of the thrombospondin gene family. J. Mol. Evol. 36, 509–516. 123. Kyriakides, T. R., Zhu, Y. H., Smith, L. T., Bain, S. D., Yang, Z., Lin, M. T., Danielson, K. G., Iozzo, R. V., LaMarca, M., McKinney, C. E., Ginns, E. I., and Bornstein, P. (1998). Mice that lack thrombospondin 2 display connective tissue abnormalities that are associated with disordered collagen fibrillogenesis, an increased vascular density, and a bleeding diathesis. J. Cell Biol. 140, 419–430. 124. Pringle, G., and Dodd, C. (1990). Immunoelectron microscopic localization of the core protein of decorin near the d and e bands of tendon collagen fibrils by use of monoclonal antibodies. J. Histochem. Cytochem. 38, 1405–1411. 125. Hedlund, H., Mengarelli-Widholm, S., Heinegård, D., Reinholt, F. P., and Svensson, O. (1994). Fibromodulin – distribution and association with collagen. Matrix 14, 227–232. 126. Glimcher, M. J. (1989). Mechanisms of calcification in bone: Role of collagen fibrils and collagen-phosphoprotein complexes in vitro and in vivo. In “Clinical Impact of Bone and Connective Tissue Markers” (E. Lindh, and J. Thorell, Eds.), Academic Press, London, pp. 137–170. 127. Traub, W., Arad, T., and Weiner, S. (1989). Three-dimensional ordered distribution of crystals in turkey tendon collagen fibers. Proc. Natl Acad. Sci. 86, 9822–9826. 128. McEwen, B. F., Song, M. J., and Landis, W. J. (1991). Quantitative determination of the mineral distribution in different collagen zones of calcifying tendon using high voltage electron microscopic tomography. J. Comp. Assis. Microscopy 3, 201–210. 129. Price, P. A., Otsuka, A. A., Poser, J. W., Kristaponis, J., and Raman, N. (1976). Characterization of a gamma-carboxyglutamic acid-containing protein from bone. Proc. Natl Acad. Sci. USA 73, 1447–1451. 130. Price, P. A., Lothringer, J. W., Baukol, S. A., and Reddi, A. H. (1981). Developmental appearance of the vitamin K-dependent protein of bone during calcification. Analysis of mineralizing tissues in human, calf, and rat. J. Biol. Chem. 256, 3781–3784. 131. Ducy, P., Desbois, C., Boyce, B., Pinero, G., Story, B., Dunstan, C., Smith, E., Bonadio, J., Goldstein, S., Gundberg, C., Bradley, A., and Karsenty, G. (1996). Increased bone formation in osteocalcin-deficient mice. Nature 382, 448–452. 132. Dickson, I. R., Poole, A. R., and Veis, A. (1975). Localisation of plasma α2HS glycoprotein in mineralising human bone. Nature 256, 430–432. 133. McKee, M. D., Nanci, A., and Khan, S. R. (1995). Ultrastructural immunodetection of osteopontin and osteocalcin as major matrix components of renal calculi. J. Bone Miner. Res. 10, 1913–1929.

84 134. Sharif, M., Kirwan, J. R., Elson, C. J., Granell, R., and Clarke, S. (2004). Suggestion of nonlinear or phasic progression of knee osteoarthritis based on measurements of serum cartilage oligomeric matrix protein levels over five years. Arthritis Rheum. 50, 2479–2488. 135. Saxne, T., Heinegård, D., and Månsson, B. (2002). Markers of joint tissue turnover in osteoarthritis. In “The Many Faces of Osteoarthritis” (K. Kuettner, and V. C. Hascall, Eds.), Birkhauser Verlag, Basel, pp. 285–292. 136. den Broeder, A. A., Joosten, L. A., Saxne, T., Heinegard, D., Fenner, H., Miltenburg, A. M., Frasa, W. L., van Tits, L. J., Buurman, W. A., van Riel, P. L., van de Putte, L. B., and Barrera, P. (2002). Long term anti-tumour necrosis factor alpha monotherapy in rheumatoid arthritis: effect on radiological course and prognostic value of markers of

Dick Heinegård ET AL. cartilage turnover and endothelial activation. Ann. Rheum. Dis. 61, 311–318. 137. Crnkic, M., Månsson, B., Larsson, L., Geborek, P., Heinegård, D., and Saxne, T. (2003). Serum cartilage oligomeric matrix protein (COMP) decreases in rheumatoid arthritis patients treated with infliximab or etanercept. Arthritis Res. Ther. 5, R181–R185. 138. Lindqvist, E., Eberhardt, K., Bendtzen, K., Heinegård, D., and Saxne, T. (2005). Prognostic laboratory markers of joint damage in rheumatoid arthritis. Ann. Rheum. Dis. 64, 196–201. 139. Larsson, E., Erlandsson, H. H., Larsson, A., Månsson, B., Saxne, T., and Klareskog, L. (2004). Corticosteroid treatment of experimental arthritis retards cartilage destruction as determined by histology and serum COMP. Rheumatology (Oxford) 43, 428–434.

Chapter 5

Proteoglycans and Glycosaminoglycans Tim Hardingham

Wellcome Trust Centre for Cell-Matrix Research, Faculty of Life Sciences, University of Manchester, Manchester M13 9PT, United Kingdom

V. Leucine-rich Proteoglycans in Cartilage and Bone VI. Perlecan in Cartilage References

I. Introduction II. Glycosaminoglycans III. Proteoglycans in Cartilage IV. Aggrecan

I. INTRODUCTION

(~0.1% w/w). This contrast in composition is also reflected in the proteoglycans present in the tissue. In cartilage the major proteoglycan is aggrecan, which forms supramolecular aggregates and is important in expanding and hydrating the matrix. It contains mainly chondroitin sulfate and increasing amounts of keratan sulfate with age. There also lesser amounts of the lower-molecular-weight leucine-rich proteoglycans, including decorin, fibromodulin, and lumican, which are collagen fibril-associated, and also biglycan. These also contain chondroitin sulfate, but with varying degrees of epimerization to dermatan sulfate. In bone there is no aggrecan and the collagen fibril associated proteoglycans and biglycan are the predominant forms. The cells of bone and cartilage also contain some cell-surface integral membrane proteoglycans, which predominantly contain heparan sulfate. These are very important in cell–matrix interactions and cell signaling, and control many aspects of chondrocyte and osteoblast function, but as the cell density is low they do not contribute to a large fraction of the tissue content of proteoglycans.

Proteoglycans are important components of extracellular matrices (ECM) and have multiple functions that depend on both their protein and carbohydrate constituents. They form a special class of glycoproteins with attached long unbranched and highly charged glycosaminoglycan chains [1, 2]. These chains are strongly hydrophilic and dominate the physical properties of proteoglycans, but the proteins to which they are attached are quite diverse in structure and form several distinct protein families. Proteoglycans are most abundant in those tissues where the ECM is highly hydrated. Cartilage and bone are tissues that both contain large expanded ECM, although the composition of the two tissues are strikingly different. Cartilage contains 70–75% (w/w) water and it has a high fibrillar collagen content (~20% w/w) and also a high proteoglycan content (5–7% w/w). In contrast, bone matrix is mainly mineral, reinforced with fibrillar collagens (~5% w/w), it contains only 10% (w/w) water and it has a correspondingly low proteoglycan content Dynamics of Bone and Cartilage Metabolism

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TIM HARDINGHAM

Figure 1

Glycosaminoglycan linkages to protein.

II. GLYCOSAMINOGLYCANS The glycosaminoglycans are long-chain unbranched polysaccharides consisting of repeating disaccharide units [2] with typically polydisperse chains of average size 10–50 kDa (Figs 1, 2). They form distinct types based on the sugars that form the repeating disaccharide. Chondroitin sulfate/dermatan sulfate are synthesized with a common backbone, which is distinct from that which forms heparin/heparan sulfate. Keratan sulfate has a third different type of disaccharide repeat. Hyaluronan is based on a fourth type of disaccharide repeat and is the only

nonsulfated glycosaminoglycan. It also differs fundamentally from the other glycosaminoglycans, as it is not synthesized attached to a protein as a proteoglycan, but as a long unbranched polysaccharide of exceptional length up to 5 × 106 Da.

A. Chondroitin Sulfate/Dermatan Sulfate In chondroitin sulfate the repeated disaccharide contains D-N-acetylgalactosamine β1–4 D-glucuronate, in β1–3-linked units. There is usually one sulfate group per

Figure 2 Chondroitin sulfate/dermatan sulfate chain structures. Different sulfation of chondroitin sulfate chains and variable epimerization of glucuronate to iduronate in dermatan sulfate chains generates patterns of distinct chain sequence and biological functions.

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disaccharide on the 4 or 6 position of the galactosamine, there is also a low frequency of non-sulfated disaccharides and even more rarely disulfated disaccharides (Fig. 2). The chains are synthesized intracellularly, attached via a characteristic xylose-galactose-galactose linkage region to serine residues on proteoglycans (Fig. 1) by the stepwise addition of single sugar residues from nucleotide sugar intermediates to the non-reducing end of the growing chain, followed by sulfation as the chain grows. This takes place at a late stage of biosynthesis in the Golgi region of the cell immediately prior to secretion. Dermatan sulphate is synthesized as a chondroitin sulfate chain in which some of the glucuronate residues in the chain are epimerized to iduronate intracellularly by an epimerase, which catalyzes the inversion of the configuration of the carboxyl group at carbon 5 of the sugar ring (Fig. 2). This moves it from an equitorial to an axial position and it may also be accompanied by 2-sulfation of the iduronate. The proportion of glucuronate residues that are epimerized to iduronate is quite variable from a few percent up to 90% of the total. The presence of iduronate residues has been shown to alter the chain conformation and different properties have been identified in chains with block structures of glucuronate and iduronate sequences compared with those with more mixed alternating sequences, but how this influences the biological functions of the chains is unclear. Mechanisms controlling the biosynthesis and composition of chondroitin sulfate/dermatan sulfate chains are not well characterized but, as proteoglycans containing chondroitin sulfate and dermatan sulfate can be expressed by the same cells, there are clearly intracellular mechanisms for assembling different GAG chain structures on newly synthesized proteoglycan proteins.

B. Keratan Sulfate Keratan sulfate contains a different disaccharide unit to that of chondroitin sulfate. It consists of D-galactose, β1–3 D-N-acetylglucosamine, in β1–4-linked units. This is also known as a lactosamine repeat and it is found in a broad class of glycoproteins including ovarian mucins, but in keratan sulfate the chain is sulfated during synthesis primarily at the 6 position of the galactose and some N-acetylglucosamine residues are also 6-sulfated. The lactosamine chains that form keratan sulfate are synthesized on two types of protein linkage (Fig. 1). It can be formed as an extension of an oligosaccharide O-linked to threonine, but it can also be synthesized as an extension of an N-linked oligosaccharide of the complex type, attached to asparagine. Both types of keratan sulfate are found in the proteoglycans of cartilage and bone (see below).

C. Disaccharide Sequences in Glycosaminoglycan Chains There is increasing evidence that glycosaminoglycan chains are not randomly sulfated or epimerized polymers and that they contain distinct patterns that may include some unique sequences (Fig. 2). Monoclonal antibodies have been produced that recognize unusual sequences in chondroitin sulfate (CS) chains and these reveal distinctive tissue distributions of the epitopes [3]. The abundance of some CS epitopes has also been shown to change during tissue development and in disease, such as in the cartilage in experimental joint disease [3]. A study of CS epitopes in synovial fluid from joints with trauma-induced ligament damage also showed increased expression of these epitopes [4]. So, although the structures of GAGs are based on repeating disaccharides, the selective sulfation of chondroitin sulfate and keratan sulfate chains and the epimerization of glucuronate to iduronate in dermatan sulfate chains produces chains with wide variations in structure and different properties (Fig. 2). These results also suggest that the cellular synthesis of glycosaminoglycans is much more closely controlled than has been previously suspected and that much has yet to be understood about the significance of the many well-documented changes in GAG composition that occur during tissue development, during aging and that accompany pathology. Further progress in this area depends on the development of improved methods for determining sequences in GAG chains. Some progress has been made in the analysis of glycosaminoglycan composition by using specific enzymes to degrade GAG chains and generate a sensitive quantitative analysis after fluorescent derivatization of the released disaccharides [5, 6]. This technique, FACE (fluorescenceassisted carbohydrate electrophoresis), permits the analysis of glycosaminoglycans in small tissue samples. However, for sequence analysis it requires techniques such as mass spectrometry in combination with specific enzyme or chemical cleavage methods to derive unique solutions to accurate mass determination. This is currently only possible with short oligosaccharides (4–5 sugars), but is likely soon to be applicable to longer oligosaccharides.

III. PROTEOGLYCANS IN CARTILAGE Proteoglycans make a major contribution to the biomechanical properties of cartilage and this is particularly important for articular cartilage as the tissue has a loadbearing function. It is the structure and organization of the macromolecules in the extracellular matrix that provide these properties [7] and they result from the composite

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structure of cartilage and the contribution made by fibrillar and nonfibrillar components. In engineering terms cartilage is a fiber-reinforced composite solid matrix swollen with water (Fig. 3). Cartilage contains several distinct collagens, the most abundant is type II, but there are also types IX, XI, and VI. The collagen fibrillar network is of fine fibers, which have no preferred orientation in the mid zone of the cartilage, although at the articular surface they are parallel to the surface and with one preferred orientation, and they are rather more perpendicular to the surface in deeper zones. The collagen provides a framework for the tissue that gives it shape and form. The triple helical collagen molecules (II, IX, and XI) are organized into fibrils with overlapping and cross-linking of adjacent molecules and the fibrils laterally associate into longer fibers. The structure of collagen gives it impressive tensile properties and these are utilized in cartilage in a special way to produce a tissue that is not only strong in tension, but also resistant to compression. This is achieved by the very high content of proteoglycan, primarily aggrecan, in the interfibrillar matrix. The aggrecan at high concentration draws water into the tissue as it creates a large osmotic swelling pressure (Fig. 3). This occurs because all the negatively charged anionic groups on aggrecan carry with them mobile counter ions such as Na+. This creates a large difference in concentration of ions inside cartilage compared with outside and an imbalance amongst the freely diffusible anions and cations. Water is drawn into the tissue because of this osmotic imbalance and because aggrecan is too large and immobile to redistribute itself. The water thus swells and expands the aggrecan-rich matrix. This places the collagen

Figure 3

network under tension and an equilibrium is achieved when tension in the collagen network balances the swelling pressure, i.e. when no more water enters the tissue because the force is insufficient to stretch the collagen network any further. At this equilibrium, with the tissue swollen with water, it has good compressive resilience, as any new load on the tissue now places the collagen network under further tension. Another feature of the composite collagen/ aggrecan organization is also important, as not only is aggrecan greatly restricted in its ability to move within the matrix and the collagen/aggrecan network is stiff and resists deformation, but aggrecan offers great resistance to any fluid flow and redistribution of water. The tissue thus behaves largely as a stiff elastic polymer to sudden impact loading, but shows some slow inelastic deformation with sustained loads [8]. However, removal of all loads leads to a redistribution of water and a return to the preloading equilibrium position. The articular cartilage thus forms a tough but compliant load-bearing surface and these characteristics depend on the integrity of the collagen network and the retention within it of a high concentration of aggrecan (Fig. 3).

IV. AGGRECAN A. Aggrecan Structure Aggrecan is a proteoglycan with a core protein of high molecular weight (~250 000) encoded by a single gene that is expressed predominantly in cartilaginous tissues [1, 2]. It is a member of a family of four related proteoglycans,

Physical properties of cartilage depend on a combination of the tensile properties of collagen fibers and the osmotic swelling properties of aggrecan. Together they result in a compressively resilient tissue suitable for load bearing.

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Figure 4

The aggrecan family of hyaluronan-binding proteoglycans and the related link protein family. Protein domains with related sequences are identified.

which includes versican with a broad tissue distribution and neurocan and brevican, which are restricted to neural tissues (Fig. 4). Aggrecan has three globular and two extended domains (Figs 4, 5). It is highly glycosylated with ~90% carbohydrate, mainly in two types of glycosaminoglycan chains, chondroitin sulfate and keratan sulfate (Table I). Each aggrecan contains ~100 chondroitin sulfate chains, which are typically ~20 kDa each and the chains are either 4-sulfated, 6-sulfated, or, usually, both. There are fewer keratan sulfate chains (up to 60) and they are usually of shorter length (5–15 kDa). The chondroitin sulfate chains are all attached to the long extended domain between globular domains 2 and 3, but the keratan sulfate

Figure 5

is more widely distributed. It is most abundant in a keratan sulfate-rich region just C-terminal to the G2 domain. They are also attached elsewhere on both extended domains and on the globular domains G1 and G2. Aggrecan also contains a variable number of O-linked oligosaccharides and N-linked oligosaccharides. The O-linked oligosaccharides have a linkage to protein similar to keratan sulfate and it appears that during biosynthesis some O-linked oligosaccharides are extended and sulfated to form keratan sulfate chains, whereas others are not. Variation in the proportion extended to form keratan sulfate and the length of the chains synthesized is thus likely to account for the large differences in the content of keratan sulfate in

Structure of aggrecan and its protein and glycosaminoglycan components.

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TIM HARDINGHAM Table I.

Aggrecan Composition and Polyanionic Properties

Aggrecan Protein Chondroitin sulfate Keratan sulfate

Mol Wt

No of chains

% of total mass

1–4 million 245 000 10 000–30 000 5000–20 000

1 1 100 20–50

100 5–10 70–90 5–20

Polyanion properties: Sulfate groups ~4500 mole per mole. Carboxyl groups ~4200 mole per mole. Total charged groups ~8700 mole per mole.

aggrecan from different cartilaginous tissues and the increase in cartilage that occurs with age. The primary gene product of aggrecan, the protein core, is thus variably glycosylated by the chondrocytes in which it is expressed to give secreted macromolecules that are polydisperse and with a range of composition (Table I). They always contains a high glycosaminoglycan content, but with variable numbers of chains of variable lengths and with some different patterns of sulfation. Although it is clear that many factors affect the synthesis of glycosaminoglycan chains and oligosaccharides, it is not yet fully understood how the composition of aggrecan relates to the age and site of the tissue, or to the range of growth factors and cytokines acting on the chondrocytes in which it is expressed. All three of the globular domains of the aggrecan protein contain sequences that are highly conserved amongst aggrecan from different vertebrate species, but the extended domains are less well conserved [2]. There is, for example, considerable variation in the length of the keratan sulfaterich region amongst different species and even this region has considerable polymorphisms in the human population [9]. This suggests that small variations in the large amount of glycosaminoglycan present on aggrecan are of little consequence to its major function, but that the functions of the globular structures are more sensitive to changes in sequence. Whilst the G1 and G2 domains have related structures (see below), the G3 domain contains four quite different protein modules.

B. Hyaluronan Hyaluronan is a long unbranched glycosaminoglycan based on a disaccharide repeat D-N-acetylglucosamine β1–4 D-glucuronate, in β1–3-linked units. It is made up of many different cell types and is unsulfated and not synthesized attached to protein. There are three mammalian hyaluronan synthases [10] and HAS-2 is most active

in cartilage. The synthase enzyme contains activity to transfer both sugar residues to the growing HA chain and it is able to translocate the chain across the plasma membrane. The mechanism of biosynthesis is quite different from the other glycosaminoglycans, as not only is no protein acceptor required for chain synthesis, but the HA chain grows by the addition of sugars at the reducing terminal end. HA is synthesized by many cells from early in development and deletion of HAS-2 is embryonically lethal in transgenic mice [11]. During cartilage formation as mesenchymal cells differentiate into chondrocytes, HA synthesis is downregulated to about 1% of total glycosaminoglycan synthesis at a level appropriate for extracellular aggregate formation with aggrecan. In aging articular cartilage the HA content steadily rises and can approach 10% of total glycosaminoglycan.

C. The Molecular Interactions Involved in Aggregation The globular N-terminal protein domain (G1) of aggrecan (Fig. 5) contains a lectin-like binding site with high affinity for hyaluronan and is responsible for the formation of aggregates [12]. As hyaluronan is a long unbranched polysaccharide chain, with an average molecular weight of up to several million, each chain can bind a large number of aggrecans to form aggregates up to several hundred million in molecular weight (Fig. 6). The binding of each aggrecan to hyaluronan is further stabilized by a cartilage link protein (45 kDa) [13]. The G1 domain of aggrecan contains three protein motifs, an immunoglobulin fold (Ig-fold) and two copies of a hyaluronan-binding motif, or link module, also referred to as the proteoglycan tandem repeat (PTR) (Fig. 4). This module is present in tandem in all members of the hyaluronan-binding family of proteoglycans and in link protein (Fig. 4), but it is also present as a single copy in the cell-surface hyaluronan-binding receptor, CD-44 and in TSG-6, a secreted matrix protein, whose synthesis is induced by inflammatory cytokines. The structure of the recombinant link module from TSG-6 has now been determined by NMR and shown to be related to the mammalian type-C lectin family of carbohydrate binding motifs [14]. Link protein has a structure very similar to the aggrecan G1 domain as it contains an Ig-fold and two link modules (Fig. 4), which suggests that they share a common gene ancestor. The G2 domain of aggrecan lacks an Ig-fold, but it contains two copies of the link module, as in G1 and link protein. However, it lacks hyaluronanbinding properties [15] and it has no role in the formation of aggregates.

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Figure 6

Electron micrograph of aggrecan aggregate.

The binding of aggrecan to hyaluronan exhibits a high affinity with a Kd of 2 × 10−8 M and it is specific for HA with no significant binding to other glycosaminoglycans. The affinity remains high throughout the pH range 6–9, but it is dissociated at pH 3. At neutral pH the binding is reversibly dissociated at temperatures up to 65°C, but at higher temperatures it is irreversibly denatured. The G1 domain recovers function after exposure to many denaturing agents; guanidine hydrochloride, urea and potassium thiocyanate and solvents, such as ethanol, acetone, or ether. It is also resistant to proteinase attack. There is also recovery of activity after reduction and re-oxidation of disulfide bonds. These properties all suggest that the native structure of the G1 domain is resistant to denaturation and strongly thermodynamically preferred [16]. The G1 domain of aggrecan thus appears to be a tough, stable structure suited to its long lifetime in the extracellular matrix.

D. The Function of Cartilage Link Protein Link protein participates directly in stabilizing the aggregate structure (Figs 4, 6). It forms a ternary complex with aggrecan G1 and HA, with its PTR binding to HA and its Ig-fold interacting with G1 domain. Aggregates formed in the presence of link protein do not dissociate significantly at physiological ionic strength and pH. Thermal stability is also enhanced as no dissociation of link-protein stabilized aggregates is detected up to 55–60°C.

The interaction between the Ig-folds of link protein and G1 in the aggregate structure appears to “lock” aggrecan onto the hyaluronan chain to form the compact ternary unit of the native aggregate structure [17]. Three other members of a link protein gene family have now been identified, all of which contain the same protein modules and, interestingly, each is positioned in the genome closely in tandem with one of the four aggrecan proteoglycan family members [18]. Expression of the cartilage link protein (gene CRTL1) is not restricted to cartilage, but is found in other tissues and it has been shown to be able to stabilize aggregates formed with HA and versican [19]. Mice homozygous for a gene mutation causing CRTL1 deficiency resulted in perinatal lethality and severe skeletal malformation. Many features of this phenotype were able to be rescued by genetically driving CRTL1 expression under a cartilage specific promoter in the mice lacking CRTL1 [20]. These results showed that CRTL1 had a broad pattern of tissue expression and that its function was not replaced by the other link family members. The different biological functions of the other members of the link family have yet to be determined.

E. The Role of Aggregation in Cartilage Structure and Function Aggrecan is synthesized and secreted continuously by chondrocytes (Fig. 7). It follows the same intracellular pathways of synthesis as other secretory proteins [1, 2].

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the G2 domain always remained separate from the G1 domain and away from the hyaluronan. In the presence of link protein the packing density of G1–G2 did not change, but the shadowed protein now appeared as a continuous coating to the central hyaluronan chain and not as discrete globular complexes. This appearance, of a continuous protein coat on the hyaluronan, is also seen in preparations of intact aggregates, which may imply some co-operative association between adjacent G1–link protein complexes along the hyaluronan chain [22]. Figure 7

Extracellular assembly and turnover of aggrecan in cartilage.

G. Functions of the G1–G2 Region in Aggrecan Turnover The mRNA is translated on membrane-bound ribosomes into the RER, followed by translocation to the Golgi for the main steps of O-glycosylation and glycosaminoglycan chain synthesis. The glycosaminoglycans appear to be synthesized rapidly on aggrecan as part of a highly concerted process that occurs just before secretion. There is no intracellular storage of the finished molecule prior to its release. Link protein is also synthesized along the same intracellular pathway. However, hyaluronan is not synthesized within these same compartments within the cell, but is formed by a synthase enzyme located at the plasma membrane. HA is thus synthesized and translocated by an enzyme directly into the extracellular matrix [10]. Aggrecan (and link protein) thus encounter hyaluronan only after their secretion by the chondrocyte into the extracellular matrix. Aggregation is, therefore, an extracellular mechanism for the assembly of aggrecan into higher order structures (Figs 6, 7), which helps immobilize it within the cartilage extracellular matrix, thereby supporting the biomechanical properties of this tissue.

F. Molecular Organization in Aggregates Determination of the stoichiometry of aggregation showed that when hyaluronan was saturated with bound aggrecan the weight ratio was about 1 to 140. This suggests that for each mole of aggrecan (MW 2 million) bound to hyaluronan there was a minimum hyaluronan mass of ~7000. This corresponds to each aggrecan occupying a section of hyaluronan of 32–36 sugars. Electron microscopy of aggregates showed their structure with a large number of aggrecan monomers attached to a central filament of hyaluronan (Fig. 6). In preparations visualized by rotary shadowing [21] the isolated G1 domain bound to hyaluronan with no evidence of co-operativity and the isolated aggrecan G1–G2 fragment bound to hyaluronan;

The extended segment between G1 and G2 appears rather stiffened, as its length in rotary shadowed images was remarkably constant (21 +/− 3 nm) [22]. This segment contains some keratan sulfate, which may contribute to its extended structure. This region also contains the major sites for proteinase attack on aggrecan, which is important in both normal and cytokine-stimulated catabolism by extracellular proteinases [2] (Fig. 7). Specific cleavage sites in this region lead to the search for “aggrecanases”, which were active in aggrecan turnover, and later ADAMTS-4 and -5 were identified as novel proteinases involved in cartilage catabolism [23]. Other metalloproteinase cleavage sites also occur in this region and both types of enzyme appear active in cartilage degeneration in joint pathology. Proteolytic cleavage in the IGD region of aggrecan releases the G2 domain along with the attached large glycosaminoglycan-bearing region from the aggregate. Tight packing of the G1 domain and link protein on hyaluronan in the aggregate structure and their resistance to proteinases leads to a progressive increase in the cartilage with age of hyaluronan associated with aggrecan G1 and link protein. This is readily evident in extracts of old human cartilage, where it has been estimated that only half the G1 domain in the tissue is still part of large aggrecan molecules, with the other half present as truncated molecules lacking the main CS attachment region [24]. It is interesting to note that, in young developing cartilage, the amount of hyaluronan is about 1% of the aggrecan, which is the optimum content to bind all the aggrecan at maximum density into large aggregates. It may also imply that the synthesis and turnover of hyaluronan and aggrecan in cartilage are closely co-ordinated. There is a progressive increase in hyaluronan in cartilage with age to almost 10% of the aggrecan. This increase thus occurs, at least in part, because a large proportion of the hyaluronan is occupied with the accumulated G1 domain fragments of aggrecan and link protein.

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H. G3 Domain The C-terminal G3 domain of aggrecan contains a composite structure of three protein modules (Figs 4, 5). They include a complement regulatory protein domain at the C-terminus, an adjacent mammalian type C-lectin domain and two alternatively spliced epidermal growth factor-like domains. In a nanomelic chick with severe skeletal malformation a primary defect in aggrecan was identified that resulted in a stop codon in the translated region towards G3 and resulted in the expression of a truncated aggrecan that was not glycosylated or secreted [25]. This total deficiency of aggrecan led to a failure in cartilage growth and development. The high conservation of all the motifs in the G3 region amongst aggrecan from different species, and amongst all known members of the aggrecan family, suggested that the G3 domain had interactive properties that may play a role in extracellular matrix organization. The three domains present in G3 are similar to those present in the selectins, where there are specific carbohydrate-binding properties in which the EGF domain appears important for the lectin binding specificity. This raised the question of whether there are conditions under which the alternatively spliced EGF forms are expressed that confer specific interactive properties. Although weak carbohydrate binding of recombinant G3 was detected, other results have shown that the C-lectin domains of all the aggrecan family interact with tenascin-R [26]. This was shown to be a protein–protein interaction with no carbohydrate involvement and binding was with the fibronectin type III repeats in tenascin. The natural ligands for aggrecan G3 domain in cartilage may thus be proteins rather than carbohydrates. This has been substantiated in further studies, which have shown specific calcium-dependent binding of recombinant human aggrecan G3 constructs containing the C-LEC domain to tenascin-C, tenascin-R, fibulin-1, fibulin-2, and PRELP [27]. Those constructs containing both EGF motifs showed particularly high affinity for tenascin-R and fibulin-2, and mRNA transcripts for this alternatively spliced form of aggrecan were detected in RNA from human articular cartilage. The potential for aggrecan G3 domain to interact with other matrix ligands was thus clearly demonstrated with the C-LEC domain providing a site for protein–protein interaction. Preparations of aggrecan from cartilage have been shown by rotary shadowing electron microscopy to contain less than half of the molecules with an intact G3 domain, and the G3 content of articular cartilage falls with age [28]. It is thus clear that in mature tissue much of the function of aggrecan in the matrix does not rely on an intact G3 domain, but its function could be important during cartilage

development or in responses to tissue damage, when the G3 site on aggrecan may help to confer additional stability in the matrix. The glycosaminoglycan attachment region of aggrecan between the G2 and G3 domains is the least well-conserved part of the protein. There are significant differences in the amino acid sequence amongst different species. For example, adjacent to the G2 domain is a keratan sulfate-rich region that contains characteristic hexapeptide repeats; there are 23 in bovine aggrecan, 12 in human and only some poorly conserved repeats in rodent and chicken aggrecans [26]. This results in a large variation in the maximum keratan sulfate content found in aggrecan from these different species. There is also evidence of significant polymorphism in the KS repeats investigated in the human population [9]. Keratan sulfate is also found attached to the protein outside this KS-rich region, but chondroitin sulfate is found exclusively within the CS1/CS2 region between G2 and G3. The CS1 region contains 20 amino acid repeats, of which there are 15 in the rat and 29 in human aggrecan [29] and the CS2 region has repeats of about 100 amino acids, which are more highly variable. These sequence changes result in variations in aggrecan protein molecular weight, for example the protein of human aggrecan is considerably larger than that of the rat (255 kDa compared with 225 kDa).

V. LEUCINE-RICH PROTEOGLYCANS IN CARTILAGE AND BONE A. The Leucine-Rich Family Members of the leucine-rich family of proteoglycans (Table II) are found in most tissues of the body. They are characterized by the inclusion of repeating leucine-rich motifs within the protein sequence structure that are similar to a consensus sequence identified in a broad family of proteins, ranging from mammalian RNAse inhibitor and FSH receptor, through drosophila Toll and chaopterin proteins, to yeast adenylate cyclase [30]. All these family members contain multiple leucine-rich repeats, which vary in number from 3 to 30. The structure of the first member to be crystallized [31], RNAse inhibitor, showed the leucinerich motif to form a β sheet/α helix structure, which packs with the β sheets parallel when several are present to give a curved structure that for RNAse inhibitor formed a doughnut with a bite taken out. Decorin and biglycan were the first leucine-rich proteoglycans to be cloned, but ten other members are now known [30, 32]. They fall into three main subfamilies and several are only sometimes expressed as proteoglycans with attached glycosaminoglycan

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TIM HARDINGHAM Table II. Protein kDa

Class 1

Class 2

Class 3

Others

Decorin Biglycan Asporin Fibromodulin Lumican Keratocan PRELP Osteoadherin Epiphycan Osteoglycin Opticin Chondroadherin

40 40 40 42 38 38 44 42 35 35 35 45

Small Leucine-Rich Proteoglycans LRR

GAG

N-Glyc

S-Tyr

FTGβ

Binding collagen

12 12 12 12 12 12 12 12 7 7 7 10

CS/DS CS/DS – KS KS KS – KS CS/DS KS – –

no no yes yes yes yes no yes yes yes – yes

3 2 – 3–5 3–5 3–5 3–4 3–5 3–4 2–3 1 ?

yes yes ? yes yes yes ? ? ? ? ? ?

yes no ? yes yes ? ? ? no yes yes yes

Classes of ECM proteoglycans and proteins of small leucine-rich repeat family. Properties: Protein Mol Mass (kDa) LRR, number of leucine-rich repeats; GAG, attached glycosaminoglycans; N-Glyc, attached N-glycans; S-Tyr, presence of sulfated tyrosines; TGFβ, binding to TGFβ; binding to fibrillar collagens.

chains (Table II). Decorin and biglycan are present in the matrices of most tissues, including cartilage and bone, although as noted below they have different distribution within tissues. Fibromodulin has a more restricted distribution in cartilage, tendon, and sclera, but it is also present in bone and in skin. Lumican as its name implies is found in cornea, but it is expressed in muscle and intestine and is also significantly expressed in cartilage and bone. Other nonproteoglycan members of the leucine-rich family of proteins have been detected in cartilage. These include osteoadherin, a 42 kDa protein, which has been shown to mediate cell attachment, although not to facilitate cell spreading. PRELP is a proline/arginine-rich end leucine-rich protein, which is closely related to fibromodulin.

B. Decorin and Biglycan Decorin contains a single site for GAG attachment to a serine, four residues from the N-terminus and biglycan has two sites for GAG attachment, which are also both close to the N-terminus and outside the region of leucinerich repeats [33]. The glycosaminoglycan chains attached during biosynthesis are chondroitin sulfate/dermatan sulfate type and the GAG structure varies with the tissue and cell type in which the molecule is synthesized. For example, in bone the chains are chondroitin sulfate and free of iduronate, whereas in skin the chains invariably are dermatan sulfate and contain iduronate; decorin from articular cartilage also contains dermatan sulfate, but in laryngeal cartilage the decorin contains only chondroitin sulfate. The functional significance of this variation in GAG chain structure is not yet explained.

In spite of the close structural relationship between decorin and biglycan there is much to suggest that they have some different functions [30]. Immunolocalization of decorin and biglycan during embryonic development shows that, despite frequently being expressed in the same tissue at the same time, they have contrasting distributions [34]. Decorin is most abundant in matrix, whereas biglycan is mainly pericellular and, where cells mainly express decorin, they poorly express biglycan and vice versa. This suggests that decorin and biglycan differ in the interactions that determine their matrix distribution and their expression is controlled independently. The properties of decorin have been investigated more thoroughly than those of biglycan or of the other leucinerich repeat proteoglycans. Decorin was shown to delay collagen fibril formation in vitro and was shown to bind to collagen fibrils at the d and e region of the gap zone [35]. This was a property of the protein as removal of the GAG chain had no effect on collagen binding. Fibromodulin was subsequently shown also to bind to fibrillar collagen in the gap zone, but at an independent site [36]. In contrast, biglycan appears not to affect collagen fibril formation in vitro and to lack strong affinity for collagen. Based on the crystal structure of ribonuclease inhibitor a model of decorin was proposed, which formed an open ring structure to which a collagen triple helix might bind [37]; however, direct investigation of the X-ray crystal structure of decorin showed a more open structure which formed decorin dimers [38, 39]. The mode of interaction with collagen fibrils has thus yet to be finalized. The effects of decorin on collagen fibril formation were evident in a mouse model in which decorin expression was transgenically disrupted [40]. The major phenotype was a fragile skin with collagen fibers of irregular and abnormal diameter. It is, however, interesting to note that

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the cartilage and bone in these animals showed no gross defects and any consequences of the lack of decorin were either more subtle, or its role in fibril development is compensated by other mechanisms in these tissues. The knockout of biglycan in mice results in a phenotype, with shorter long bones and lower peak bone mass, which lead to an osteoporotic-like condition [41]. They also later developed osteoarthritis, which may result from tendon complications. The basis of the low bone mass appeared to result from fewer osteoblasts and a progressive decline in bone marrow stem cell numbers with age. In contrast to the in vitro results, which showed that biglycan did not affect collagen fibrillogenesis, type I collagen fibril structure was abnormal in the biglycan-deficient animals, but it is unclear if this was a direct or indirect effect. The generation of mice lacking both decorin and biglycan showed a more severe phenotype, with reduced cortical and trabecular bone and fragile skin [41]. The morphology of collagen fibrils was very abnormal, particularly in bone. The results suggested that decorin and biglycan share some functions and in single knockouts they can substitute for each other, but only to a limited extent. Decorin, biglycan, and fibromodulin have been shown to bind TGFβ with both high affinity (Kd 1–20 nM) and lower affinity sites [42]. The interaction was with the protein and removal of the GAG chain increased binding. The interaction was strongest with active TGFβ dimer and was weak with the inactive latent precursor molecule. Both bone and cartilage are known to contain TGFβ bound in their matrices and the leucine-rich proteoglycans may provide sites for their localization. As TGFβ stimulates matrix protein production by chondrocytes and osteoblasts, a matrix store of this factor that might be released at times of matrix damage could be part of a mechanism for an early response to stimulate cells for tissue repair. Bone matrix decorin has been reported to bind TGFβ and enhance its inhibitory effect on the proliferation of osteoblastic cells [43] and on monocytes, but in other systems TGFβ induction of biglycan synthesis was inhibited by decorin [44]. There are therefore potential ways in which decorin, biglycan, and fibromodulin may act both directly and through TBFβ to modulate cell differentiation and proliferation, which may explain some of the range of abnormalities in animals in which their expression is deleted. The properties identified for decorin therefore suggest a whole range of ways in which it may contribute to matrix organization and it may also help to regulate factors that control cell function. Some of these properties are shared with biglycan. Investigation of the expression of decorin and biglycan in different tissues during embryogenesis by in situ hybridization reveals an interesting pattern where

frequently both are expressed in the same cells, but the strong expression of one is accompanied by weak expression of the other [34]. In the developing limb cartilage, biglycan was in the outer rim of cartilage, which lacked decorin, whereas the central region of cartilage was more rich in matrix and contained decorin and lacked biglycan. The surrounding noncartilaginous tissue contained biglycan, but lacked decorin. As development proceeded, biglycan was enriched in the territorial capsule of chondrocytes, whereas decorin was predominantly expressed in the interterritorial matrix. In mature cartilage, the content of biglycan decreased with age, whereas that of decorin increased and biglycan expression was stimulated by TGFβ, but decorin was inhibited [45]. There is much evidence to show that these structurally related proteoglycans have some different tissue functions in the extracellular matrix.

C. Fibromodulin Fibromodulin contains leucine-rich repeat sequences similar to decorin and biglycan, but it contains no ser-gly GAG attachment sequence. However, it does contain four N-linked oligosaccharide acceptor sequences and it has been shown that these can be substituted with N-linked keratan sulfate chains [46]. These site are within the leucine-rich repeat region, but there is also a tyrosine-rich sequence close to the N-terminus which can be sulfated. Fibromodulin thus has a high anionic charge even though it carries no CS/DS GAG chain. Fibromodulin is expressed in early development in both chondrocytes and osteoblasts and in mature tissue is particularly abundant in tendon. Fibromodulin knockout mice have abnormal and fewer collagen fibrils and showed ectopic tendon calcification and there was also a compensatory up-regulation of lumican in tissues such as tail tendon [46]. This appeared to correspond to the fact that both fibromodulin and lumican bind to the same site on fibrillar collagen, which is an independent site to decorin [36].

D. Lumican Lumican is a leucine-rich repeat proteoglycan that is present in cornea, but is only weakly expressed in cartilage and bone. It is most similar in sequence to fibromodulin, it also contains N-linked keratan sulfate, but lacks the tyrosine-rich sequence. It binds to fibrillar collagen similarly to fibromodulin as noted above [47]. In the lumican/ fibromodulin knockout mouse there was a more pronounced tendon phenotype, where lumican absence caused early

96

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defects and fibromodulin absence caused them to persist, but there were no major bone/cartilage abnormalities.

E. Other Leucine-Rich Family Members and Nonglycanated Forms Investigation of tissue extracts from cartilage [48] and intervertebral disk [49] have revealed leucine-rich repeat proteoglycans lacking the glycosaminoglycan chains. This may arise because in decorin and biglycan the N-terminal peptide, which includes the site of GAG attachment, has been shown to be sensitive to mild proteolytic cleavage. The removal of the peptide with the attached GAG chain could thus occur as a result of proteolytic action in the extracellular matrix. However, molecules may also be synthesized and secreted lacking GAG chains and this may be controlled during cell expression. More detailed analysis of structure and biosynthesis is required to show which of these alternative explanations is the major cause of the nonglycanated forms and also if they have any functional significance.

VI. PERLECAN IN CARTILAGE Perlecan is a high-molecular-weight, multidomain protein, which contains heparan sulfate and chondroitin sulfate chains [50]. It was first identified as a component of the matrix in a basement membrane-producing tumor [50, 51]. In rotary shadowed EM perlecan appeared as an elongated core protein containing globular domains with glycosaminoglycan chains attached to one end. Sequence analysis revealed five distinct domains containing tandem arrays of seven different protein motifs. Immunohistochemical studies demonstrated that perlecan was present in all basement membranes and was also located in other stromal ECMs such as the dermis of the skin and in the pericellular region and matrix of cartilage [51–53]. A major site of GAG attachment in perlecan was shown to be domain I, but domain V also has GAG chains. Perlecan was thought to be an obligate HS proteoglycan, but from cartilage it is a composite proteoglycan carrying both HS and CS chains. Part of the phenotype of perlecannull mice, generated by homologous recombination, was disruption of skeletal development in which the cartilage showed disorganized collagenous components and a reduction in content of the major proteoglycan, aggrecan [52, 53]. Mutations in the human perlecan gene have been identified in Schwartz–Jampel syndrome with characteristic cartilage abnormalities [54]. Furthermore, the addition of exogenous perlecan to cultured chondrocytes has been

reported to promote the re-expression of the chondrogenic phenotype [55]. In cartilage, the pericellular localization of much of the perlecan means that it is strategically positioned to participate in extracellular growth factor and cytokine signals acting on chondrocytes to regulate their metabolism [51], but precise mechanisms have yet to be determined. Perlecan has a number of signaling functions in other tissues affecting cell proliferation and angiogenesis and it has been implicated in the regulation of bone formation [56].

References 1. Hardingham, T. E., and Fosang, A. J. (1992). Proteoglycans: many forms, many functions. FASEB J. 6, 861–870. 2. Fosang, A. J., and Hardingham, T. E. (1996). Matrix proteoglycans. In “Extracellular Matrix” (W. D. Comper, Ed.), Vol 2, pp 200–229, Harwood Academic Publishers, Netherlands. 3. Caterson, B., Mahmoodian, F., Sorrell, J. M., Hardingham, T. E., Bayliss, M. T., Carney, S. L., Ratcliffe, A., and Muir, H. (1990). Modulation of native chondroitin sulphate structure in tissue development and in disease. J. Cell. Science 97, 411–417. 4. Hazell, P. K., Dent, C., Fairclough, J. A., Bayliss, M. T., and Hardingham, T. E. (1995). Changes in glycosaminoglycan epitope levels in knee joint fluid following injury. Arthr. Rheum. 38, 953–959. 5. Calabro, A., Benavides, M., Tammi, M., Hascall, V., and Midura, R. (2000). Microanalysis of enzyme digests of hyaluronan and chondroitin/ dermatan sulfate by fluorophore-assisted carbohydrate electrophoresis (FACE). Glycobiology 10, 273–281. 6. Calabro, A., Hascall, V., and Midura, R. (2000). Adaptation of FACE methodology for microanalysis of total hyaluronan and chondroitin sulfate composition from cartilage. Glycobiology 10, 283–293. 7. Hardingham, T. E. (2004). Articular cartilage. In “Oxford Textbook of Rheumatology, 3rd Edition” (D. A. Isenberg, P. J. Maddison, P. Woo, D. N. Glass, F. C. Breedfeld, Eds), pp 325–334, Oxford Medical Publications, Oxford, UK. 8. Mow, V. C., Hou, J. S., Owens, J. M., and Ratcliffe, A. (1990). Biphasic and quasilinear viscoelastic theories for hydrated soft tissues. In Biomechanics of Diarthrodial Joints, (V. C. Mow, A. Ratcliffe, and S. L-Y Woo, Eds), pp 215–260, Springer-Verlag, New York. 9. Doege, K. J., Coulter, S. N., Meek, L. M., Maslen, K., and Wood, J. G. (1997). A human-specific polymorphism in the coding region of the aggrecan gene. Variable number of tandem repeats produce a range of core protein sizes in the general population. J. Biol. Chem. 272, 13974–13979. 10. Spicer, A. P., and McDonald, J. A. (1998). Characterization and molecular evolution of a vertebrate hyaluronan synthase gene family. J. Biol. Chem. 272, 1923–1932. 11. Camenisch, T. D., Spicer, A. P., Brehm-Gibson, T., Biesterfeldt, J., Augustine, M. L., Calabro, A., Kubalak, S., Klewe, S. E., and McDonald, J. A. (2000). Disruption of hyaluronan synthase-2 abrogates normal cardiac morphogenesis and hyaluronan-mediated transformation of epithelium to mesenchyme, J. Clin. Invest. 106, 349–360. 12. Hardingham, T. E., and Muir, H. (1972). The specific interaction of hyaluronic acid with cartilage proteoglycans. Biochim. Biophys. Acta 279, 401–405. 13. Hardingham, T. E. (1979). The role of link-protein in the structure of cartilage proteoglycans aggregates. Biochem. J. 177, 237–247. 14. Kohda, D., Morton, C. J., Parkar, A. A., Hatanaka, H., Inagaki, F. M., Campbell, I. D., and Day, A. J. (1996). Solution structure of the link

Chapter 5 Proteoglycans and Glycosaminoglycans

15.

16. 17.

18.

19.

20.

21.

22.

23. 24.

25.

26.

27.

28.

29.

30.

31.

32.

module: a hyaluronan-binding domain involved in extracellular matrix stability and cell migration. Cell 86, 767–775. Fosang, A. J., and Hardingham, T. E. (1989). Isolation of the N-terminal globular domains from cartilage proteoglycan: identification of G2 domain and its lack of interaction with hyaluronate and link protein. Biochem. J. 261, 801–809. Hardingham, T. E. (1981). Proteoglycans: their structure, interactions and molecular organization in cartilage. Biochem. Soc. Trans. 9, 489–497. Ratcliffe, A., and Hardingham, T. E. (1983). Cartilage proteoglycan binding and link protein: radioimmunoassays and the detection of masked determinants in aggregates. Biochem. J. 213, 371–378. Spicer, A. P., Joo, A., Bowling, R. A. (2003). A hyaluronan binding link protein gene family whose members are physically linked adjacent to chrondroitin sulfate proteoglycan core protein genes – The missing links. J. Biol. Chem. 278, 21083–21091. Shi, S. L., Grothe, S., Zhang, Y. P., O’Connor-McCourt, M. D., Poole, A. R., Roughley, P. J., Mort, J. S. (2004). Link protein has greater affinity for versican than aggrecan J. Biol. Chem. 279, 12060–12066. Czipri, M., et al. (2003). Genetic rescue of chondrodysplasia and the perinatal lethal effect of cartilage link protein deficiency. J. Biol. Chem. 278, 39214–39223. Morgelin, M., Paulsson, M., Hardingham, T. E., Heinegard, D., and Engel, J. (1988). Cartilage proteoglycans: assembly with hyaluronate and link protein as studied by electron microscopy. Biochem. J. 253, 175–185. Morgelin, M., Paulsson, M., Heinegard, D., Aebi, U., and Engel, J. (1995). Evidence of a defined spatial arrangement of hyaluronan in the central filament of cartilage proteoglycan aggregates. Biochem. J. 307, 595–601. Nagase, H., Kashiwagi, M. (2003). Aggrecanases and cartilage matrix degradation. Arthritis Res. Ther. 5, 94–103. Hardingham, T. E., and Bayliss, M. T. (1990). Proteoglycans of articular cartilage: changes in ageing and joint disease. Semin. Arthritis Rheum. 20, 12–33. Vertel, B. M., Walters, L. M., Grier, B., Maine, N., and Goetinck, P. F. (1993). Nanomelic chondrocytes synthesize, but fail to translocate, a truncated aggrecan precursor. J. Cell. Science 104, 939–948. Asberg, A., Miura, R., Bourdoulous, S., Shimonaka, M., Heinegard, D., Schachner, M., Ruoslahti, E., and Yamaguchi, Y. (1997). The C-type lectin domains, a family of aggregating chondroitin sulfate proteoglycans, bind tenascin-R by protein-protein interactions independent of carbohydrate moiety. Proc. Natl Acad. Sci. USA 94, 10116–10121. Day, J. M., Olin, A. I., Murdoch, A. D., Canfield, A., Sasaki. T., Timpl, R., Hardingham, T. E., and Aspberg, A. (2004). Alternative splicing of aggrecan G3 domain influences interactions with tenascin-C and other extracellular matrix proteins. J. Biol. Chem. 279, 12511–12518. Dudhia, J., Davidson, C. M., Wells, T. M., Vynios, D. H., Hardingham, T. E., Bayliss, M. T. (1996). Age related changes in the content of the carboxy-terminal region of aggrecan in human articular cartilage. Biochem. J. 313, 933–940. Doege, K. J., Sasaki, M., Horigan, E., Hassell, J. R., and Yamada, Y. (1991). Complete coding sequence and deduced primary structure of the human large aggregating proteoglycan, aggrecan. Human specific repeats and alternatively spliced forms. J. Biol. Chem. 266, 894–902. Iozzo, R. V. (1997). The family of the small leucine-rich proteoglycans: key regulators of matrix assembly and cell growth. Crit. Rev. Biochem. Mol. Biol. 32, 141–174. Kobe, B., and Deisenhofer, J., (1995). A structural basis for the interactions between leucine-rich repeats and protein ligands. Nature (London) 374, 183–186. Ameye, L., Young, M. F. (2002). Mice deficient in small leucine-rich proteoglycans: novel in vivo models for osteoporosis, osteoarthritis, Ehlers-Danlos syndrome, muscular dystrophy and corneal diseases. Glycobiology 12, 107R–116R.

97 33. Fisher, L. W., Termine, J. D., and Young, M. F. (1989). Deduced protein sequence of bone small proteoglycan I (biglycan) shows homology with proteoglycan II (decorin) and several non-connective tissue proteins in a variety of species. J. Biol. Chem. 264, 4571–4576. 34. Bianco, P., Riminucci, M., Young, M. F., Termine, J. D., and Robey, P. G. (1990). Expression and localization of the two small proteoglycans biglycan and decorin in developing human skeletal and non-skeletal tissues. J. Histochem. Cytochem. 38, 1549–1563. 35. Pringle, G. A., and Dodd, C. M. (1990). Immunoelectron microscopic localization of the core protein of decorin near the d and e bands of tendon collagen fibrils by the use of monoclonal antibodies. J. Histochem. Cytochem. 38, 1405–1411. 36. Hedbom, H., and Heinegard, D. (1993). Binding of fibromodulin and decorin to separate sites on fibrillar collagens. J. Biol. Chem. 268, 27307–27312. 37. Weber, I. T., Harrison, R. W., and Iozzo, R. V. (1997). Model structure of decorin and implications for collagen fibrillogenesis. J. Biol. Chem. 271, 31767–31770. 38. Scott, P. G., Grossmann, J. G., Dodd, C. M., Sheehan, J. K., Bishop, P. N. (2003). Light and X-ray scattering show decorin to be a dimer in solution J. Biol. Chem. 278, 18353–18359. 39. Scott, P. G., McEwan, P. A., Dodd, C. M., Bergmann, E. M., Bishop, P. N., Bella, J. (2004). Crystal structure of the dimeric protein core of decorin, the archetypal small leucine-rich repeat proteolycan. Proc. Natl Acad. Sci. 101, 15633–15638. 40. Danielson, K. G., Baribault, H., Holmes, D. F., Graham, H., Kadler, K. E., and Iozzo, R. V. (1997). Targetted disruption of decorin leads to abnormal collegen morphology and skin fragility. J. Cell. Biol. 136, 729–743. 41. Ameye, L., Aria, D., Jepsen, K., Oldberg, A., Xu, T., and Young, M., (2002). Abnormal collagen fibrils in tendons of biglycan/fibromodulin deficient mice lead to gait impairment, ectopic ossification and osteoarthritis. FASEB J., 16, 673–680. 42. Hildebrand, A., Romaris, M., Rasmussen, L. M., Heinegard, D., Twardzik, D. R., Border, W. A., and Ruoslahti, E. (1994). Interaction of the small interstitial proteoglycans biglycan, decorin and fibromodulin with transforming growth factor β. Biochem. J. 302, 527–534. 43. Takeuchi, Y., Kodama, Y., and Matsumoto, T. (1994). Bone matrix decorin binds transforming growth factor-b and enhances its bioactivity. J. Biol. Chem. 269, 32634–32638. 44. Hausser, H., Groning, A., Haslik, A., Schonherr, E., Kresse, H. (1994). Selective inactivity of TGF-β/decorin complexes. FEBS Lett. 353, 243–245. 45. Roughly, P. J., Melching, L. I., Recklies, A. D. (1994). Changes in the expression of decorin and biglycan in human articular cartilage with age and regulation by TGF-β. Matrix. Biol. 14, 51–59. 46. Plaas, A. H. K., Neame, P. J., Nivens, C. M., and Reiss, L. (1990). Identification of the keratan sulfate attachment sites on bovine fibromodulin. J. Biol. Chem. 265, 20634–20640. 47. Ezura, Y., Chakravarti, S., Oldberg, A., Chervoneva, I., and Birk, D. E. (2000). Differential expression of lumican and fibromodulin regulate collagen fibrillogenesis in developing mouse tendons. J. Cell Biol. 151, 779–787. 48. Sampaio, L de O., Bayliss, M. T., Hardingham, T. E., and Muir, H. (1988). Dermatan sulphate proteoglycan from human articular cartilage. Biochem. J. 254, 757–764. 49. Johnstone, B., Markopoulos, M., Neame, P., and Caterson, B. Identification and characterization of glycanated and non-glycanated forms of biglycan and decorin in the human intervertebral disc. Biochem. J. 292, 661–666. 50. Iozzo, R. V., Cohen, I. R., Grassel, S., and Murdoch, A. D. (1994). The biology of perlecan: the multifaceted heparan sulphate proteoglycan of basement membranes and pericellular matrices. Biochem. J. 302, 625–639. 51. Iozzo, R. V., and Murdoch, A. D. (1996). Proteoglycans of the extracellular environment: clues from the gene and protein side offer novel

98 perspectives in molecular diversity and function. FASEB J. 10, 598–614. 52. Costell, M., Gustafsson, E., Aszodi, A., Morgelin, M., Bloch, W., Hunziker, E., Addicks, K., Timpl, R., and Fassler, R. (1999). Perlecan maintains the integrity of cartilage and some basement membranes. J. Cell Biol. 147, 1109–1122. 53. Arikawa-Hirasawa, E., Watanabe, H., Takami, H., Hassell, J. R., Yamada, Y. (1999). Perlecan is essential for cartilage and cephalic development. Nature Genet. 23, 354–358. 54. Nicole, S., Davoine, C-S., Topaloglu, H., Cattolico, L., Barral, D., Beighton, P., Ben Hamida, C., Hammouda, H., Cruaud, C., White, P. S., Samson, D., Urtizberea, J. A., Lehmann-Horn, F., Weissenbach, J.,

TIM HARDINGHAM Hentati, F., and Fontaine, B. (2000). Perlecan, the major proteoglycan of basement membranes, is altered in patients with Schwartz-Jampel syndrome (chondrodystrophic myotonia). Nature Genet. 26, 480–483. 55. French, M. M., Smith, S. E., Akanbi, K., Sanford, T., Hecht, J., Farach-Carson, M. C., and Carson, D. D. (1999). Expression of the heparan sulfate proteoglycan, perlecan, during mouse embryogenesis and perlecan chondrogenic activity in vitro. J. Cell Biol. 145, 1103–1115. 56. Mongiat, M., Fu, J., Oldershaw, R., Greenhalgh, R., Gown, A. M., Iozzo, R. V. (2003). Perlecan protein interacts with extracellular matrix protein 1 (ECM1) a glycoprotein involved in bone formation and angiogenesis. J. Biol. Chem. 278, 17491–17499.

Chapter 6

Growth Factors Philippa Hulley, Graham Russell Peter Croucher

Academic Unit of Bone Biology, University of Sheffield Medical School, Sheffield S10 2RX, United Kingdom

V. Wnts VI. Additional Growth Factors VII. Summary References

I. Introduction II. Insulin-like Growth Factors III. The Transforming Growth Factor-Beta/Bone Morphogenetic Protein Superfamily IV. Fibroblast Growth Factors

I. INTRODUCTION

and BMP7, have already been released for use in local treatment of fractures and lumbar interbody spinal fusion [1]. The major skeletal growth factors include the insulin-like growth factors (IGF), members of the transforming growth factor-beta (TGF-β)/ΒΜP superfamily, the fibroblast growth factors (FGF), the platelet-derived growth factors (PDGF), members of the epidermal growth factor family (EGF), and the Wnts, and will be discussed in this chapter.

Bone and cartilage growth factors are secreted proteins that are generally produced locally in the skeletal microenvironment and act in a paracrine or autocrine fashion to regulate growth and repair. They contribute to growth not only by triggering mitosis but also by driving skeletal cells towards a committed phenotype, by enhancing differentiated function and, in some cases, protecting against apoptosis. Growth factors can be stored bound to extracellular matrix components, serving as a long-term, stable repository until released by normal matrix remodeling or damage. There are complex interactions between signaling pathways activated by different classes of growth factors, and these can vary widely in embryonic patterning compared to adult remodeling and repair. Likewise, cartilage can respond in a fundamentally different way to growth factors than bone. Thus, while there is obvious appeal in the current enthusiasm for harnessing growth factors as bone and cartilage anabolics, much careful groundwork needs to be done first. In fact, the first growth factors to be licensed as bone anabolics, bone morphogenetic protein 4 (BMP4) Dynamics of Bone and Cartilage Metabolism

Botnar Research Centre, Institute of Musculoskeletal Sciences, University of Oxford, Oxford OX3 7LD, United Kingdom

II. INSULIN-LIKE GROWTH FACTORS The insulin-like growth factors-I and -II (IGF-I and IGF-II) are polypeptide hormones structurally related to insulin. IGF-I, formerly known as somatomedin C, is synthesized in the liver under the influence of growth hormone (GH) and also locally in the skeleton. The IGFs are nonglycosylated single chain polypeptides that exist in serum primarily associated with IGF-binding proteins (IGFBPs), and with an acid-labile protein, as complexes. Since the original discovery of IGFBPs [2], a total of six 99

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100 binding proteins have been identified and named IGFBP-1 to -6 [3]. The most abundant form is IGFBP-3, which is the main binding protein in serum. Although the precise function of each of the IGFBPs has not been fully established, they are themselves under endocrine regulation and play important roles in regulating the distribution of IGFs in tissues and can have both stimulatory and inhibitory effects on IGF function [3]. Since IGFs will bind with greater affinity to the IGFBPs than to the IGF receptor, the binding proteins serve as a potential reservoir for IGF. The affinity of the binding proteins for IGF can be modulated by proteolytic activity, thereby releasing IGF for interaction with its receptor. The IGF-I receptor is structurally related to the insulin receptor and comprises two extracellular α subunits and two transmembrane β subunits which have tyrosine kinase activity. The IGF-II receptor is identical to the mannose-6-phosphate receptor. IGF-I binds with highest affinity to the IGF-I receptor, although it can also bind to the insulin and IGF-II receptors. IGF-II can associate with IGF-I, IGF-II and insulin receptors, although it has a higher affinity for the IGF-II and insulin receptors. A combination of experiments of nature in humans and some very elegant knockout mouse studies have greatly clarified the functions of these growth factors [4–6]. IGF-I and -II regulate embryonic growth, without much influence of GH. GH deficiency only mildly influences birth size whereas congenital IGF deficiency in mice and man [7–9] produces severe stunting which GH administration was unable to correct. After birth, IGF-II function seems to fall away though more human studies are needed to clarify this, and long bone growth comes strongly under GH control of IGF-I release. GH does not only act via the liver since it also has direct effects on the growth plate [10, 11]. Both IGF-I and insulin signal via the insulin receptor substrate proteins. The insulin receptor substrate-1 (IRS-1) -/- mouse is not diabetic but develops severe lowturnover osteopenia as a result of reduced osteoblast proliferation and differentiation, and impaired osteoclastogenesis [12]. IRS-2 deficient mice are also osteopenic as a result of decreased osteoblast differentiation and mineralizing activity, but this is coupled with increased osteoclast activity, secondary to up-regulation of receptor activator of nuclear factor kappa B ligand (RANK-L) expression by the IRS-2 -/- osteoblasts [13]. IGFs have been shown to promote bone resorption in neonatal murine calvariae [14] and fetal long bones [15]. IGF-I and IGF-II can also promote the formation of osteoclasts in bone marrow cultures [14, 16] but have no effect on the activity of isolated osteoclasts [14]. These observations have led to the suggestion that the IGFs can promote bone resorption, albeit indirectly, via an effect on osteoblasts. These effects are likely to be mediated via IGF-I and -II

PHILIPPA HULLEY ET AL.

receptors which have been shown to be expressed by osteoblast-like cells [17–20]. IGFs have been shown to promote the proliferation of isolated osteoblast-like cells and osteoblasts in organ culture [21–23]. Furthermore, both IGF-I and -II promote the synthesis of bone matrix in organ culture and up-regulate type I collagen expression in isolated osteoblast-like cells [23–26]. The effect of IGF has been examined in vivo in a number of rodent models. Tobias et al. [27] demonstrated that daily subcutaneous injections of IGF-I promoted longitudinal bone growth and an increase in periosteal bone formation; however, this was associated with a decrease in trabecular bone formation. In contrast, Bagi et al. [28, 29] reported that IGF-I, when administered as a complex with IGFBP-3, could promote both trabecular and periosteal bone formation. Interestingly, Machwate et al. [30] were able to show that IGF-I could partially prevent the trabecular bone loss associated with disuse; however, IGF-I had no effect on control animals. IGF-I therapy has been tested in humans but has proven relatively ineffective and the high doses needed cause hypoglycemia. However, the need for a precise understanding of the mechanisms of action of IGF-I in adult bone is highlighted by the fact that PTH requires intact IGF-I for its anabolic effects [31], as does thyroid hormone [32]. Estrogen and mechanical strain also exert their effects on osteoblasts in part via IGF-1R [33, 34]. It is interesting to note that targeted osteoblast IGF-I over-expression [35] and osteoblast-specific IGF-1R knockout [36] affect osteoblast function but not osteoblast proliferation.

III. THE TRANSFORMING GROWTH FACTOR-BETA/BONE MORPHOGENETIC PROTEIN SUPERFAMILY Transforming growth factor-beta (TGF-β) was originally identified as a factor that transformed the phenotype of fibroblasts in vitro [37]. It has become apparent that members of the TGF-β superfamily have a number of diverse and complex activities in a wide range of tissues. These include effects on cell growth and proliferation, on apoptosis, and on differentiation and the induction of new gene expression. The TGF-β superfamily comprises a large number of structurally related gene products divided into a number of closely related groups. These include the TGF-β family, activins and inhibins, müllerian inhibiting substance, the bone morphogenetic proteins (BMPs), the cartilage-derived morphogenetic proteins, the Drosophila decapentaplegic gene product (dpp), the Vgr gene products, and glial cell line-derived neurotrophic factor. Given the

Chapter 6 Growth Factors

size of this gene family, our comments will be confined to those members of the family that have been shown to be important in regulating bone or cartilage biology.

A. Transforming Growth Factor-Beta The TGF-β family consists of three mammalian gene products (TGF-β1–3), a gene identified in chicken as TGF-β4, and an additional member of the family identified in Xenopus laevis (TGF-β5). Although mammalian cells can express each of the three isoforms, the majority of information has been generated with β1. Each of the isoforms exist as 25 kDa dimers which share more than 70% amino acid identity. TGF-β is produced as a latent complex of approximately 100 kDa which comprises two molecules, each containing mature TGF-β associated with a precursor, which is also known as latency-associated peptide (LAP). In addition, this latent TGF-β complex has also been shown to be associated with a binding protein with a molecular mass of 190 kDa in fibroblasts and 130 kDa in platelets. The mechanism of activation of latent TGF-β to active TGF-β has not been clearly established, although it involves cleavage by proprotein convertases such as the serine endoprotease, furin, or plasmin [38–40]. Parathyroid hormone, glucocorticoids, and local growth factors can promote cells to activate latent TGF-β. Members of the TGF-β family elicit their biological effect via interactions with specific TGF-β receptors. Two TGF-β receptors (TβRI and TβRII) with masses of 53 and 70 kDa, respectively, have been described [41]. Further analysis has demonstrated both receptors to be functional serine/threonine kinases that are thought to operate as a heterodimer [42, 43], with the TβRII required for ligand binding and both TβRI and TβRII needed for signaling. Studies have suggested that the functional signaling complex is likely to be a tetramer consisting of two TβRI and two TβRII molecules [44]. Interestingly, the type II receptor can bind both TGF-β and TGF-β3 but it has less affinity for TGF-β2. An additional receptor, the type III receptor (betaglycan), appears to be required to present TGF-β2 to the type I/type II signaling complex [45]. The TGF-β receptor complex is known to interact with specific signaling mediators. Studies in Drosophila of the decapentaplegic gene identified a gene named Mothers against dpp (Mad) which is thought to mediate this signaling [46]. Mad has been shown to be a homolog of the small body size (sma) genes of C. elegans [47]. At least nine members of this gene family have also been identified [48] and these are now known as the Smad family of mediators of TGF-β signaling [49]. These molecules translocate to the nucleus where they regulate gene transcription, either by

101 binding directly to DNA, forming co-operative complexes with other transcription factors, or by displacing nuclear factors. There are three sub-classes, R-Smad (Smads-1, -2, -3, -5, -8), Co-smad (Smad-4) and I-Smad (Smad-6 and -7). TGF-β receptors activate Smads-2 and -3, which dimerize with Smad-4 and move to the nucleus, unless inhibited by first binding to an I-Smad. BMPs signal in the same way using R-Smads -1, -5, and -8 [38, 50]. Both TGF-β and BMPs can activate the mitogen-activated protein kinases ERK and p38 and signal independently of the Smads. The role of TGF-β in the skeleton has been the subject of considerable research interest. Early studies demonstrated the presence of two cartilage-inducing factors (CIP-A and -B) in demineralized bone matrix [51], which were subsequently shown to be identical to TGF-β1 and TGF-β2, respectively [52, 53]. The cartilage-inducing properties of TGF-β are illustrated by its ability to co-operate with Wnt signaling to drive differentiation of chondrocytes and suppress adipogenesis in human marrow stromal cultures [54]. Further, in the adult mouse TGF-β plays an essential role in maintaining articular cartilage thickness [55]. Bone has been shown to be a major source of TGF-β1, with reports estimating there to be as much as 200 µg of TGF-β1 per kg of tissue. Osteoblasts produce both the 100 kDa TGF-β precursor [56] and a complex containing the 190 kDa TGF-β binding protein [57]. The latter complex has been suggested to be deposited in bone during synthesis [58]. Factors that promote bone resorption, including 1,25-dihydroxyvitamin D3, parathyroid hormone, and interleukin-1, have been shown to promote the release of TGF-β from fetal rat or neonatal calvarie [59]. In addition, inhibition of bone resorption also prevents release of TGF-β [59]. These data suggest that TGF-β is liberated as bone is resorbed; however, osteoclasts themselves have been shown to produce TGF-β. Northern blot analysis demonstrated the presence of the mRNA for each of the TGF-β2, TGF-β3, and TGF-β4 isoforms in avian osteoclasts, although TGF-β2 was reported to be the principal secreted form [60]. Zheng et al. [61] have also shown that osteoclast-like cells derived from giant cell tumors of bone express the mRNA for TGF-β1. Osteoclasts can activate latent TGF-β in vitro [60, 62] and MMP-9 has been implicated [63]. Although active TGF-β can be released during bone resorption, the effect of this factor has not been fully established because responses appear to be dependent upon the dose of TGF-β studied and the experimental system used. Chenu et al. [64] demonstrated that TGF-β could inhibit the formation of osteoclast-like cells in vitro, a response that was suggested to be mediated by preferentially stimulating the differentiation of cells towards a granulocytic phenotype rather than cells with characteristics of osteoclasts. Shinar and Rodan [65] also reported a decrease

102 in osteoclast-like cell formation in murine bone marrow cultures; however, at low concentrations, TGF-β was reported to increase osteoclast-like cell formation. The effects of TGF-β on bone resorption in organ culture have also been conflicting. TGF-β has been reported to stimulate an increase in bone resorption in a murine neonatal calvaria system [66, 67]. In contrast, high doses of TGF-β cause an inhibition of bone resorption in fetal rat long bones [67, 68]. However, in an ovariectomized rat model, direct injection of TGF-β into the bone marrow space caused a decrease in both the osteoclast surface and numbers of osteoclasts, suggesting that, in vivo, TGF-β may inhibit bone resorption [69]. Although TGF-β appears to have variable effects on bone resorption, a significant body of data has demonstrated that this molecule may be important in promoting bone formation. Genetic evidence comes from Camurati Engelman disease where missense mutations of beta1-LAP (β1-LAP) disrupt the association of β1-LAP and TGF-β1 in the propeptide complex, causing excessive release of mature TGF-β1 [70] and endosteal and perisosteal thickening of the diaphyses. Consistent with this observation, multiple daily injections of TGF-β1 onto the parietal bones of neonatal rats or into the subcutaneous tissue overlying the calvaria of mice have been show to result in an increase in bone formation at the site of injection [71–73]. Beck et al. [74] demonstrated that a single application of TGF-β could induce healing of skull defects in rabbits. Rosen et al. [75] have demonstrated that systemic administration of TGF-β2 to both juvenile and adult rats could also stimulate bone formation. Although the mechanism of increased bone formation is unclear, it has been suggested that TGF-β promotes the proliferation and recruitment of new osteoblasts. In support of this, several studies have shown that TGF-β is chemotactic for bone cells such as rat osteosarcoma and osteoblast-like cells derived from fetal rat calvaria [76, 77]. However, the effect of TGF-β1 on the proliferation of osteoblast-like cells is somewhat unclear, with reports demonstrating either increases [78–80] or decreases in the proliferation of osteoblast-like cells [81, 82], and biphasic effects dependent upon cell density [83]. However, these data are confounded by the fact that many of the cells examined in these studies were osteosarcoma cells or more differentiated osteoblasts rather than osteoblast precursors. Studies have also shown that TGF-β can modulate the differentiated function of these cells. These include studies that demonstrate that TGF-β can both stimulate and inhibit the production or expression of alkaline phosphatase, type I collagen, and osteonectin [82, 84, 85]. However, the effects of TGF-β1 on expression of osteocalcin production and mineralization appear more consistent. A number of reports have demonstrated

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that TGF-β inhibits the expression [85, 86] and synthesis of osteocalcin in osteoblast-like cells [86]. Furthermore, studies have also shown a decrease in the formation of bone nodules in vitro [85, 87, 88]. This ability of a growth factor to build bone in vivo or when administered intermittently, but to repress bone markers in vitro is a confusing but also consistent observation for several growth factors. The explanation could be as simple as internalization and downregulation of receptors following continuous exposure to growth factors, which could be reversed by a washout period. Understanding the underlying signaling will be critical in allowing us to effectively utilize growth factors as bone anabolics. However, TGF-β is not a particularly attractive candidate because it is not clearly anabolic and has a plethora of systemic effects.

B. Bone Morphogenetic Proteins The bone morphogenetic proteins are a large family of proteins that share common structural features with TGF-β1 and other members of the TGF-β superfamily. Members of the BMP family can be further divided into a number of related subgroups based upon sequence identity. BMP family members are synthesized as precursors which are proteolytically processed to yield a mature protein containing seven conserved cysteine residues. Functionally, BMPs exist as homodimers that are linked by disulfide bonds, although a functional heterodimer has been reported [89]. In a similar manner to the TGF-β isoforms, members of the BMP family bind to type I and type II receptors, both of which are required for signal transduction. Two different type I BMP receptors have been identified: BMP receptor-IA (BMPR-IA, also known as ALK-3 (activin receptor-like kinase-3)) and BMP receptor-IB (BMPR-IB or ALK-6) [90, 91]. Although these two receptors are specific for BMP family members, several members can also bind to the activin type I receptor (Act R-I or ALK-2) [90]. A BMP type II receptor (BMPR-II) has also been identified which is specific for BMPs [92, 93]; however, some BMPs can also bind to the activin type II receptors, albeit with reduced affinity [94]. Like TGF-β, signal transduction is mediated by members of the Smad family, with Smad1 and Smad5 being specifically implicated in BMP signaling. Urist [95] made the important discovery that demineralized bone induced new bone formation when implanted into ectopic sites in rodents. Subsequent studies demonstrated that members of the BMP family contributed to this activity and that this family of molecules plays an important role in bone development [96]. BMP-2 and -4 directly induce expression of Sry-type high-mobility group

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(HMG)-box containing transcription factor (SOX9) in all chondroprogenitors, concurrent with expression of cartilage specific type II collagen [97]. During limb bud development BMPs participate not only in chondrogenesis (via BMP-R1B), but also apoptotic response of mesenchymal cells, digit identity and the generation, maintenance, and regression of the apical ectodermal ridge which controls limb outgrowth. Together with FGF, Indian hedgehog, and parathyroid hormone-related peptide (PTHrP), BMPs also regulate the process of endochondral ossification. BMPs -2, -4, -5, and -7 can all induce new bone formation in vivo [98–100] and heterodimers of the different members, including BMP4/7, increase this activity [101]. BMPs appear to initiate a complex cascade of events leading to new bone formation, whether ectopic or within existing skeletal structures, which includes the migration of mesenchymal cells to these sites, their differentiation to chondroblasts and chondrocytes, mineralization of cartilage, angiogenesis, osteoblast differentiation, bone formation, and later on, remodeling of this bone [102]. BMPs may be required to initiate these events, which are subsequently supported by other factors; alternatively, they may modulate the behavior of cells at more than one stage. Several members of the BMP family have been shown to exhibit a chemotactic activity. For example, Cunningham et al. [103] demonstrated that BMP-3 and BMP-4 can promote migration of human monocytes. Furthermore, BMP-2, but neither BMP-4 or BMP-6, are chemotactic for human osteoblasts [104]. BMPs have been shown to determine the differentiation pathways of uncommitted mesenchymal cells. Wang et al. [105] demonstrated that BMP-2 could promote the differentiation of C3H10T1/2 cells, which is a murine mesenchymal cell line, into cells with characteristics of osteoblasts or adipocytes. High concentrations of BMP-2 promoted differentiation into osteoblasts whereas low concentrations favored differentiation into adipocytic cells [105]. Furthermore, Katagiri et al. [106] reported that BMP-2 treatment of C2C12 myoblastic cells resulted in an increase in the expression of alkaline phosphatase, osteocalcin, and PTH-dependent cAMP production and a concomitant reduction in myotube formation. In addition, BMP-6 was shown to promote the differentiation of a rat mesenchymal cell line into cells with characteristics of osteoblasts [107]. BMPs also have effects on committed osteoblast precursors with a number of reports demonstrating that BMPs -2, -3, -4, -6, and -7 can promote differentiation to an osteoblast phenotype, including the up-regulation of alkaline phosphatase activity, type I collagen synthesis, and the expression of osteocalcin [100– 112]. Studies have shown that BMPs can regulate the production of IGFs by bone cells. BMP-2 up-regulates the expression of IGF-I and -II and down-regulates the

103 expression of IGFBP-5 in rat calvarial osteoblasts [113, 114]. BMP-7 promotes expression of IGF-II but not IGF-I in human osteosarcoma cells [115]. Furthermore, BMP-7 stimulates production of IGFBP-3 and -5 but inhibits IGFGP-4 production in these cells, suggesting that BMP-7 can regulate the IGF system in a number of different ways [115]. The importance of BMPs in skeletal development has been exemplified by the demonstration that mutations in the BMP-5 gene in mice lead to the short ear phenotype [116], whereas mutations in growth differentiation factor (GDF-5 or BMP-14) lead to the development of brachypodism [117]. Thomas et al. [118] have reported that a mutation in CDMP-1, the human homolog of GDF-5, is associated with a chondrodysplasia characterized by shortening of the bones of the appendicular skeleton and abnormalities of the bones in the hands and feet (Hunter–Thomson type). Furthermore, it has been suggested that abnormalities in the BMP-2 and -4 genes may be associated with fibrodysplasia ossificans progressiva because mutations in the Drosophila homolog, dpp, result in similar abnormalities [119]. BMP-3 is a negative BMP, opposing the effects of BMP-2 in vitro, and BMP-3null mice have increased bone mineral density and trabecular volume [120]. Recently, the discovery of sclerostin (SOST) and its association with the control of adult bone mass has brought the BMP pathway into focus as a major therapeutic target. SOST is a BMP antagonist, similar to noggin, chordin, and gremlin [121], found primarily in osteocytes, osteoblasts, and hypertrophic chondrocytes [122, 123]. It competes with BMP for access to BMP receptors and blocks BMP signaling via the SMADs, but may also antagonize Wnt signalling. SOST inhibits proliferation of osteoblastic precursors and inhibits alkaline phosphatase activity, in addition to causing apoptosis of human osteoblasts [122]. A null mutation in the SOST gene causes high bone mass in patients with sclerosteosis due to the unopposed action of BMPs [121, 124]. Since SOST is likely to exert its effect on BMPs extracellularly, it should be possible to target this molecule pharmaceutically in the patient population, thus enhancing BMP action and generating bone.

IV. FIBROBLAST GROWTH FACTORS The fibroblast growth factors are a family of heparin binding growth factors that are potent regulators of proliferation and differentiation. The family is thought to consist of at least nine members which share 30–50% amino acid sequence identity and contain two conserved cysteine residues. The prototypes of this family are acidic and basic FGF (FGF-1 and FGF-2, respectively), which were

104 originally identified as factors in extracts of pituitary that stimulated the growth of 3T3 cells [125, 126]. The two proteins that comprised this activity (FGF-1 and FGF-2) share approximately 55% amino acid sequence identity. FGF-1 is approximately 16–18 kDa in size, whereas FGF-2 is slightly larger, approximately 18 kDa, although larger forms have been identified. Neither FGF-1 or -2 possesses a signal sequence, suggesting that these two members of the family are not released via the classical pathway. However, studies have confirmed that they are released from cells by mechanisms that do not involve cell death and that inhibitors of exocytosis can prevent this release [127]. Extracellular FGF can bind to heparin or to heparan sulfate and can therefore be sequestered into the extracellular matrix which acts as a reservoir of this growth factor; alternatively, it can be found bound to cell surface heparin sulfate proteoglycans. FGFs elicit a biological response by interacting with members of the FGF receptor (FGFR) family of transmembrane tyrosine kinases. There are four members of the FGFR family, all of which have a similar general structure; however, alternate mRNA splicing can lead to a large number of isoforms of these receptors [128]. The mechanism by which FGF binds to its receptor and signals to the cell has not been fully determined. Studies have shown that FGF can bind to FGFRs in the absence of heparan sulfate [129]. However, binding of FGF to heparan sulfate appears to increase the affinity of FGF for its receptor [130] and is important for subsequent receptor dimerization and activation [128]. This has led to the suggestion that heparan sulfate acts as a low-affinity receptor for FGF by binding FGF and allowing its presentation to the highaffinity FGFR for signaling. Thus, the receptor complex consists of two FGFR molecules, one heparan sulfate molecule and either one or two FGF molecules [128]. FGF-2 is synthesized by osteoblasts [131, 132], is stored in the bone matrix, and can be released during bone resorption [131, 133]. FGFs have distinct effects on precursor and differentiating osteoblasts – they are potently mitogenic for immature osteoblasts, but repress differentiation by impairing alkaline phosphatase expression and mineralization. Furthermore, apoptosis of differentiated osteoblasts is strongly increased, and this also happens in osteoblasts transfected with the FGF-R activating mutations which cause Apert (S252W) and Crouzon (C342Y) syndromes [134]. Despite these stage-specific effects, recent studies in vivo suggest that systemic administration of FGF-2 results in an increase in the number of osteoblasts and new bone formation [135–137]. Further in vivo evidence for a positive role of FGFs comes from the disruption of the FGF2 gene in mice which causes inhibition of long bone formation [138] and the FGF18 knockout which has

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delayed ossification and cranial suture closure [139, 140]. Conditional knockout of FGFR2 also interferes with osteoblast proliferation and osteogenesis [141]. In addition to their osteogenic effects, FGFs affect osteoclasts. FGF-1 and -2 have been reported to stimulate bone resorption in organ culture [142, 143] and FGF-2 has been shown to stimulate the formation of osteoclasts in murine bone marrow cultures [144]. FGFs work in concert with other growth factors, inducing synthesis of TGF-β, IGF-I, HGF, and VEGF, which are all mitogenic for osteoprogenitor cells, and also inhibit synthesis of the IGF inhibitory IGF-BP5 [145]. Signaling events leading to proliferation and osteoblast differentiation include activation of the mitogen-activated protein kinase, ERK, which stimulates AP1 and drives expression of procollagen, osteopontin, osteocalcin, and VEGF [145]. FGF2 up-regulates cbfa1/ Runx2 expression via activation of PKC [146], as well as inducing ERK-mediated phosphorylation and activation of cbfa1/Runx2 [147]. In addition, FGF signaling co-operatively interacts with that of BMP4 [148, 149]. In humans, the importance of the FGF/FGFR system in bone development has been demonstrated by the identification of genetic mutations that lead to skeletal abnormalities. These mutations are often found in the receptors for FGF and are typified by the creation of an unpaired cysteine residue which can lead to ligand-independent dimerization and constitutive activation [150]. Mutations in FGFR-1 and -2 are found in the craniosynostosis syndromes [150, 151], including Apert [152], Pfeiffer [153, 154], Jackson–Weiss [155], and Crouzon syndromes [156]. Mutations in FGFR-3 have also been shown to lead to skeletal abnormalities. These include a point mutation (Gly380Arg) in the transmembrane region of FGFR-3 that leads to the development of achondroplasia [157] and mutations in the tyrosine kinase domain, Asn540Lys and Lys650Glu, that lead to hypochondroplasia and thanatophoric dysplasia, respectively [158, 159]. In addition, FGF-23 mutations cause hypophosphatemic rickets and high circulating levels of FGF-23 are found in patients with osteomalacia associated with malignancy [160, 161].

V. Wnts The Wnts are perhaps not classical growth factors but are secreted, lipid-modified glycoproteins that actively stimulate bone and cartilage growth during development and in later life (for excellent reviews see [162–164]). There are 19 human Wnts, and at least six of these (Wnt 1, 4, 5a, 7a, 7b, and 14) are made by osteoblast lineage cells. The canonical Wnt signaling pathway (also the most active in osteoblasts) involves a sub-class of Wnts (including 1,

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2, 3a, 8) binding to a receptor complex composed of low-density lipoprotein receptor-related proteins -5 or -6 (Lrp5/6), and the 7-spanning transmembrane Frizzled (Fzd) receptors. This ligand–receptor complex, via activation of the phosphoprotein dishevelled (Dsh), disrupts the multiprotein complex of casein kinase-1 (CK-1), adenomatous polyposis coli (APC), axin, and glycogen synthase kinase-3β (GSK-3β) which normally serves to target β-catenin for degradation. In the absence of this default repression, cellular levels of β-catenin rise, β-catenin translocates to the nucleus and acts with T-cell factor and lymphoid enhancer-binding factor (TCF/LEF) transcription factors to modulate expression of a wide variety of genes, including up-regulation of proliferative cyclin D1 and c-myc. Wnts seem to act on osteoblasts by repressing adipocyte differentiation (Wnt 1 and 10b), enhancing proliferation (Wnt 1, 3a, 6, 7b) and promoting expression of early differentiation markers such as alkaline phosphatase (Wnt 1, 2, and 3a). Later differentiation markers such as osteocalcin are in fact repressed by β-catenin in the mouse [165], suggesting a useful bone anabolic effect via the precursor and young osteoblast compartments only. Human and mouse mutations affecting bone development and the regulation of bone mass have been found in multiple players in the Wnt signaling pathway. Activating mutations are associated with weak activation of β-catenin and high bone mass (HBM) syndrome [166–169]. The classical mutation in Lrp5 prevents binding of the secreted antagonist, dickkopf (DKK) to Lrp5, thus lifting a tonic restraint on Wnt signaling. Increased bone mass is also seen in LRP5 mutants with conditions such as osteopetrosis type I, endosteal hyperostosis, autosomal dominant osteosclerosis and van Buchem disease [170]. Loss of function mutations in LRP5 causes low bone mass in mice and the osteoporosis pseudoglioma (OPPG) syndrome in man [171–173]. Lrp5 mutations seem to affect only osteoblastic bone formation and not osteoclasts – markers of bone turnover are normal in OPPG and high bone mass patients [163]. Typical Wnt-deficient phenotypes are also seen in mice with a point mutation in Lrp6, which reduces the efficiency of the pathway, causing patterning defects and delayed ossification during development and a low bone mass phenotype in the adult [174]. A loss of function nonsense mutation in Wnt3 itself causes the severe patterning defect, tetra-amelia, a complete failure of arm and leg outgrowth in humans [175]. In the normal population, a variety of different LRP5 polymorphisms are associated with either osteoporosis or elevated BMD, indicating a clear contribution of this pathway to the determination of BMD [176]. The full spectrum of Wnts, their antagonists and receptors, come into play during patterning of the embryonic

skeleton and we do not yet have the full picture. It is evident that Wnt7a expressed in the limb ectoderm serves primarily to repress formation of cartilage, thus confining developing bones to the center of limbs [177]. In contrast Wnt 3a stimulates chondrogenesis, and seems to do this in concert with BMP2 [177]. In the adult, cartilage metabolism is also affected by the Wnt pathway, as evidenced by the association of a SNP in secreted frizzled related protein 3 (sFRP3) with osteoarthritis in women [178]. The sFRPs are secreted decoy proteins which normally bind to Wnt in direct competition with membrane receptor Fzd, thereby reducing pathway activation. The mutated sFRP3 is unable to effectively block Wnt signaling. Given that the Wnt signaling pathway is active in most tissues of the body, it is not surprising that mutations in the pathway cause diseases ranging through Alzheimer’s, polycystic kidneys, pulmonary fibrosis, familial exudative vitreoretinopathy, and a variety of cancers [164]. The development of therapies based on manipulation of pathway components is likely to be a complicated business, but carriers of the LRP5 gain-of-function mutation are completely normal except for their high bone mass, giving hope that weak activators of LRP5 signaling may be bone anabolic without significant side-effects.

VI. ADDITIONAL GROWTH FACTORS The growth factors described in the previous sections represent those factors that have been most widely studied in bone and cartilage. However, a number of other growth factors have also been shown to have effects on the cells of the skeleton. These include platelet-derived growth factors (PDGF), members of the epidermal growth factor family, such as epidermal growth factor (EGF), amphiregulin, and transforming growth factor-alpha (TGF-α), and vascular endothelial growth factor (VEGF, also known as vascular permeability factor).

A. Platelet-derived Growth Factors The platelet-derived growth factors now include PDGF-A, -B, -C, and -D and activate receptors PDGF-α and -β. PDGF-A and -B are encoded by separate genes and dimerize to form AA, AB, or BB isoforms. Similarly the α and β subunits of the receptor can dimerize to form αα, ββ, and αβ receptors (reviewed in reference [179]). PDGF-AA binds to the αα receptor, PDGF-AB can bind to both αα and αβ receptors, whereas PDGF-BB can bind to each of the three forms of the receptor [180]. PDGF can be deposited in bone matrix [133] and be released during

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bone resorption or it can be released by osteoblasts to act in either a paracrine or autocrine manner. Indeed, both PDGF-AA and -BB can bind to osteoblasts [181–184], and studies have shown that both osteoblasts and bonelining cells express the PDGF-α receptor in situ [185]. Developing chondrocytes also express PDGF and receptors, and require PDGF for normal development. PDGF has been shown to promote collagen synthesis [181], but these effects are likely to be secondary to their strong stimulation of cell proliferation [182]. In long-term culture, continuous exposure to PDGF decreases alkaline phosphatase expression, type I collagen, and the appearance of mineralized nodules, which is consistent with an inhibitory effect on the differentiated function of the osteoblast [186]. However, short-term exposure to PDGF has the opposite effect and simulates mineralized nodule formation [187], which is consistent with the effects observed in vivo. The local administration of PDGF following osteotomy has been shown to accelerate healing and promote bone formation [188]. Furthermore, systemic administration of PDGF has been shown to prevent bone loss in ovariectomized rats, an effect that is accompanied by an increase in osteoblast number [189]. PDGF has also been shown to have effects on bone resorption. Tashjian et al. [190] demonstrated that partially purified PDGF promoted resorption of neonatal mouse calvariae, an effect that was thought to be mediated by an increase in the local production of PGE2. Subsequent studies demonstrated that PDGF-AB and PDGF-BB, but not PDGF-AA, have the ability to promote bone resorption [191]. PDGF can also promote the production of interleukin-6, a factor that has been implicated in promoting bone resorption by rodent osteoblasts [192]. Furthermore, studies have shown that PDGF-BB, but not PDGF-AA, can up-regulate osteoblast expression of collagenase-1 which may contribute to the degradation of type I collagen and the activation of bone resorption [193]. However, despite its strong mitogenic and bone-building effect in vivo, there are at present no reported human genetic defects in PDGF ligands or receptors with a strong skeletal phenotype.

mitogenic effect via the EGFR and activation of ERK, Akt, c-fos, and c-jun. It inhibits osteoblastic differentiation in vitro but the amphiregulin knockout mouse has reduced tibial trabecular bone, indicating that it normally functions as a bone builder [197]. Similarly, TGF-α can promote PGE2 production in osteoblastic cells [198]. EGF and TGF-α have been shown to promote bone resorption in vitro [199–201]. Studies have also demonstrated that TGF-α can stimulate bone resorption in vivo and that this may contribute to the development of hypercalcemia of malignancy [202].

C. Vascular Endothelial Growth Factor VEGF is a polypeptide growth factor that specifically promotes the proliferation of vascular endothelial cells, an effect that is mediated via VEGF receptors known as flt-1 and KDR. While VEGF essentially works via alterations in the vascularization of bone and cartilage, it is a critically required growth factor for both bone development and fracture repair. Studies have shown that rodent and human osteoblast-like cells express the mRNA for VEGF and also produce the VEGF protein [203–205]. VEGF induces osteoblast differentiation, migration, and is essential for bone healing [206]. Futhermore, the expression of VEGF is promoted by prostaglandins, 1,25dihydroxyvitamin D3, and IGF-I [203–205]. Because blood vessels and endothelial cells are found closely associated with osteoblasts, the VEGF produced by these cells may promote both endothelial cell proliferation and the paracrine production of other growth factors [207]. VEGF is an essential factor for chondrocyte differentiation and is necessary for the survival of hypoxic chondrocytes [208]. Production of VEGF by hypertrophic chondrocytes recruits blood vessels into developing bones, a requirement for subsequent chondroclast invasion and trabecular bone formation [4].

VII. SUMMARY B. Epidermal Growth Factor Family EGF, amphiregulin, and TGF-α are structurally related glycoproteins that bind to the same receptor, EGFR (also called cErb1/HER1), and have been reported to have similar effects on bone. EGF receptors are expressed by osteoblast-like cells [194, 195] and treatment of such cells with EGF results in an increase in cell proliferation [194] and the production of PGE2 [196]. Amphiregulin is strongly induced in osteoblasts by PTH and exerts a potent

The most exciting recent advances in skeletal growth factor biology have emerged from genetic and transgenic approaches and these have greatly enhanced our understanding of the role played by growth factors in the developing and adult skeleton. A more coherent picture of the interplay between different hormones, growth factors and cytokines during development of cartilage and bone is emerging, and with the advent of inducible and tissuespecific knockout and transgenic animal models, we are now beginning to investigate actions of growth factors in the

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adult skeleton separately from their role in development. The development of PTH as a bone anabolic and approved treatment for osteoporosis has triggered a flood of detailed mechanistic studies, which are already highlighting the central involvement of growth factors in orchestrating some of the effects of this therapy. There remain a great many open questions, particularly relating to the actions of growth factors on osteoclasts and on mature articular chondrocytes. Continued research into the skeletal growth factors will open up their use as anabolic and repair agents for bone and cartilage.

References 1. Lane, N. E., and Kelman, A. (2003). A review of anabolic therapies for osteoporosis. Arthritis Res. Ther. 5, 214–222. 2. Hintz, R. L., and Liu, F. (1977). Demonstration of specific plasma protein binding sites for somatomedin. J. dm. Endocrinol. Metab. 45, 988–995. 3. Clemmons, D. R. (1997). Insulin-like growth factor binding proteins and their role in controlling IGF action. Cytokine Growth Factor Rev. 8, 45–62. 4. van der Eerden, B. C., Karperien, M., and Wit, J. M. (2003). Systemic and local regulation of the growth plate. Endocr. Rev. 24, 782–801. 5. Dupont, J., and Holzenberger, M. (2003). Biology of insulin-like growth factors in development. Birth Defects Res. C. Embryo Today 69, 257–271. 6. Sjogren, K., Jansson, J. O., Isaksson, O. G., and Ohlsson, C. (2002). A model for tissue-specific inducible insulin-like growth factor-I (IGF-I) inactivation to determine the physiological role of liver-derived IGF-I. Endocrine 19, 249–256. 7. Powell-Braxton, L., Hollingshead, P., Warburton, C., Dowd, M., PittsMeek, S., Dalton, D., Gillett, N., and Stewart, T. A. (1993). IGF-I is required for normal embryonic growth in mice. Genes Dev. 7, 2609–2617. 8. Liu, J. P., Baker, J., Perkins, A. S., Robertson, E. J., and Efstratiadis, A. (1993). Mice carrying null mutations of the genes encoding insulinlike growth factor I (Igf-1) and type 1 IGF receptor (Igf1r). Cell 75, 59–72. 9. Woods, K. A., Camacho-Hubner, C., Savage, M. O., and Clark, A. J. (1996). Intrauterine growth retardation and postnatal growth failure associated with deletion of the insulin-like growth factor I gene. N. Engl. J. Med. 335, 1363–1367. 10. Wang, J., Zhou, J., Powell-Braxton, L., and Bondy, C. (1999). Effects of Igf1 gene deletion on postnatal growth patterns. Endocrinology 140, 3391–3394. 11. Gevers, E. F., van der Eerden, B. C., Karperien, M., Raap, A. K., Robinson, I. C., and Wit, J. M. (2002). Localization and regulation of the growth hormone receptor and growth hormone-binding protein in the rat growth plate. J. Bone Miner. Res. 17, 1408–1419. 12. Ogata, N., Chikazu, D., Kubota, N., Terauchi, Y., Tobe, K., Azuma, Y., Ohta, T., Kadowaki, T., Nakamura, K., and Kawaguchi, H. (2000). Insulin receptor substrate-1 in osteoblasts is indispensable for maintaining bone turnover. J. Clin. Invest. 105, 935–943. 13. Akune, T., Ogata, N., Hoshi, K., Kubota, N., Terauchi, Y., Tobe, K., Takagi, H., Azuma, Y., Kadowaki, T., Nakamura, K., and Kawaguchi, H. (2002). Insulin receptor substrate-2 maintains predominance of anabolic function over catabolic function of osteoblasts. J. Cell. Biol. 159, 147–156.

107 14. Hill, P. A., Reynolds, J. J., and Meikle, M. C. (1995). Osteoblasts mediate insulin-like growth factor-I and -II stimulation of osteoclast formation and function. Endocrinology 136, 124–131. 15. Slootweg, M. C., Most, W. M., van Beek, E., Schot, L. P. C., Papapoulous, S. E., and Lowik, C. W. G. M. (1992). Osteoclast formation together with interleukin-6 production in mouse long bones is increased by insulin-like growth factor-I. J. Endocrinol. 132, 433–438. 16. Mochizuki, H., Hakeda, Y., Wahatsuki, R., Usui, N., Akashi, S., Sato, T., Tanaka, K., and Kumegawa, M. (1992). Insulin-like growth factor-I supports formation and activation of osteoclasts. Endocrinology 131, 1075–1080. 17. Mohan, S., Linkhart, T., Rosenfield, R., and Baylink, D. (1989). Characterisation of the receptor for insulin-like growth factor II in bone cells. J. Cell. Physiol. 140, 169–176. 18. Centrella, M., McCarthy, T. L., and Canalis, E. (1990). Receptors for insulin-like growth factors-I and -II in osteoblast-enriched cultures from fetal rat bone Endocrinology 126, 39–44. 19. Slootweg, M. C., Hoogerbrugge, C. M., de Poorter, T. L., Duursma, S. A., and van Buul-Offers, S. C. (1990). The presence of classical insulin-like growth factor (IGF) type-I and -II receptors on mouse osteoblasts: Autocrine/paracrine growth effect of IGFs? J. Endocrinol. 125, 271–277. 20. Cantrell, P. R., Frolik, C. A., Ellis, L. F., and Williams, D. C. (1991). Binding of insulin-like growth factors to cultured rat calvarial cells with differing biologic responses. J. Bone Miner. Res. 6, 851–857. 21. Fournier, B., Ferralli, J. M., Price, P. A., and Schlaeppi, J. M. (1993). Comparison of the effects of insulin-like growth factors-I and -II on the human osteosarcoma cell line OHS-4. J. Endocrinol. 136, 173–180. 22. Hock, J. M., Centrella, M., and Canalis, E. (1988). Insulin-like growth factor I has independent effects on bone matrix formation and cell replication. Endocrinology 122, 254–260. 23. Pfeilschifter, J., Oechsner, M., Naumann, A., Gronwald, R. G. K., Minne, H. W., and Ziegler, R. (1990). Stimulation of bone matrix apposition in vitro by local growth factors: A comparison between insulin-like growth factor I, platelet-derived growth factor, and transforming growth factor β. Endocrinology 127, 69–75. 24. Canalis, E. (1993). Insulin like growth factors and the local regulation of bone formation. Bone 14, 273–276. 25. Schmid, C., Guler, H.-P., Rowe, D., and Froesch, E. D. (1989). Insulinlike growth factor I regulates type I procollagen messenger ribonucleic acid steady state levels in bone of rats. Endocrinology 125, 1575–1580. 26. Strong, D. D., Beachler, A. L., Wergedal, J. E., and Linkhart, T. A. (1991). Insulin-like growth factor II and transforming growth factor β regulate collagen expression in human osteoblast-like cells in vitro. J. Bone Miner. Res. 6, 15–23. 27. Tobias, J. H., Chow, J. W. M., and Chambers, T. J. (1992). Opposite effects of insulin-like growth factor-I on the formation of trabecular and cortical bone in adult female rats. Endocrinology 131, 2387–2392. 28. Bagi, C. M., Brommage, R., Deleon, L., Adams, S., Rosen, D., and Sommer, A. (1994). Benefit of systemically administered rhIGF-I and rhIGF-I/IGFBP-3 on cancellous bone in ovariectomized rats. J. Bone Miner. Res. 9, 1301–1312. 29. Bagi, C. M., Deleon, E., Brommage, R., Adams, S., Rosen, D., and Sommer, A. (1995). Systemic administration of rhIGF-I or rhIGF-I/IGFBP-3 increases cortical bone and lean body mass in ovariectomized rats. Bone 16, 263S–269S. 30. Machwate, M., Zerath, E., Holy, X., Pastoureau, P., and Marie, P. J. (1994). Insulin-like growth factor-I increases trabecular bone formation and osteoblastic cell proliferation in unloaded rats. Endocrinology 134, 1031–1038. 31. Bikle, D. D., Sakata, T., Leary, C., Elalieh, H., Ginzinger, D., Rosen, C. J., Beamer, W., Majumdar, S., and Halloran, B. P. (2002). Insulin-like growth factor I is required for the anabolic actions of parathyroid hormone on mouse bone. J. Bone Miner. Res. 17, 1570–1578. 32. Huang, B. K., Golden, L. A., Tarjan, G., Madison, L. D., and Stern, P. H. (2000). Insulin-like growth factor I production is essential for anabolic

108

33.

34.

35.

36.

37.

38.

39.

40.

41.

42.

43.

44.

45. 46.

47.

48.

effects of thyroid hormone in osteoblasts. J. Bone Miner. Res. 15, 188–197. Cheng, M. Z., Rawlinson, S. C., Pitsillides, A. A., Zaman, G., Mohan, S., Baylink, D. J., and Lanyon, L. E. (2002). Human osteoblasts’ proliferative responses to strain and 17beta-estradiol are mediated by the estrogen receptor and the receptor for insulin-like growth factor I. J. Bone Miner. Res. 17, 593–602. Kasukawa, Y., Stabnov, L., Miyakoshi, N., Baylink, D. J., and Mohan, S. (2002) Insulin-like growth factor I effect on the number of osteoblast progenitors is impaired in ovariectomized mice. J. Bone Miner. Res. 17, 1579–1587. Zhao, G., Monier-Faugere, M. C., Langub, M. C., Geng, Z., Nakayama, T., Pike, J. W., Chernausek, S. D., Rosen, C. J., Donahue, L. R., Malluche, H. H., Fagin, J. A., and Clemens, T. L. (2000). Targeted overexpression of insulin-like growth factor I to osteoblasts of transgenic mice: increased trabecular bone volume without increased osteoblast proliferation. Endocrinology 141, 2674–2682. Zhang, M., Xuan, S., Bouxsein, M. L., von Stechow, D., Akeno, N., Faugere, M. C., Malluche, H., Zhao, G., Rosen, C. J., Efstratiadis, A., and Clemens, T. L. (2002). Osteoblast-specific knockout of the insulinlike growth factor (IGF) receptor gene reveals an essential role of IGF signaling in bone matrix mineralization. J. Biol. Chem. 277, 44005–44012. Roberts, A. B., Anzano, M. A., Lamb, L. C., Smith, J. M., and Sporn, M. B. (1981). New class of transforming growth factors potentiated by epidermal growth factor. Proc. Natl. Acad. Sci. USA. 78, 5339–5343. Canalis, E., Economides, A. N., Gazzerro, E (2003). Bone morphogenetic proteins, their antagonists and the skeleton. Endocrine Rev. 24, 218–235. Sato, Y., and Rifkin, D. B. (1989). Inhibition of endothelial cell movement by pericytes and smooth muscle cells: Activation of a latent transforming growth factor- β1 -like molecule by plasmin during co-culture. J. Cell Biol. 109, 309–315. Lyons, R. M., Gentry, L. E., Purchio, A. F., and Moses, H. L. (1990). Mechanism of activation of latent recombinant transforming growth factor-β by plasmin. J. Cell Biol. 110, 1361–1397. Cheifetz, S., Like, B., and Massague, J. (1986). Cellular distribution of type I and type II receptors for transforming growth factor-β. J. Biol. Chem. 261, 9972–9978. Lin, H. Y., Wang, X.-R., Ng-Eaton, E., Weinberg, R. A., and Lodish, H. F. (1992). Expression cloning of the TGF β type II receptor, a functional transmembrane serine/threonine kinase. Cell 68, 775–785. Bassing, C. H., Yingling, J. M., Howe, D. J., Wang, T., He, W. W., Gustafson, M. L., Shah, P., Donahoe, P. K., and Wang, X.-F. (1994). A transforming growth factor β type I receptor that signals to activate gene expression. Science 263, 87–89. Weiss-Garcia, F., and Massague, J. (1996). Complementation between kinase-defective and activation defective TGF-β receptors reveals a novel form of receptor co-operativity essential for signaling. EMBO J. 15, 276–289. Alevizopoulos, A., and Mermod, N. (1997). Transforming growth factor-β: The breaking open of a black box. Bio Essays 19, 581–591. Sekelsky, J. J., Newfeld, S. J., Raftery, L. A., Chartoff, E. H., and Gelbart, W. M. (1995). Genetic characterization and cloning of mothers against dpp, a gene required for decapentaplegic function in Drosophila melanogaster. Genetics 139, 1347–1358. Savage, C., Das, P., Finelli, A. L., Townsend, S. R., Sun, C.-Y., Baird, S. E., and Padgett, R. W. (1996). Caenorhabditis elegans genes sma-2, sma-3 and sma-4 define a conserved family of transforming growth factor β pathway components. Proc. Natl. Acad. Sci. USA 93, 790–794. Kawabata, M., Imamura, T., and Miyazono, K. (1998). Signal transduction by bone morphogenetic proteins. Cytokine Growth Factor Rev. 9, 49–61.

PHILIPPA HULLEY ET AL. 49. Derynck, R., Gelbart, W. M., Harland, R. M., Heldin, C.-H., Kem, S. E., Massague, J., Melton, D. A., Mlodzik, M., Padgett, R. W., Roberts, A. B., Smith, J., Thomsen, G. H., Vogelstein, B., and Wang, X.-F. (1996). Nomenclature: Vertebrate mediators of TGF β family signals. Cell 87, 173. 50. Katagiri, T., and Takahashi, N. (2002). Regulatory mechanisms of osteoblast and osteoclast differentiation. Oral Diseases 8, 147–159. 51. Seyedin, S. M., Thomas, T. C., Thompson, A. Y., Rosen, D. M., and Piez, K. A. (1985). Purification and characterization of two cartilageinducing factors from bovine demineralised bone. Proc. Natl. Acad. Sci. USA 82, 2267–2271. 52. Seyedin, S. M., Thompson, A. Y., Bentz, H., Rosen, D. M., McPherson, J. M., Conti, A., Siegel, N. R., Galluppi, G. R., and Piez, K. A. (1986). Cartilage-inducing factor-A. Apparent identity to transforming growth factor-β. J. Biol. Chem. 261, 5693–5695. 53. Centrella, M., McCarthy, T. L., and Canalis, E. (1991). Transforming growth factor-beta and remodeling of bone. J. Bone J. Surg. 73, 1418–1428. 54. Zhou, S., Eid, K., Glowacki, J. (2004). Cooperation between TGF-β and Wnt pathways during chondrocyte and adipocyte differentiation of human marrow stromal cells. J. Bone Miner. Res. 19, 463–470. 55. Scharstuhl, A., Vitters, E. L., van der Kraan, P. M., van den Berg, W. B. (2003). Reduction of osteophyte formation and synovial thickening by adenoviral overexprssion of TGF β /BMP inhibitors during experimental osteoarthritis. Arthritis and Rheumatism 48, 3442–3451. 56. Bonewald, L. F., Wakefield, L., Oreffo, R. O. C., Escobedo, A., Twardzik, D. R., and Mundy, G. R. (1991). Latent forms of transforming growth factor-beta (TGF-β) derived from bone cultures. Identification of a naturally occuring 100-kDa complex with similarity to recombinant latent TGF-β. Mol. Endocrinol. 5, 741–751. 57. Dallas, S. L., Park-Snyder, S., Miyazono, K., Twardzik, D., Mundy, G. R., and Bonewald, B. L. (1994). Characterization and autoregulation of latent transforming growth factor β (TGF-β) complexes in osteoblast-like cell lines: Production of a latent complex lacking the latent TGF β-binding protein (LTBP). J. Biol. Chem. 269, 6815–6822. 58. Bonewald, L. F., and Dallas, S. L. (1994). Role of active and latent transforming growth factor β in bone formation. J. Cell. Biochem. 55, 350–357. 59. Pfeilschifter, J., and Mundy, G. R. (1987). Modulation of type β transforming growth factor activity in bone cultures by osteotropic hormones. Proc. Natl. Acad. Sci. USA 84, 2024–2028. 60. Oursler, M. J. (1994). Osteoclast synthesis and secretion and activation of latent transforming growth factor β. J. Bone Miner. Res. 9, 443–451. 61. Zheng, M. H., Fan, Y., Wysocki, S. J., Lau, A. T.-T., Robertson, T., Beilharz, M., Wood, D. J., and Papadimitriou, J. M. (1994). Gene expression of transforming growth factor β 1 and its type II receptor in giant cell tumors of bone. Am. J. Pathol. 145, 1095–1104. 62. Oreffo, R. O. C., Mundy, G. R., Seyedin, S. M., and Bonewald, L. F. (1989). Activation of the bone derived latent TGF beta complex by isolated osteclasts. Biochem. Biophys. Res. Commun. 158, 817–823. 63. Karsdal, M. A., Hjorth, P., Henriksen, K., Kirkegaard, T., Nielsen, K. L., Lou, H., Delaisse, J. M., Foged, N. T. (2003). Transforming growth factor-beta controls human osteoclastogenesis through the p38 MAPK and regulation of RANK expression. J. Biol. Chem. 278, 44975–44987. 64. Chenu, C., Pfeilschifter, J., Mundy, G. R., and Roodman, G. D. (1988). Transforming growth factor β inhibits formation of osteoclast-like cells in long-term human marrow cultures. Proc. Natl. Acad. Sci. USA 85, 5683–5687. 65. Shinar, D. M., and Rodan, G. A. (1990). Biphasic effects of transforming growth factor β on the production of osteoclast-like cells in mouse bone marrow cultures: The role of prostaglandins in the generation of these cells. Endocrinology 126, 3153–3158. 66. Tashjian, A. H., Voelkel, E. E., Lazzaro, M., Singer, F. R., Roberts, A. B., Derynck, R., Winkler, M. E., and Levine, L. (1985). Alpha and beta human transforming growth factors stimulate prostaglandin production

109

Chapter 6 Growth Factors

67.

68.

69.

70.

71.

72.

73.

74.

75.

76. 77.

78.

79.

80.

81.

82.

83.

84.

and bone resorption in cultured mouse calvaria. Proc. Natl. Acad. Sci. USA 82, 4535–4538. Dieudonne, S. C., Foo, P., Van Zoelen, E. J. J., and Burger, E. H. (1991). Inhibiting and stimulating effects of TGF-β1 on osteoclastic bone resorption in fetal mouse bone organ cultures. J. Bone Miner. Res. 6, 479–487. Pfeilschifter, J. P., Seyedin, S., and Mundy, G. R. (1988). Transforming growth factor beta inhibits bone resorption in fetal rat long bone cultures. J. Clin. Invest. 82, 680–690. Beaudreuil, J., Mbalaviele, G., Cohen-Solal, M., Morieux, C., De Vernejoul, M. C., and Orcel, P. (1995). Short-term local injections of transforming growth factor β1 decrease ovariectomy-stimulated osteoclastic resorption in vivo in rats. J. Bone Miner. Res. 10, 971–977. Saito, T., Kinoshita, A., Yoshiura, K., Makita, Y., Wakui, K., Honke, K., Niikawa, N., and Taniguchi, N. (2001). Domain-specific mutations of a transforming growth factor (TGF)-beta 1 latency-associated peptide cause Camurati-Engelmann disease because of the formation of a constitutively active form of TGF-beta 1. J. Biol. Chem. 276, 11469–11472. Noda, M., and Camilliere, J. J. (1989). In vivo stimulation of bone formation by transforming growth factor β. Endocrinology 124, 2991–2994. Mackie, E. J., and Trechsel, U. (1990). Stimulation of bone formation in vivo by transforming growth factor β remodeling of woven bone and lack of inhibition by indomethacin. Bone 11, 295–300. Marcelli, C., Yates, A. J., and Mundy, G. R. (1990). In vivo effects of human recombinant transforming growth factor β on bone turnover in normal mice. J. Bone Miner. Res. 10, 1087–1096. Beck, L. S., DeGuzman, L., Lee, W. P., Xu, Y., McFatridge, L. A., Gillett, N. A., and Amento, E. P. (1991). TGF-β1 induces bone closure of skull defects. J. Bone Miner. Res. 6, 1257–1265. Rosen, D., Miller, S. C., Deleon, E., Thompson, A. Y., Bentz, H., Mathews, M., and Adams, S. (1994). Systemic administration of recombinant transforming growth factor beta 2 (rTGF-β2) stimulates parameters of cancellous bone formation in juvenile and adult rats. Bone 15, 355–359. Lucas, P. A. (1989). Chemotactic response of osteoblast-like cells to transforming growth factor beta. Bone 10, 459–463. Pfeilschifter, J., Wolf, O., Naumann, A., Minne, H. W., Mundy, G. R., and Ziegler, R. (1990). Chemotactic response of osteoblast-like cells to transforming growth factor β. J. Bone Miner. Res. 5, 825–830. Robey, P. G., Young, M. F., Flanders, K. C., Roche, N. S., Kondaiah, P., Reddi, A. H., Termine, J. D., Sporn, M. B., and Roberts, A. (1987). Osteoblasts synthesize and respond to transforming growth factor type-β (TGF-β) in vitro. J. Cell Biol. 105, 457–463. Uneno, S., Yamamoto, I., Yamamuro, T., Okumura, H., Ohta, S., Lee, K., Kasai, R., and Konishi, J. (1989). Transforming growth factor β modulates proliferation of osteoblastic cells: Relation to its effect on receptor levels for epidermal growth factor. J. Bone Miner. Res. 4, 165–171. Chen, T. L., Bates, R. L., Dudley, A., Hammonds, R. G., and Amento, E. P. (1991). Bone morphogenetic protein-2b stimulation of growth and osteogenic phenotypes in rat osteoblast-like cells: Comparison with TGF-β1. J. Bone Miner. Res. 6, 1387–1393. Pfeilschifter, J., D’Souza, S. M., and Mundy, G. R. (1987). Effects of transforming growth factor beta on osteoblastic osteosarcoma cells. Endocrinology 121, 212–218. Noda, M., and Rodan, G. (1986). Type β transforming growth factor inhibits proliferation and expression of alkaline phosphatase in murine osteoblast-like cells. Biochem. Biophys. Res. Commun. 140, 56–65. Centrella, M., McCarthy, T. L., and Canalis, E. (1987). Transforming growth factor β is a bidirectional regulator of replication and collagen synthesis in osteoblast-enriched cell cultures from fetal rat bone J. Biol. Chem. 262, 2869–2874. Noda, M., and Rodan, G. A. (1987). Type β transforming growth factor (TGF-β) regulation of alkaline phosphatase expression and other

85.

86.

87.

88.

89.

90.

91.

92.

93.

94.

95. 96. 97. 98.

99.

100.

phenotype-related mRNAs in osteoblastic rat osteosarcoma cells. J. Cell Physiol. 133, 426–437. Harris, S. E., Bonewald, L. E., Harris, M. A., Sabatini, M., Dallas, S., Feng, J. Q., Ghosh-Choudhury, N., Wozney, J., and Mundy, G. R. (1994). Effects of transforming growth factor β on bone nodule formation and expression of bone morphogenetic protein 2, osteocalcin, osteopontin, alkaline phosphatase, and type I collagen mRNA in long-term cultures of fetal calvarial osteoblasts. J. Bone Miner. Res. 9, 855–863. Pirskanen, A., Jaaskelainen, T., and Maenpaa, P. H. (1994). Effects of transforming growth factor β1 on the regulation of osteocalcin synthesis in human MG-63 osteosarcoma cells. J. Bone Miner. Res. 9, 1635–1642. Antosz, M. E., Bellows, C. G., and Aubin, J. E. (1989). Effects of transforming growth factor β and epidermal growth factor on cell proliferation and the formation of bone nodules in isolated fetal rat calvaria cells. J. Cell Physiol. 140, 386–395. Chen, T. L., and Bates, R. L. (1993). Recombinant human transforming growth factor beta 1 modulates bone remodeling in a mineralizing bone organ culture. J. Bone Miner. Res. 8, 423–434. Sampath, T. K., Coughlin, J. E., Whetstone, R. M., Banach, D., Corbett, C., Ridge, R. J., Ozkaynak, E., Oppermann, H., and Rueger, D. C. (1990). Bovine osteogenic protein is composed of dimers of OP-1 and BMP-2 A, two members of the transforming growth factor-β superfamily. J. Biol. Chem. 265, 13198–13250. ten Dijke, P., Yamashita, H., Sampath, T. K., Reddi, A. H., Estevez, M., Riddle, D. L., Ichijo, H., Heldin, C.-H., and Mijazono, K. (1994). Identification of type I receptors for osteogenic protein-1 and bone morphogenetic protein-4. J. Biol. Chem. 269, 16985–16988. Koenig, B. B., Cook, J. S., Wolsing, D. H., Ting, J., Tiesman, J. P., Correa, P. E., Olson, C. A., Pecquet, A. L., Ventura, E., Grant, R. A., Chen, G.-X., Wrana, J. L., Massague, J., and Rosenbaum, J. S. (1994). Characterization and cloning of a receptor for BMP-2 and BMP-4 from NIH 3T3 cells. Mol. Cell. Biol. 14, 5961–5974. Rosenzweig, B. L., Imamura, T., Okadome, T., Cox, G. N., Yamashita, H., ten Dijke, P., Heldin, C.-H., and Miyazono, K. (1995). Cloning and characterization of a human type II receptor for bone morphogenetic proteins. Proc. Natl Acad, Sci. USA 92, 7632–7636. Liu, F., Ventura, F., Doody, J., and Massague, J. (1995). Human type II receptor for bone morphogenetic proteins (BMPs): Extension of the two-kinase receptor model to the BMPs. Mol. Cell. Biol. 15, 3479–3486. Yamashita, H., ten Dijke, P., Huylebroeck, D., Sampath, T. K., Andries, M., Smith, J. C., Heldin, C.-H., and Miyazono, K. (1995). Osteogenic protein-1 binds to activin type II receptors and induces certain activin-like effects. J. Cell Biol. 130, 217–226. Urist, M. R. (1965). Bone: Formation by autoinduction. Science 150, 893–899. Sakou, T. (1998). Bone morphogenetic proteins: From basic studies to clinical approaches. Bone 22, 591–603. Shum, L., and Nuckolls, G. (2002). The life cycle of chondrocytes in the developing skeleton. Arthritis Res. 4, 94–106. Wang, E. A., Rosen, V., D’Alessandro, J. S., Bauduy, M., Cordes, P., Harada, T., Israel, D. I., Hewick, R. M., Kerns, K. M., LaPan, P., Luxenberg, D. P., McQuaid, D., Moutsatsos, I. K., Nove, J., and Wozney, J. M. (1990). Recombinant human bone morphogenetic protein induces bone formation. Proc. Natl Acad. Sci. USA 87, 2220–2224. D’Alessandro, J. S., Cox, K. A., Israil, D. L., LaPan, P., Moutsatsos, I. K., Nove, J., Rosen, V., Ryan, M. C., Wozney, J. M., and Wang, E. A. (1991). Purification, characterization and activities of recombinant bone morphogenetic protein-5. J. Bone Miner. Res. 6, S153. Sampath, T. K., Maniacal, J. C., Hauschka, P. V., Jones, W. K., Sasak, H., Tucker, R. F., White, K. H., Coughlin, J. E., Tucker, M. M., Pang, R. H. L., Corbett, C., Ozkaynak, E., Oppermann, H., and Rueger, D. C. (1992). Recombinant human osteogenic protein-1 (hOP-1) induces new bone formation in vivo with a specific activity comparable with natural bovine osteogenic protein and stimulates

110

101.

102.

103.

104.

105.

106.

107.

108.

109.

110.

111.

112.

113.

114.

115.

116.

117.

PHILIPPA HULLEY ET AL. osteoblast proliferation and differentiation in vitro. J. Biol. Chem. 267, 20352–20362. Aono, A., Hazama, M., Notoya, K., Taketomi, S., Yamasaki, H., Tsukuda, R., Sasaki, S., and Fujisawa, Y. (1995). Potent ectopic boneinducing activity of bone morphogenetic protein -4/7 heterodimer. Biochem. Biophys. Res. Commun. 210, 670–677. Reddi, A. H. (1997). Bone morphogenetic proteins: An unconventional approach to isolation of first mammalian morphogens. Cytokine Growth Factor Rev. 8, 11–20. Cunningham, N. S., Paralkar, V., and Reddi, A. H. (1992). Osteogenin and recombinant bone morphogenetic protein 2B are chemotactic for human monocytes and stimulated transforming growth factor β1 mRNA expression. Proc. Natl Acad. Sci. USA 89, 11740–11744. Lind, M., Eriksen, E. E., and Bunger, C. (1996). Bone morphogenetic protein-2 but not bone morphogenetic protein-4 and -6 stimulates chemotactic migration of human osteoblasts, human marrow osteoblasts, and U2-OS cells. Bone 18, 53–57. Wang, E. A., Israel, D. I., Kelly, S., and Luxemberg, D. P. (1993). Bone morphogenetic protein-2 causes commitment and differentiation in C3H10T1/2 and 3T3 cells. Growth Factors 9, 57–71. Katagiri, T., Yamaguchi, A., Komaki, M., Abe, E., Takahashi, N., Ikeda, T., Rosen, V., Wozney, J. M., Fujisawa-Sehara, A., and Suda, T. (1994). Bone morphogenetic protein-2 converts the differentiation pathway of C2C12 myoblasts into the osteoblast lineage. J. Cell Biol. 127, 1755–1766. Gitelman, S. E., Kirk, M., Ye, J.-Q., Filvaroff, E. H., Kahn, A. J., and Derynck, R. (1995). Vgr-l/BMP-6 induces osteoblastic differentiation of pluripotential mesenchymal cells. Cell Growth Differ. 6, 827–836. Takuwa, Y., Ohse, C., Wang, E. A., Wozney, J. M., and Yamashita, K. (1991). Bone morphogenetic protein-2 stimulates alkaline phosphatase activity and collagen synthesis in cultured osteoblastic cells, MC3T3-El. Biochem. Biophys. Res. Common. 174, 96–101. Maliakal, J. C., Asahina, I., Hauschka, P. V., and Sampath, T. K. (1994). Osteogenic protein-1 (BMP-7) inhibits cell proliferation and stimulates the expression of markers characteristic of osteoblast phenotype in rat osteosarcoma (17/2.8) cells. Growth Factors 11, 227–234. Hughes, F. J., Collyer, J., Stanfield, M., and Goodman, S. (1995). The effect of bone morphogenetic protein-2, -4 and -6 on differentiation of rat osteoblast cells in vitro. Endocrinology 136, 2671–2677. Chen, T. L., Bates, R. L., Dudley, A., Hammonds, R., and Amento, E. P. (1991). Bone morphogenetic protein-2b stimulation of growth and osteogenic phenotypes in rat osteoblast-like cells: Comparison with TGF-β1. J. Bone Miner. Res. 6, 1387–1393. Zhou, H., Hammonds, R., Findlay, D. M., Martin, T. J., and Ng, K. W. (1993). Differential effects of transforming growth factor-beta 1 and bone morphogenetic protein 4 on gene expression and differentiated function of preosteoblasts. J. Cell Physiol. 155, 112–119. Canalis, E., and Gabbitas, B. (1994). Bone morphogenetic protein 2 increases insulin-like growth factor I and II transcripts and polypeptide levels in bone cell cultures. J. Bone Miner. Res. 9, 1999–2005. Gabbitas, B., and Canalis, E. (1995). Bone morphogenetic protein-2 inhibits the synthesis of insulin-like growth factor-binding protein-5 in bone cell cultures. Endocrinology 136, 2397–2403. Knutsen, R., Honda, Y., Strong, D. D., Sampath, T. K., Baylink, D. J., and Mohan, S. (1995). Regulation of insulin-like growth factor system components by osteogenic protein-1 in human bone cells. Endocrinology 136, 857–865. Kingsley, D. M., Bland, A. E., Grubber, J. M., Marker, P. C., Russell, L. B., Copeland, N. G., and Jenkins, N. A. (1992). The mouse short ear skeletal morphogenesis locus is associated with defects in a bone morphogenetic member of the TGF beta superfamily. Cell 71, 339–410. Storm, E. E., Huynh, T. V., Copeland, N. G., Jenkins, N. A., Kingsely, D. M., and Lee, S.-J. (1994). Limb alterations in brachypodism

118.

119. 120.

121.

122.

123.

124.

125. 126.

127.

128. 129.

130.

131.

132.

133.

134.

135.

mice due to mutations in a new member of the TGF β-superfamily. Nature 368, 639–643. Thomas, J. T., Lin, K., Nandedkar, M., Camargo, M., Cervenka, J., and Luyten, F. P. (1996). A human chondrodysplasia due to a mutation in a TGF-β superfamily member. Nat. Genet. 12, 315–317. Kaplan, F. S., Tabas, J. A., and Zasloff, M. A. (1990). Fibrodysplasia ossificans progressiva: A clue from the fly? Calcif. Tissue Int. 46, 117–125. Daluiski, A., Engstrand, T., Bahamonde, M. E., Gamer, L. W., Agius, E., Stevenson, S. L., Cox, K., Rosen, V., and Lyons, K. M. (2001). Bone morphogenetic protein–3 is a negative regulator of bone density. Nat. Genet. 27, 84–88. Balemans, W., Ebeling, M., Patel, M., VanHul, E., Olson, P., Dioszegi, M., et al. (2001). Increased bone density in sclerosteosis is due to the deficiency of a novel secreted protein (SOST). Hum. Mol. Genet. 10, 537–543. Sutherland, M. K., Geoghegan, J. C., Yu, C., Turcott, E., Skonier, J. E., Winkler, D. G., and Latham, J. A. (2004). Sclerostin promotes the apoptosis of human osteoblastic cells: A novel regulation of bone formation. Bone 35, 828–835. van Bezooijen, R. L., Roelen, B. A., Visser, A., van der Wee-Pals, L., de Wilt, E., Karperien, M., Hamersma, H., Papapoulos, S. E., ten Dijke, P., and Lowik, C. W. (2004). Sclerostin is an osteocyteexpressed negative regulator of bone formation, but not a classical BMP antagonist. J. Exp. Med. 199, 805–814. Brunkow, M. E., Gardner, J. C., VanNess, J., Paeper, B. W., Kovacevich, B. R., Proll, S., et al. (2001). Bone dysplasia sclerosteosis results from the loss of the SOST gene product, a novel cysteine knot-containing protein. Am. J. Hum. Genet. 68, 577–589. Armelin, H. A. (1973). Pituitary extracts and steroid hormones in the control of 3T3 growth. Proc. Natl Acad. Sci. USA 70, 2702–2706. Gospodarowicz, D. (1974). Localization of a fibroblast growth factor and its effect alone and with hydrocortisone on 3T3 cell growth. Nature 326, 123–127. Migantti, P., Morimoto, T., and Rifkin, D. G. (1992). Basic fibroblast growth factor, a protein devoid of secretory signal sequence, is released by cells via a pathway independent of the endoplasmic reticulum-golgi complex. J. Cell Physiol. 151, 81–93. Green, P. J., Walsh, F. S., and Doherty, P. (1996). Promiscuity of fibroblast growth factor receptors. BioEssays 18, 639–646. Roghani, M., Mansukhani, A., Dell’Era, P., Bellosta, P., Basilico, C., Rifkin, D. B., and Moscatelli, D. (1994). Heparin increases the affinity of bFGF for its receptor, but is not required for binding. J. Biol. Chem. 269, 3976–3984. Ornitz, D. M., and Leder, P. (1992). Ligand specificity and heparin dependence of fibroblast growth factor receptors 1 and 3. J. Biol. Chem. 267, 16305–16311. Globus, R. K., Plouet, J., and Gospodarowicz, D. (1989). Cultured bovine bone cells synthesise basic fibroblast growth factor and store it in their extracellular matrix. Endocrinology 124, 1539–1547. Hurley, M. J., Abreu, C., Gronowicz, G., Kawaguchi, H., and Lorenzo, J. (1994). Expression and regulation of basic fibroblast growth factor mRNA levels in mouse osteoblastic MC3T3-E1 cells. J. Biol. Chem. 268, 9392–9396. Hauschka, P. V., Mavrakos, A. E., lafrati, M. D., Doleman, S. E., and Klagsbrun, M. (1986). Growth factors in bone matrix. Isolation of multiple types by affinity chromatography on heparin-sepharose. J. Biol. Chem. 261, 12665–12674. Mansukhani, A., Bellosta, P., Sahni, M., and Basilico, C. (2000). Signaling by fibroblast growth factors (FGF) and fibroblast growth factor receptor 2 (FGFR2)-activating mutations blocks mineralization and induces apoptosis in osteoblasts. J. Cell Biol. 149, 1297–1308. Mayahara, H., Ito, T., Nagai, H., Miyajima, H., Tsukuda, R., Taketomi, S., Mizoguchi, J., and Kato, K. (1993). In vivo stimulation of endosteal bone formation by basic fibroblast growth factor in rats. Growth Factors 9, 73–80.

111

Chapter 6 Growth Factors 136. Nakamura, T., Hanada, K., Tamura, M., Shibanushi, T., Nigi, H., Tagawa, M., Fukumoto, S., and Matsumoto, T. (1995). Stimulation of endosteal bone formation by systemic injections of recombinant basic fibroblast growth factor in rats. Endocrinology 136, 1276–1284. 137. Nagai, H., Tsukuda, T., and Mayahara, H. (1995). Effect of basic fibroblast growth factor (bFGF) on bone formation in growing rats. Bone 16, 367–373. 138. Montero, A., Okada, Y., Tomita, M., Ito, M., Tsurukami, H., Nakamura, T., Doetschman, T., Coffin, J. D., and Hurley, M. M. (2000). Disruption of the fibroblast growth factor-2 gene results in decreased bone mass and bone formation. J. Clin. Invest. 105, 1085–1093. 139. Liu, Z., Xu, J., Colvin, J. S., and Ornitz, D. M. (2002). Coordination of chondrogenesis and osteogenesis by fibroblast growth factor 18. Genes Dev. 16, 859–869. 140. Ohbayashi, N., Shibayama, M., Kurotaki, Y., Imanishi, M., Fujimori, T., Itoh, N., Takada, S. (2002). FGF18 is required for normal cell proliferation and differentiation during osteogenesis and chondrogenesis. Genes Dev. 16, 870–879. 141. Yu, K., Xu, J., Liu, Z., Sosic, D., Shao, J., Olson, E. N., Towler, D. A., Ornitz, D. M. (2003). Conditional inactivation of FGF receptor 2 reveals an essential role for FGF signaling in the regulation of osteoblast function and bone growth. Development 130, 3063–3074. 142. Shen, V., Kohler, G., Huang, J., Huang, S. S., and Peck, W. A. (1989). An acidic fibroblast growth factor stimulates DNA synthesis, inhibits collagen and alkaline phosphatase synthesis and induces resorption in bone. Bone Miner. 7, 407–408. 143. Simmons, H. A., and Raisz, L. G. (1991). Effects of acid and basic fibroblast growth factor and heparin on resorption of cultured fetal rat long bones. J. Bone Miner. Res. 6, 1301–1305. 144. Hurley, M. M., Lee, S. K., Raisz, L. G., Bernecker, P., and Lorenzo, J. (1998). Basic fibroblast growth factor induces osteoclast formation in murine bone marrow cultures. Bone 22, 309–316. 145. Marie, P. J. (2003). Fibroblast growth factor signalling controlling osteoblast differentiation. Gene 316, 23–32. 146. Kim, H. J., Kim, J. H., Bae, S. C., Choi, J. Y., Kim, H. J., Ryoo, H. M. (2003). The protein kinase C pathway plays a central role in the fibroblast growth factor-stimulated expression and transactivation activity of Runx2. J. Biol. Chem. 278, 319–326. 147. Xiao, G., Jiang, D., Gopalakrishnan, R., Franceschi, R. T. (2002). Fibroblast growth factor 2 induction of the osteocalcin gene requires MAPK activity and phosphorylation of the osteoblast transcription factor, Cbfa1/Runx2. J. Biol. Chem. 277, 36181–36187. 148. Kubota, K., Iseki, S., Kuroda, S., Oida, S., Iimura, T., Duarte, W. R., Ohya, K., Ishikawa, I., Kasugai, S. (2002). Synergistic effect of fibroblast growth factor-4 in ectopic bone formation induced by bone morphogenetic protein-2. Bone 31, 465–471. 149. Warren, S. M., Brunet, L. J., Harland, R. M., Economides, A. N., and Longaker, M. T. (2003). The BMP antagonist noggin regulates cranial suture fusion. Nature 422, 625–629. 150. Burke, D., Wilkes, D., Blundell, T. L., and Malcolm, S. (1998). Fibroblast growth factor receptors: Lessons from the genes. Trends Biochem. Sci. 23, 59–62. 151. Muenke, M., and Schell, U. (1995). Fibroblast-growth factor receptor mutations in human skeletal disorders. Trends Genet. 11, 308–313. 152. Wilkie, A. O. M., Slaney, S. P., Oldridge, M., Poole, M. D., Ashworth, G. J., Hockley, A. D., Hayward, R. D., David, D. J., Pulleyn, L. J., Rutland, P., Malcolm, S., Winter, R. M., and Reardon, W. (1995). Apert syndrome results from localized mutations of FGFR2 and is allelic with Crouzon syndrome. Nat. Genet. 9, 165–172. 153. Muenke, M., Schell, U., Hehr, A., Robin, N. H., Losken, H. W., Schinzel, A., Pulleyn, L. J., Rutland, P., Reardon, W., Malcolm, S., and Winter, R. M. (1994). A common mutation in the fibroblast growth factor receptor 1 gene in Pfeiffer syndrome. Nat. Genet. 8, 269–274. 154. Rutland, P., Pulleyn, L. J., Reardon, W., Baraitser, M., Hayward, R., Jones, B., Malcolm, S., Winter, R. M., Oldridge, M., Slaney, S. F.,

155.

156.

157.

158.

159.

160.

161.

162. 163. 164.

165.

166.

167.

168.

169.

Poole, M. D., and Wilkie, A. O. M. (1994). Identical mutations in the FGFR2 gene cause both Pfeiffer and Crouzon syndrome phenotypes. Nat. Genet. 9, 173–176. Jabs, E. W., Li, X., Scott, A. F., Meyers, G., Chen, W., Eccles, M., Mao, J.-I., Charnas, L. R., Jackson, C. E., and Jaye, M. (1994). Jackson-Weiss and Crouzon syndromes are allelic with mutations in fibroblast growth factor receptor 2. Nat. Genet. 8, 275–279. Readon, W., Winter, R. M., Rutland, P., Pulleyn, L. J., Jones, B. M., and Malcolm, S. (1994). Mutations in the fibroblast growth factor receptor 2 gene cause Crouzon syndrome. Nat. Genet. 8, 98–103. Rousseau, F., Bonaventure, J., Legeai-Mallet, L., Pelet, A., Rozet, J.-M., Maroteauz, P., Le Merrer, M., and Munnich, A. (1994). Mutations in the gene encoding fibroblast growth factor receptor-3 in achondroplasia. Nature 371, 252–254. Bellus, G. A., McIntosh, I., Smith, E. A., Aylsworth, A. S., Kaitila, L., Horton, W. A., Greenshaw, G. A., Hecht, J. T., and Francomano, C. A. (1995). A recurrent mutation in the tyrosine kinase domain of fibroblast growth factor receptor 3 causes hypochondroplasia. Nat. Genet. 10, 357–359. Tavormina, P. L., Shiang, R., Thompson, L. M., Zhu, Y.-Z., Wilkin, D. J., Lachman, R. S., Wilcox, W. R., Rimoin, D. L., Cohn, D. H., and Wasmuth, J. J. (1995). Thanatophoric dysplasia (types I and II) caused by distinct mutations in fibroblast growth factor receptor 3. Nat. Genet. 9, 321–328. ADHR Consortium (2000). Autosomal dominant hypophosphataemic rickets is associated with mutations in FGF23. Nat. Genet. 26, 345–348. Shimada, T., Muto, T., Urakawa, I., Yoneya, T., Yamazaki, Y., Okawa, K., Takeuchi, Y., Fujita, T., Fukumoto, S., and Yamashita, T. (2002). Mutant FGF-23 responsible for autosomal dominant hypophosphatemic rickets is resistant to proteolytic cleavage and causes hypophosphatemia in vivo. Endocrinology 143, 3179–3182. Harada, S., and Rodan, G. A. (2003). Control of osteoblast function and regulation of bone mass. Nature 423, 349–355. Westendorf, J. J., Kahler, R. A., and Schroeder, T. M. (2004). Wnt signaling in osteoblasts and bone diseases. Gene 341, 19–39. Moon, R. T., Kohn, A. D., De Ferrari, G. V., Kaykas, A. (2004). Wnt and beta-catenin signalling: Diseases and therapies. Nat. Rev. Gene. 5, 689–699. Kahler, R. A., and Westendorf, J. J. (2003). Lymphoid enhancer factor-1 and beta-catenin inhibit Runx2-dependent transcriptional activation of the osteocalcin promoter. J. Biol. Chem. 278, 11937–11944. Johnson, M. L., Gong, G., Kimberling, W., Recker, S. M., Kimmel, D. B., and Recker, R. B. (1997). Linkage of a gene causing high bone mass to human chromosome 11 (11q12–13). Am. J. Hum. Genet. 60, 1326–1332. Little, R. D., Carulli, J. P., Del Mastro, R. G., Dupuis, J., Osborne, M., Folz, C., Manning, S. P., Swain, P. M., Zhao, S. C., Eustace, B., Lappe, M. M., Spitzer, L., Zweier, S., Braunschweiger, K., Benchekroun, Y., Hu, X., Adair, R., Chee, L., FitzGerald, M. G., Tulig, C., Caruso, A., Tzellas, N., Bawa, A., Franklin, B., McGuire, S., Nogues, X., Gong, G., Allen, K. M., Anisowicz, A., Morales, A. J., Lomedico, P. T., Recker, S. M., Van Eerdewegh, P., Recker, R. R., and Johnson, M. L. (2002). A mutation in the LDL receptor-related protein 5 gene results in the autosomal dominant high-bone-mass trait. Am. J. Hum. Genet. 70, 11–19. Boyden, L. M., Mao, J., Belsky, J., Mitzner, L., Farhi, A., Mitnick, M. A., Wu, D., Insogna, K., and Lifton, R. P. (2002). High bone density due to a mutation in LDL-receptor-related protein 5. N. Engl. J. Med. 346, 1513–1521. Babij, P., Zhao, W., Small, C., Kharode, Y., Yaworsky, P. J., Bouxsein, M. L., Reddy, P. S., Bodine, P. V., Robinson, J. A., Bhat, B., Marzolf, J., Moran, R. A., and Bex, F. (2003). High bone mass in mice expressing a mutant LRP5 gene. J. Bone Miner. Res. 18, 960–974.

112 170. Van Wesenbeeck, L., Cleiren, E., Gram, J., Beals, R. K., Benichou, O., Scopelliti, D., Key, L., Renton, T., Bartels, C., Gong, Y., Warman, M. L., De Vernejoul, M. C., Bollerslev, J., and Van Hul, W. (2003). Six novel missense mutations in the LDL receptor-related protein 5 (LRP5) gene in different conditions with an increased bone density. Am. J. Hum. Genet. 72, 763–771. 171. Gong, Y., Slee, R. B., Fukai, N., Rawadi, G., Roman-Roman, S., Reginato, A. M., Wang, H., Cundy, T., Glorieux, F. H., Lev, D., Zacharin, M., Oexle, K., Marcelino, J., Suwairi, W., Heeger, S., Sabatakos, G., Apte, S., Adkins, W. N., Allgrove, J., Arslan-Kirchner, M., Batch, J. A., Beighton, P., Black, G. C., Boles, R. G., Boon, L. M., Borrone, C., Brunner, H. G., Carle, G. F., Dallapiccola, B., De Paepe, A., Floege, B., Halfhide, M. L., Hall, B., Hennekam, R. C., Hirose, T., Jans, A., Juppner, H., Kim, C. A., Keppler-Noreuil, K., Kohlschuetter, A., LaCombe, D., Lambert, M., Lemyre, E., Letteboer, T., Peltonen, L., Ramesar, R. S., Romanengo, M., Somer, H., Steichen-Gersdorf, E., Steinmann, B., Sullivan, B., Superti-Furga, A., Swoboda, W., van den Boogaard, M. J., Van Hul, W., Vikkula, M., Votruba, M., Zabel, B., Garcia, T., Baron, R., Olsen, B. R., and Warman, M. L., Osteoporosis-Pseudoglioma Syndrome Collaborative Group. (2001). LDL receptor-related protein 5 (LRP5) affects bone accrual and eye development. Cell 107, 513–523. 172. Kato, M., Patel, M. S., Levasseur, R., Lobov, I., Chang, B. H., Glass, D. A. 2nd, Hartmann, C., Li, L., Hwang, T. H., Brayton, C. F., Lang, R. A., Karsenty, G., and Chan, L. (2002). Cbfa1-independent decrease in osteoblast proliferation, osteopenia, and persistent embryonic eye vascularization in mice deficient in Lrp5, a Wnt coreceptor. J. Cell Biol. 157, 303–314. 173. Fujino, T., Asaba, H., Kang, M. J., Ikeda, Y., Sone, H., Takada, S., Kim, D. H., Ioka, R. X., Ono, M., Tomoyori, H., Okubo, M., Murase, T., Kamataki, A., Yamamoto, J., Magoori, K., Takahashi, S., Miyamoto, Y., Oishi, H., Nose, M., Okazaki, M., Usui, S., Imaizumi, K., Yanagisawa, M., Sakai, J., and Yamamoto, T. T. (2003). Low-density lipoprotein receptor-related protein 5 (LRP5) is essential for normal cholesterol metabolism and glucose-induced insulin secretion. Proc. Natl Acad. Sci. USA 100, 229–234. 174. Kokubu, C., Heinzmann, U., Kokubu, T., Sakai, N., Kubota, T., Kawai, M., Wahl, M. B., Galceran, J., Grosschedl, R., Ozono, K., and Imai, K. (2004). Skeletal defects in ringelschwanz mutant mice reveal that Lrp6 is required for proper somitogenesis and osteogenesis. Development 131, 5469–5480. 175. Niemann, S., Zhao, C., Pascu, F., Stahl, U., Aulepp, U., Niswander, L., Weber, J. L., and Muller, U. (2004). Homozygous WNT3 mutation causes tetra-amelia in a large consanguineous family. Am. J. Hum. Genet. 74, 558–563. 176. Koay, M. A., Woon, P. Y., Zhang, Y., Miles, L. J., Duncan, E. L., Ralston, S. H., Compston, J. E., Cooper, C., Keen, R., Langdahl, B. L., MacLelland, A., O’Riordan, J., Pols, H. A., Reid, D. M., Uitterlinden, A. G., Wass, J. A., and Brown, M. A. (2004). Influence of LRP5 polymorphisms on normal variation in BMD. J. Bone Miner. Res. 19, 1619–1627. 177. Tuan, R. S. (2003). Cellular signaling in developmental chondrogenesis: N-cadherin, Wnts, and BMP-2. J. Bone Joint Surg. Am. 85-A Suppl 2, 137–141. 178. Loughlin, J., Dowling, B., Chapman, K., Marcelline, L., Mustafa, Z., Southam, L., Ferreira, A., Ciesielski, C., Carson, D. A., and Corr, M. (2004). Functional variants within the secreted frizzled-related protein 3 gene are associated with hip osteoarthritis in females. Proc. Natl Acad. Sci. USA 101, 9757–9762. 179. Betsholtz, C. (2003). Biology of platelet-derived growth factors in development. Birth Defects Res. C. Embryo Today 69, 272–285. 180. Seifert, R. A., Hart, C. E., Phillips, P. E., Forstrom, J. W., Ross, R., Murray, M. J., and Bowen-Pope, D. F. (1989). Two distinct subunits associate to create isoform-specific platelet-derived growth factor receptors. J. Biol. Chem. 264, 8771–8778.

PHILIPPA HULLEY ET AL. 181. Centrella, M., McCarthy, T. L., and Canalis, E. (1989). Platelet derived growth factor enhances deoxyribonucleic acid and collagen synthesis in osteoblast-enriched cultures from fetal rat parietal bone. Endocrinology 125, 13–19. 182. Centrella, M., McCarthy, T. L., Kusmik, W. F., and Canalis, E. (1991). Relative binding and biochemical effects of heterodimeric and homodimeric isoforms of platelet-derived growth factor in osteoblast-enriched cultures from rat fetal bone. J. Cell Physiol. 147, 420–426. 183. Gilardetti, R. S., Chaibi, M. S., Stroumza, C. J., Williams, S. R., Antoniades, H. N., Carnes, D. C., and Graves, D. T. (1991). High affinity binding of PDGF-AA and PDGF-BB to normal human osteoblastic cells and modulation by interleukin-1. Am. J. Physiol. 261, C980–C985. 184. Pfeilschifter, J., Krempien, R., Naumann, A., Gronwald, R. G. K., Hoppe, J., and Ziegler, R. (1992). Differential effects of platelet-derived growth factor isoforms on plasminogen activator activity in fetal rat osteoblasts due to isoform-specific receptor functions. Endocrinology 130, 2059–2066. 185. Horner, A., Bord, S., Kemp, P., Grainger, D., and Compston, J. E. (1996). Distribution of platelet-derived growth factor (PDGF) A chain mRNA, protein, and PDGF-a receptor in rapidly forming bone. Bone 19, 353–362. 186. Yu, X. H., Hsieh, S. C., Bao, W., and Graves, D. T. (1997). Temporal expression of PDGF receptors and PDGF regulatory effects on osteoblastic mineralizing cultures. Am. J. Physiol. 41, C1709–C1716. 187. Hsieh, S. C., and Graves, D. T. (1998). Pulse application of plateletderived growth factor enhances formation of a mineralizing matrix while continuous application is inhibitory. J. Cell. Biochem. 69, 169–180. 188. Nash, T. J., Howlett, C. R., Martin, C., Steele, J., Johnson, K. A., and Hicklin, D. J. (1994). Effect of platelet-derived growth factor in tibial osteotomies in rabbits. Bone 15, 203–208. 189. Mitlak, B. H., Finkelman, R. D., Hill, E. L., Li, J., Martin, B., Smith, T., D’Andrea, M., Antoniades, H. N., and Lynch, S. E. (1996). The effect of systemically administered PDGF-BB on the rodent skeleton. J. Bone Miner. Res. 11, 238–247. 190. Tashjian, A., Hohmann, E., Antoniades, H., and Levine, L. (1982). Platelet-derived growth factor stimulates bone resorption via a prostaglandin-mediated mechanism. Endocrinology 111, 118–124. 191. Cochran, D. L., Rouse, C. A., Lynch, S. E., and Graves, D. T. (1993). Effects of platelet-derived growth factor isoforms on calcium release from neonatal mouse calvariae. Bone 14, 53–58. 192. Franchimont, N., and Canalis, E. (1995). Platelet-derived growth factor stimulates the synthesis of interleukin-6 in cells of the osteoblast lineage. Endocrinology 136, 5469–5474. 193. Varghese, S., Delany, A. M., Liang, L., Gabbitas, B., Jeffrey, J. J., and Canalis, E. (1996). Transcriptional and posttranslational regulation of interstitial collagenase by platelet-derived growth factor BB in bone cell cultures. Endocrinology 137, 431–437. 194. Ng, K. W., Partridge, N. C., Niall, H., and Martin, T. J. (1983). Epidermal growth factor receptors in clonal lines of a rat osteogenic sarcoma and in osteoblast-rich rat bone cells. Calcif. Tissue Int. 35, 298–303. 195. Shupnik, M. A., and Tashjian, A. H. (1982). Epidermal growth factor and phorbol ester actions on human osteosarcoma cells. J. Biol. Chem. 257, 12161–12164. 196. Yokota, K., Kusaka, M., Ohshima, T., Yamamoto, S., Kurihara, N., Yoshino, T., and Kumagawa, M. (1986). Stimulation of prostaglandin E2 synthesis in cloned osteoblastic cells of mouse (MC3T3-E1) by epidermal growth factor. J. Biol. Chem. 261, 15410–15415. 197. Qin, L., Tamasi, J., Raggatt, L., Li, X., Feyen, J. H., Lee, D. C., Dicicco-Bloom, E., and Partridge, N. C. (2004). Amphiregulin is a novel growth factor involved in normal bone development and in the cellular response to parathyroid hormone stimulation. J. Biol. Chem. 280, 3974–3981.

Chapter 6 Growth Factors 198. Harrison, J. R., Lorenzo, J. A., Kawaguchi, H., Raisz, L. G., and Pilbeam, C. (1994). Stimulation of prostaglandin E2 production by interleukin-1 alpha and transforming growth factor alpha in osteoblastic MC3T3-E1 cell. J. Bone Miner. Res. 9, 817–823. 199. Raisz, L. G., Simmons, H. A., Sandberg, A. L., and Canalis, E. (1980). Direct stimulation of bone resorption by epidermal growth factor. Endocrinology 107, 207–212. 200. Ibbotson, K. J., Harrod, J., Gowen, M., D’Souza, S., Smith, D. D., Winkler, M. E., Derynck, R., and Mundy, G. R. (1985). Stimulation of bone resorption in vitro by synthetic transforming growth factor alpha. Science 228, 1007–1009. 201. Stern, P. H., Krieger, N. S., Nissenson, R. A., Williams, R. D., Winkler, M. E., Derynck, R., and Strewler, G. J. (1985). Human transforming growth factor alpha stimulates bone resorption in vitro. J. Clin. Invest. 76, 2016–2020. 202. Yates, A. J., Boyce, B. R., Favarato, G., Aufdemorte, T. B., Marcelli, C., Kester, M. B., Walker, R., Langton, B. C., Bonewald, L. F., and Mundy, G. R. (1992). Expression of human transforming growth factor alpha by Chinese hamster ovarian tumors in nude mice causes hypercalcemia and increased osteoclastic bone resorption. J. Bone Miner. Res. 7, 847–853. 203. Harada, S-L., Nagy, J. A., Sullivan, K. A., Thomas, K. A., Endo, N., Rodan, G. A., and Rodan, S. B. (1994). Induction of vascular endothelial growth factor by prostaglandin E2 and E1 in osteoblasts. J. Clin. Invest. 93, 2490–2496.

113 204. Wang, D. S., Yamazaki, K., Nohtomi, K., Shizume, K., Ohsumi, K., Shibuya, M., Demura, H., and Sato, K. (1996). Increase of vascular endothelial growth factor mRNA expression by 1,25-dihydroxy vitamin 03 in human osteoblast-like cells. J. Bone Miner. Res. 11, 472–479. 205. Goad, D. L., Rubin, J., Wang, H., Tashjian, A. H., and Patterson, C. (1996). Enhanced expression of vascular endothelial growth factor in human SaOS-2 osteoblast-like cells and murine osteoblasts induced by insulin-like growth factor I. Endocrinology 137, 2262–2268. 206. Street, J., Bao, M., deGuzman, L., Bunting, S., Peale, Jr, F. V., Ferrara, N., Steinmetz, H., Hoeffel, J., Cleland, J. L., Daugherty, A., van Bruggen, N., Redmond, H. P., Carano, R. A. D., and Filvaroff, R. H. (2002). Vascular endothelial growth factor stimulates bone repair by promoting angiogenesis and bone turnover. Proc. Natl Acad. Sci. USA 99, 9656–9661. 207. Wang, D. S., Miura, M., Demura, H., and Sato, K. (1997). Anabolic effects of 1,25-dihyroxyvitamin D3 on osteoblasts are enhanced by vascular endothelial growth factor produced by osteoblasts and by growth factors produced by endothelial cells. Endocrinology 138, 2953–2962. 208. Maes, C., Stockmans, I., Moermans, K., Van Looveren, R., Smets, N., Carmeliet, P., Bouillon, R., and Carmeliet, G. (2004). Soluble VEGF isoforms are essential for establishing epiphyseal vascularization and regulating chondrocyte development and survival. J. Clin. Invest. 113, 188–199.

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Chapter 7

Prostaglandins and Proinflammatory Cytokines Lawrence G. Raisz, M.D. Joseph A. Lorenzo, M.D.

Interim Director, Musculoskeletal Institute, Board of Trustees Distinguished Professor of Medicine, University of Connecticut Health Center, 263 Farmington Avenue, MC-3805, Farmington, CT 06030 Director, Bone Biology Research, Professor of Medicine, University of Connecticut Health Center, 263 Farmington Avenue, MC 1317, Farmington, CT 06030–1317.

IV. The Role that Proinflammatory Cytokines have in Bone and Cartilage Metabolism References

I. Introduction II. Prostaglandins III. The Role that Cytokines have in Osteoclast Formation and Function

I. INTRODUCTION

in skeletal tissues. The source of prostaglandins, arachidonate, may be itself a mediator, as well as leukotrienes, which are products of lipoxygenase [2, 3]. Prostaglandins are multifunctional regulators in that they have both stimulatory and inhibitory effects on bone formation and bone resorption as well as biphasic effects on cartilage. Largely because bone has been studied more extensively, a reasonable description of the pattern of physiologic and pathologic responses can now be provided, although there are still many gaps. The role of prostaglandins in cartilage has been less completely studied. Interpretation is further confounded by the fact that many different cell types, species, and in vitro and in vivo conditions have been examined. Identification of specific receptors for prostaglandins, particularly prostaglandin E2 (PGE2), which is the most abundant product in bone cells, has provided a substantial amount of new information. Both transgenic mouse models in which specific receptors

This chapter has been extensively revised since the previous edition to accommodate the remarkable amount of new information that has been developed during the last 5 years in studies of prostaglandins and cytokines in bone and cartilage. Only key references from the previous edition have been retained and the reader is referred to that edition for background information on earlier studies.

II. PROSTAGLANDINS Prostaglandins are critical local regulators of skeletal metabolism that have been shown to play key roles in both physiologic regulation and pathologic responses [1]. In addition to prostaglandins, which are produced by cyclooxygenase, other lipid mediators can be produced Dynamics of Bone and Cartilage Metabolism

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have been deleted and selective receptor agonists and antagonists are now available for study.

A. Regulation of Prostaglandin Synthesis Many types of cells in the skeleton can produce prostaglandins and this production is highly regulated. Prostaglandin production in osteoblastic cells can be stimulated by cytokines, growth factors, nitric oxide, and mechanical forces. Prostaglandins themselves increase their own production (auto-regulation) [4]. This has also been shown for prostaglandin production in chondrocytes. In addition hematopoietic cells, particularly cells of the monocyte/macrophage lineage, and synovial cells can play a prominent role in prostaglandin production in inflammatory disorders. Regulation of prostaglandin production is largely through changes in the inducible form of cyclooxygenase (COX-2). However, the constitutive cyclooxygenase (COX-1) may play a role, particularly in rapid initial responses in which release of arachidonate by phospholipase leads to immediate production of prostaglandins. This may occur in response to both mechanical forces and inflammation [5]. However, the full response of skeletal tissue to mechanical force and inflammation as well as in fracture repair requires COX-2 [6–11]. Recent studies have suggested that the response of the skeleton to changes in ion concentration might also be prostaglandin dependent. High calcium concentrations can induce COX-2 [12]. The stimulation of bone resorption in organ culture that occurs in acid culture medium appears to be prostaglandin dependent [13, 14]. The physiologic importance of this is unknown. However, prostaglandins can inhibit osteoclast function [15]. Thus, if the high concentrations of calcium and hydrogen ions that are produced during osteoclastic bone resorption can induce COX-2, this might provide a mechanism for stopping their activity and, hence, limiting the depth of resorption cavities. Studies using mice in which the COX-2 gene has been deleted clearly demonstrate a critical role for this enzyme in the stimulation of bone resorption by a number of agonists [10] in fracture healing [6, 9], in the osteolysis in response to implants [8], and probably also in heterotopic ossification, based on clinical data [16], as well as on the diminished response to BMP-2 implants in COX-2 knockout animals [11]. The major product of COX-2 in bone cells and probably also in cartilage cells appears to be PGE2 and, hence, membrane-associated PGE2 synthase is also critical for bone resorption and other responses [17]. Studies on the induction of COX-2 have suggested that both the protein kinase-A (PKA) and protein kinase-C (PKC) pathways are involved. However, studies using

selective prostaglandin receptor agonists have supported a greater role for cAMP and PKA activation, particularly through the prostaglandin EP2 receptor [18]. The regulation of COX-2 in cartilage has been less well studied, but chondrocytes appear to show similar responses to those of osteoblasts. Cytokines such as IL-1, growth factors, and mechanical forces can all induce COX-2 in chondrocytes [19–22]. The role of COX-2 in the response of the skeleton to mechanical forces has received increasing attention [23]. A cellular model for mechanical force, the application of fluid shear stress to osteoblast or osteocyte cell cultures, has confirmed the critical role of COX-2 [7, 24]. The anabolic response to mechanical loading can be blocked by selective COX-2 inhibitors [25, 26]. The effects of fluid shear appear to be mediated through cAMP and an ERK signaling pathway [12, 27] and to increased function of gap junctions [28–30]. Signal transduction can occur through cAMP, but likely also through other pathways.

B. Prostaglandins and Bone Resorption Studies using receptor knockout animals, selective agonists, and specialized cell culture models have helped to identify the multiple mechanisms and target cells of prostaglandin bone resorption. Probably the most important effect is the ability of PGE2 to stimulate RANKL production in cells of the osteoblast lineage and perhaps other marrow cells [31]. This effect can be diminished by knocking out either the EP2 or the EP4 receptor, but the latter appears to have a somewhat greater role [32–36]. In fetal rat organ cultures EP4 agonists stimulate resorption, while EP2 agonists are ineffective [37]. In addition to stimulating RANKL, PGE2 may inhibit the production of the decoy receptor osteoprotegerin (OPG), although this has not been found in all of the models tested [38–40]. While the EP2 receptor plays a role in the response of osteoblasts to PGE2, it also has a role in cells of the hematopoietic lineage that give rise to osteoclast [36]. Addition of PGE2 can increase osteoclast production by spleen cells treated with maximally effective concentrations of macrophage colony stimulating factor (M-CSF) and RANKL [41, 42]. Knockout of EP2 receptors in spleen cells reduces the number of osteoclast produced in response to PGE2 [36, 42]. In vivo studies have shown that the response to lipopolysaccharide is decreased in EP4 knockout mice, while the hypercalcemic effect of PGE2 is reduced in EP2 receptor knockout mice [40, 43]. In addition to their predominant effect to stimulate the formation, differentiation, and extend the lifespan

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of osteoclast, prostaglandins have a direct inhibitory effect on osteoclast function. This effect may be mediated by the EP4 receptor [15].

chondrocytes, largely through induction of COX-2 [21, 65, 67, 68]. In addition to a critical role for COX-2, the microsomal PGE synthase is required for these responses [69]. The role of specific prostaglandin receptors has been studied in collagen antibody-induced arthritis in mice. Animals lacking the EP4 receptor show reduced bone destruction and cartilage collagen breakdown in this model [70]. Prostaglandins have complex effects on chondrocyte replication, differentiation and cell death [71]. In an experimental model of osteoarthritis dual inhibition of 5 lipoxygenase and cyclooxygenase resulted in decreased cell death [72]. In cultured chondrocytes from humans with osteoarthritis, nitric oxide induction of cell death is enhanced by induction of COX-2 and production of PGE2. On the other hand PGE2 has been shown to stimulate production and degradation of proteoglycan and increase collagen synthesis in rat chondrocyte cultures [73, 74]. These effects may be mediated by activation of the EP2 and EP4 receptors [75].

C. Prostaglandins and Bone Formation Exogenous PGE can stimulate both endosteal and periosteal bone formation, that is, both modeling and remodeling of the skeleton. This effect persists even in old experimental animals [44, 45]. The role of cell replication in this response is not entirely clear. Under different conditions PGE2 can inhibit or stimulate osteoblastic cell replication [46, 47]. On the other hand, differentiation and colony formation in osteoblastic cell cultures is consistently enhanced by PGE2 [48–50]. Here again, there is a biphasic effect of PGE2, in that collagen synthesis is inhibited in differentiated osteoblasts and in some osteoblastic cell lines [51]. The anabolic effects of PGE2 may be mediated by both EP2 and EP4 receptors. Agonists for both receptors can increase collagen synthesis in fetal rat calvarial organ cultures [37]. A selective EP2 agonist was found to stimulate local bone formation and enhance fracture healing [52, 53]. Inhibitors of the EP4 receptor can impair the anabolic response to PGE2 [54, 55] and selective agonists can increase bone formation and prevent bone loss [56]. There are also data indicating that prostaglandins of the F series can be anabolic in bone [57]. The mechanism of the anabolic effect of PGE2 is not fully understood. The enhancement of osteoblast differentiation appears to be at least in part cAMP dependent, and it probably involves stimulation of the production of endogenous growth factors [49]. Prior studies suggested that increased activity of IGF-1 or VEGF might mediate the anabolic effect [58, 59]. Stimulation of production or enhancement of the response to BMP-2 and BMP-7 have been implicated [60, 61]. Prostaglandins, particularly PGF2α, can also stimulate FGF production, thus leading to a new amplification loop, since FGF in turn can stimulate COX-2 [62, 63]. A dual pathway is likely, for example PGE2 stimulates bone sialoprotein (BSP) expression both through cAMP and FGF2 response elements in the BSP promoter [64].

E. Clinical Implications While there is some evidence that prostaglandins may play important roles in diseases of bone and cartilage, their relative importance compared to other regulators and the precise pathways by which they act are not understood [76–78]. For example, the impaired fracture healing in COX-2-deficient or NSAID-treated animals and humans could be due to effects on platelets, vessels, inflammatory mediators, or fibroblastic cells in the initial phase, as well as to effects on cartilage and bone in subsequent steps. The same may apply for their role in heterotopic ossification, prosthetic loosening and the bone loss associated with immobilization and inflammation. There is direct evidence for an anabolic effect of prostaglandins in infants given infusions of PGE1 and more recently a similar response has been observed in an older patient who developed hypertrophic osteoarthropathy after PGE1 infusion for congestive heart failure [79]. Impact loading can increase PGE2 production in human bone [80]. A role in the pathogenesis of osteoporosis has been suggested based on epidemiologic studies of anti-NSAIDs, as well as some animal studies, but the data are contradictory [81–83].

D. Prostaglandins and Cartilage The fact that NSAIDs can not only inhibit inflammatory responses but may also reduce cartilage degradation in humans and in animal models of arthritis has led to extensive study of the role of prostaglandins in cartilage and adjacent synovial tissue [65, 66]. Cytokines and nitric oxide can increase prostaglandin production in

III. THE ROLE THAT CYTOKINES HAVE IN OSTEOCLAST FORMATION AND FUNCTION Osteoclast are multinuclear giant cells with the unique ability to efficiently resorb both the mineral and organic

118 components of bone [84]. They form from mononuclear cells, which derive from myeloid lineage hematopoietic precursors. The role that cytokines play in the regulation of osteoclast has been made clearer in recent years by the discovery of a tumor necrosis factor superfamily member and its receptors that are critical for osteoclast formation, activity, and survival [85, 86]. This cytokine was named receptor activator of nuclear factor κB ligand (RANKL, TNFSF11) [87]. It was originally identified as a product of activated T-lymphocytes [87, 88] and it exists either as a membrane-bound or a soluble product whose form is determined by both alternate gene transcription and cleavage of the membranebound form by proteases [89]. Stimulators of resorption enhance production of RANKL, which binds to its receptor RANK (TNFRSF11A) on osteoclast precursor cells [86]. This interaction initiates a series of second messenger steps that mediate osteoclast formation from mononuclear precursors, resorptive activity, and osteoclast survival. In addition, a variety of cells in bone produce a soluble decoy receptor, osteoprotegerin (OPG, TNFRSF11B) [90], which binds RANKL and prevents its interaction with RANK. Thus, OPG is a negative regulator of osteoclastic resorption. Some hormonal stimulators of bone resorption, such as parathyroid hormone [91] and 1,25 (OH)2 vitamin D [92], increase the production of RANKL and inhibit the production of OPG in stromal and osteoblastic cells. This reciprocal regulation of RANKL and OPG likely enhances the resorptive activity of these agents. Other cytokines such as interleukin-1 and tumor necrosis factor [93] stimulate RANKL but have either no effect or are stimulators of OPG production [94]. The second cytokine that is critical for osteoclast differentiation is macrophage colony-stimulating factor (M-CSF or CSF-1) [95]. M-CSF was originally identified as a regulator of macrophage differentiation. However, it was subsequently found that the defect in the op/op mouse, which presents with defective osteoclast formation and osteopetrosis, is caused by a mutation in the M-CSF gene [96, 97]. This point mutation introduces a stop codon in the sequence, which prevents expression of the protein. Subsequent studies have shown that M-CSF is a mitogen for early myeloid precursor cells, which regulates the lineage commitment of myeloid precursor cells towards osteoclast [95]. Exposure of myeloid precursors to M-CSF also induces RANK expression [98]. Like RANKL, M-CSF enhances the survival of mature osteoclast [99]. However, in contrast to the actions of RANKL, it cannot initiate the terminal differentiation of osteoclast precursors. In cell culture, osteoclast precursor cells need continuous exposure to both M-CSF and RANKL to form fully differentiated mature osteoclast. PGE2 treatment of

Lawrence G. Raisz and Joseph A. Lorenzo

murine osteoblastic cells had no effect on M-CSF mRNA expression [40]. The receptor for M-CSF is the product of the protooncogene c-fms [100]. This receptor, which is also known as CSF-1R, mediates responses through activation of tyrosine kinase [95]. Recently, mice lacking CSF-1R were generated and their phenotype was similar to that of the op/op mouse [100]. As with a number of immune cells, costimulatory molecules are involved in osteoclast formation. Two costimulatory receptors have been identified as being involved in osteoclast formation. These are TREM-2 [101] and OSCAR [102], which are found on the surface of osteoclast precursor cells and for which cognate ligands are unknown. Costimulatory receptors interact with activator proteins, containing an immunotyrosinebased activation motif (ITAM), to mediate their responses [103]. In osteoclast TREM-2 associates with DAP12 [104] while OSCAR associates with FcRγ [105]. If both DAP12 and FcRγ are absent in mice, osteoclast formation is markedly inhibited [105–107].

IV. THE ROLE THAT PROINFLAMMATORY CYTOKINES HAVE IN BONE AND CARTILAGE METABOLISM The cytokines that have been most extensively studied for their ability to regulate bone and cartilage function are interleukin-1 (IL-1), tumor necrosis factor (TNF), and interleukin-6 (IL-6). These agents are proinflammatory cytokines that have potent effects in a variety of in vivo and in vitro assays. This section will focus on their actions in bone and cartilage. In addition, the role of other IL-6 family member cytokines will be briefly reviewed.

A. Effects of Interleukin-1 on Bone Interleukin-1 (IL-1) is encoded by two separate gene products, α and β, which have identical activities [108]. IL-1 was the first polypeptide mediator of immune cell function that was found to regulate bone resorption [109, 110]. It also increased prostaglandin synthesis in bone [110], an effect that may account for some of its resorptive activity. This response is mediated through its ability to increase production of cyclooxygenase 2 [111] and the effect does not appear to require nitric oxide synthesis [112]. IL-1 is the most potent peptide stimulator of in vitro bone resorption that has yet been identified [110]

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and it also has potent in vivo effects [113]. The in vitro effects of IL-1 on bone formation appear to be inhibitory [114–116]; although, it does stimulate DNA synthesis in both bone organ cultures and primary cultures of human bone cells [116, 117]. IL-1 is produced in bone [118] and IL-1 activity is present in bone marrow serum [119, 120]. Macrophages are a likely source of local bone IL-1 [121]. In addition, osteoblasts and osteoclasts may also produce it [122, 123]. IL-1 also appears to directly stimulate mature osteoclast to resorb [124]. A natural inhibitor to IL-1, IL-1 receptor antagonist (IL-1ra), is an analog of IL-1 that binds but does not activate the type 1 IL-1 receptor [125–127]. IL-1ra blocks the ability of IL-1 to stimulate resorption and PGE2 production in bone organ cultures [128]. Increased release of IL-1ra has been demonstrated from peripheral blood monocytes (PBM) of both normal and osteoporotic postmenopausal women [129]. Decreased levels of IL-1ra are also reported in the bone marrow of some women with a rapid-loss form of osteoporosis [130]. There are two known receptors for IL-1, type I and type II [131]. All known biologic responses to IL-1 appear to be mediated exclusively through the type I receptor [132]. Signaling through type I receptors involves activation of NF-κB and specific TNF receptor associated factors (TRAFs) [133, 134]. IL-1 receptor type I requires interaction with a second protein, interleukin-1 receptor accessory protein (IRAcP), to generate postreceptor signals [135–137]. IRAcP induces activation of interleukin-1 receptor-associated kinase (IRAK) [52]. Phosphorylated IRAK binds TRAF6 with subsequent downstream signaling. IL-1 receptor type II appears to have no agonist activity. Rather, there is convincing evidence that it is a decoy receptor, which prevents activation of IL-1 type I receptors [138]. IL-1 receptor type II can also synergize with IL-1ra to inhibit activation of the IL-1 receptor type I [139]. The role of IL-1 in estrogenmediated bone loss had been suspected from studies which demonstrated that the IL-1 antagonist IL-1ra blocked ovariectomy-induced bone loss [140], and was confirmed by studies demonstrating that mice lacking the IL-1 receptor type I (IL-1R1 −/−) did not lose bone mass with ovariectomy [141]. Studies of the regulation of IL-1 by estrogens in humans have been inconsistent. One group demonstrated that IL-1 levels in serum were increased 4 weeks after ovariectomy [142]. However, others failed to find a correlation between serum IL-1α, IL-1β, or IL-1ra levels and indices of bone turnover in either pre- or post-menopausal women [143] or between osteoporotic women and normals [144].

Production of IL-1 receptor type II, the decoy IL-1 receptor, is up-regulated by estrogen treatment [145, 146]. It is possible that estrogen regulates bone mass by increasing IL-1 receptor type II expression independent of any direct effects on IL-1 production. This response would decrease the ability of bone cells to respond to IL-1 and blunt the potent catabolic effects of IL-1 on bone mass.

B. Effects of Interleukin-1 on Cartilage IL-1 is a potent regulator of cartilage cell function. It inhibits cartilage cell replication, colony formation in soft agar and proteoglycan synthesis [147, 148], while it stimulates production of matrix metalloproteinases, which degrade cartilage collagen [149]. IL-1 also inhibited the production of cartilage-specific collagens [150] and it regulated expression of 1,25-dihydroxyvitamin D receptors and type X collagen production in growth plate chondrocytes but had little effect on type II collagen synthesis [151]. IL-1 may be involved in the regulation of growth plate cartilage since it is made by these cells [152]. Production of IL-1 in rat growth plate cartilage was altered by the vitamin D status of the animals [153]. This mechanism may be involved in growth. However, it is not critical since mice that cannot respond to IL-1 grow normally [154]. In vitro, IL-1 and tumor necrosis factor (TNF) act synergistically to inhibit the growth of cultured long bones and this response appears mediated by decreased chondrocyte proliferation and increased apoptosis [155].

C. Effects of Tumor Necrosis Factor on Bone Like IL-1, tumor necrosis factor (TNF) is a family of two related polypeptides (α and β) that are separate gene products [156–160]. TNFα and β have similar biologic activities and are both potent stimulators of bone resorption [110, 161, 162]. TNF also enhances the formation of osteoclast-like cells in bone marrow culture [163]. Therefore, it appears to regulate osteoclast precursor cell differentiation. TNFα was produced by human osteoblast-like cell cultures [164] and its production was stimulated by IL-1, GM-CSF and lipopolysaccharide, but not by PTH, 1,25(OH)2 vitamin D, or calcitonin. Recently, TNF was shown to stimulate osteoclast formation directly in an in vitro culture system [124, 165]. This effect was independent of RANK actions since it occurred in cells from RANK-deficient mice. However, the functional significance of this finding is questionable because in vivo administration of TNF to

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RANK-deficient mice caused only an occasional osteoclast to form [166]. Like IL-1, TNF binds to two cell surface receptors, the TNF receptor 1 or p55 and the TNF receptor 2 or p75 [167]. In contrast to IL-1, both receptors transmit biologic responses. Mice deficient in either or both TNF receptors have been made [168–170]. All of these animals appear healthy and breed normally, but lack selective immune responses. Bones from mice that lack either IL-1 receptor type I or TNF receptor 1 are similar to those of wild-type mice, except for a 36% increase in the total bone area of calvaria from IL-1 receptor type I-deficient mice, compared to wild-type animals [154]. Mice deficient in TNF (TNF −/−) fail to lose bone mass after ovariectomy [171]. This finding appears significant because production of TNF by T lymphocytes appears responsible for some of the bone loss that occurs in mice after ovariectomy [172]. TNF appears to have direct effects on osteoclast formation independent of RANKL [124]. However, this response requires that the osteoclast precursor cells be previously exposed to RANKL [173].

D. Effects of Tumor Necrosis Factor on Cartilage Like IL-1, TNFα stimulates production of degradative enzymes in cartilage and inhibits cartilage synthesis [174, 175]. Both TNF and IL-1 are implicated as mediators of inflammatory arthritis [176]. In costal chondrocytes TNF decreased alkaline phosphatase and DNA synthesis [177]. In vivo fracture healing in rat ribs was inhibited by TNFα treatment [178]. This effect was due to a decrease in cartilaginous callus formation and, possibly, a decrease in the differentiation of mesenchymal cells into chondroblasts.

E. Effects of Interleukin 6 on Bone IL-6, like IL-1 and TNF, has a wide variety of effects on immune cell function and on the replication and differentiation of other cell types [179, 180]. The receptor for IL-6 is composed of two parts: a specific IL-6 binding protein (IL-6 receptor), which can be either membrane bound or soluble, and gp-130, an activator protein that is common to a number of cytokine receptors [181]. Production of the specific IL-6 receptor may be regulated, since it is not expressed on bone cells at all times [182]. The ability of IL-6 to stimulate bone resorption in vitro is variable and depends on the assay system that is examined [183–186]. The reasons for these discrepancies are unknown. It appears that a major effect of IL-6 is to

regulate the differentiation of osteoclast progenitor cells into mature osteoclast [187, 188]. In an in vitro model of osteoclast differentiation from human bone marrow, IL-6 was found to be a potent stimulus of new osteoclast-like cell development by a mechanism that was also dependent on IL-1 [189]. In contrast, in a murine in vitro system, IL-6 was only effective when soluble IL-6 receptor was added to the cultures [182]. Implantation of Chinese hamster ovary (CHO) cells, which were genetically engineered to express murine IL-6, into nude mice produced a syndrome of hypercalcemia, cachexia, leukocytosis, and thrombocytosis [190]. The effects of IL-6 on resorption in this model were weak, but were significantly potentiated when the animals were also treated with PTHrP [191]. A major effect of IL-6 in these in vivo models was to increase the number of early osteoclast precursors (CFU-GM) in the marrow. IL-6 may mediate some of the increased bone resorption and bone pathology that is seen in the clinical syndromes of Paget’s disease [192], hypercalcemia of malignancy [193], fibrous dysplasia [194], giant cell tumors of bone [195], and Gorham–Stout disease [196]. Ovariectomy-induced bone loss in mice has been associated with increased IL-6 production [197] and mice deficient in IL-6 failed to lose bone after ovariectomy [198].

F. Effects of Interleukin 6 on Cartilage IL-6 is produced in cartilage in response to treatment with inflammatory cytokines such as IL-1 and TNF [176]. IL-6 production is increased by treatment with growth hormone or insulin-like growth factor-1 (IGF-1) in growth plate chondrocyte cultures [199]. However, in human articular cartilage cultures IGF-1 had no effect on IL-6 production [200]. IL-6 is not critical for skeletal growth since IL-6 deficient mice grow normally [201]. IL-6 also enhances cartilage production of tissue inhibitor of metalloproteinase 1 (TIMP-1), a factor that blocks the activity of metalloproteinases, which degrade cartilage [202, 203] (see also Chapter 11).

G. Effects of Other Interleukin-6 Family Members on Bone IL-6 is a member of a group of cytokines that share the gp-130 activator protein in their receptor complex [204, 205]. Each family member utilizes unique ligand receptors to generate specific binding. Besides IL-6, the cytokines of this family include interleukin 11 (IL-11), leukemia inhibitory factor (LIF), oncostatin M (OSM),

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ciliary neurotrophic factor (CNTF), and cardiotrophin-1 (CT-1). Except for CT-1, receptors for all other IL-6 family members have been detected in bone marrow stromal or osteoblastic cells [206]. In addition, IL-11-specific receptors are present in osteoclast-like cells that are generated in vitro. Signal transduction through these receptors utilizes the JAK/STAT pathway [181]. Leukemia inhibitory factor (LIF) is produced by bone cells in response to a number of resorption stimuli [207–209]. However, production can be variable and depends on the model system that is being studied. The effects of LIF on bone resorption are also variable. In a number of in vitro model systems, LIF stimulated resorption by a mechanism that is prostaglandin dependent [210] while, in other in vitro assays, it had inhibitory effects [211, 212]. Up-regulation of RANKL production seems to be the mechanism by which LIF stimulates resorption in bone organ cultures, although the role of prostaglandins in this model is unknown [213]. Mice that lack the specific LIF receptor and, hence, cannot respond to LIF, have reduced bone volume and a six-fold increase in osteoclast numbers [214]. The role of IL-6 family members in osteoclast formation has been re-examined in the light of data demonstrating that mice lacking the gp-130 activator protein have an increased number of osteoclasts compared to wild-type animals [215]. Since gp-130 is an activator of signal transduction for all members of the IL-6 family, this result argues that at least some IL-6 family members inhibit osteoclast formation and bone resorption. The mechanism by which these effects occur were recently investigated in mice that were deficient in signaling through either the gp-130-mediated STAT 1/3 pathway or the SHP2/Ras/MAPK pathway [216]. In mice deficient in STAT 1/3 signaling premature growth plate closure reduced bone length. In SHP2/Ras/MAPK deficient mice osteoclast were increased. Crossing IL-6 deficient mice with mice deficient in gp-130-mediated SHP2/Ras/MAPK signaling decreased osteoblasts without affecting the increase in osteoclast. These results demonstrate that the gp-130-mediated STAT 1/3 pathway is a major regulator of chondrocyte differentiation while gp-130mediated SHP2/Ras/MAPK signaling is an inhibitor of osteoclastogenesis.

2. Bordin, L., Priante, G., Musacchio, E., Giunco, S., Tibaldi, E., Clari, G., and Baggio, B. (2003). Arachidonic acid-induced IL-6 expression is mediated by PKC alpha activation in osteoblastic cells. Biochemistry 42, 4485–4491. 3. Traianedes, K., Dallas, M. R., Garrett, I. R., Mundy, G. R., and Bonewald, L. F. (1998). 5-Lipoxygenase metabolites inhibit bone formation in vitro. Endocrinology 139, 3178–3184. 4. Pilbeam, C. C., Raisz, L. G., Voznesensky, O., Alander, C. B., Delman, B. N., and Kawaguchi, H. (1995). Autoregulation of inducible prostaglandin G/H synthase in osteoblastic cells by prostaglandins. J. Bone Miner. Res. 10, 406–414. 5. Miyaura, C., Inada, M., Matsumoto, C., Ohshiba, T., Uozumi, N., Shimizu, T., and Ito, A. (2003). An essential role of cytosolic phospholipase A2alpha in prostaglandin E2-mediated bone resorption associated with inflammation. J. Exp. Med. 197, 1303–1310. 6. Simon, A. M., Manigrasso, M. B., and O’Connor, J. P. (2002). Cyclooxygenase 2 function is essential for bone fracture healing. J. Bone Miner. Res. 17, 963–976. 7. Bakker, A. D., Klein-Nulend, J., and Burger, E. H. (2003). Mechanotransduction in bone cells proceeds via activation of COX-2, but not COX-1. Biochem. Biophys. Res. Commun. 305, 677–683. 8. Zhang, X., Morham, S. G., Langenbach, R., Young, D. A., Xing, L., Boyce, B. F., Puzas, E. J., Rosier, R. N., O’Keefe, R. J., and Schwarz, E. M. (2001). Evidence for a direct role of cyclo-oxygenase 2 in implant wear debris-induced osteolysis. J. Bone Miner. Res. 16, 660–670. 9. Zhang, X., Schwarz, E. M., Young, D. A., Puzas, J. E., Rosier, R. N., and O’Keefe, R. J. (2002). Cyclooxygenase-2 regulates mesenchymal cell differentiation into the osteoblast lineage and is critically involved in bone repair. J. Clin. Invest. 109, 1405–1415. 10. Okada, Y., Lorenzo, J. A., Freeman, A. M., Tomita, M., Morham, S. G., Raisz, L. G., and Pilbeam, C. C. (2000). Prostaglandin G/H synthase-2 is required for maximal formation of osteoclast-like cells in culture. J. Clin. Invest. 105, 823–832. 11. Chikazu, D., Li, X., Kawaguchi, H., Sakuma, Y., Voznesensky, O. S., Adams, D. J., Xu, M., Hoshio, K., Katavic, V., Herschman, H. R., Raisz, L. G., and Pilbeam, C. C. (2002). Bone morphogenetic protein 2 induces cyclo-oxygenase 2 in osteoblasts via a Cbfa1 binding site: role in effects of bone morphogenetic protein 2 in vitro and in vivo. J. Bone Miner. Res. 17, 1430–1440. 12. Choudhary, S., Wadhwa, S., Raisz, L. G., Alander, C., and Pilbeam, C. C. (2003). Extracellular calcium is a potent inducer of cyclo-oxygenase-2 in murine osteoblasts through an ERK signaling pathway. J. Bone Miner. Res. 18, 1813–1824. 13. Krieger, N. S., Parker, W. R., Alexander, K. M., and Bushinsky, D. A. (2000). Prostaglandins regulate acid-induced cell-mediated bone resorption. Am. J. Physiol. Renal Physiol. 279, F1077–F1082. 14. Bushinsky, D. A., Parker, W. R., Alexander, K. M., and Krieger, N. S. (2001). Metabolic, but not respiratory, acidosis increases bone PGE(2). levels and calcium release. Am. J. Physiol. Renal Physiol. 281, 1058–1066. 15. Mano, M., Arakawa, T., Mano, H., Nakagawa, M., Kaneda, T., Kaneko, H., Yamada, T., Miyata, K., Kiyomura, H., Kumegawa, M., and Hakeda, Y. (2000). Prostaglandin E2 directly inhibits bone-resorbing activity of isolated mature osteoclast mainly through the EP4 receptor. Calcif. Tissue Int. 67, 85–92. 16. Romano, C. L., Duci, D., Romano, D., Mazza, M., and Meani, E. (2004). Celecoxib versus indomethacin in the prevention of heterotopic ossification after total hip arthroplasty. J. Arthroplasty 19, 14–18. 17. Saegusa, M., Murakami, M., Nakatani, Y., Yamakawa, K., Katagiri, M., Matsuda, K., Nakamura, K., Kudo, I., and Kawaguchi, H. (2003). Contribution of membrane-associated prostaglandin E2 synthase to bone resorption. J. Cell Physiol. 197, 348–356. 18. Sakuma, Y., Li, Z., Pilbeam, C. C., Alander, C. B., Chikazu, D., Kawaguchi, H., and Raisz, L. G. (2004). Stimulation of cAMP production

References 1. Pilbeam, C. C., Harrison, J. R., and Raisz, L. G. (2002). Prostaglandins and bone metabolism. in (J. P. Bilezikian, L. G. Raisz, G. A. Rodan, eds.) “Principles of Bone Biology”. Academic Press, San Diego, pp. 979–994.

122

19.

20.

21.

22.

23.

24.

25.

26.

27.

28.

29.

30.

31.

32.

33.

and cyclooxygenase-2 by prostaglandin E(2). and selective prostaglandin receptor agonists in murine osteoblastic cells. Bone 34, 827–834. Fermor, B., Jeffcoat, D., Hennerbichler, A., Pisetsky, D. S., Weinberg, J. B., and Guilak, F. (2004). The effects of cyclic mechanical strain and tumor necrosis factor alpha on the response of cells of the meniscus. Osteoarthritis Cartilage 12, 956–962. Huh, Y. H., Kim, S. H., Kim, S. J., and Chun, J. S. (2003). Differentiation status-dependent regulation of cyclooxygenase-2 expression and prostaglandin E2 production by epidermal growth factor via mitogen-activated protein kinase in articular chondrocytes. J. Biol. Chem. 278, 9691–9697. Thomas, B., Thirion, S., Humbert, L., Tan, L., Goldring, M. B., Bereziat, G., and Berenbaum, F. (2002). Differentiation regulates interleukin-1beta-induced cyclo-oxygenase-2 in human articular chondrocytes: role of p38 mitogen-activated protein kinase. Biochem. J. 362, 367–373. LeGrand, A., Fermor, B., Fink, C., Pisetsky, D. S., Weinberg, J. B., Vail, T. P., and Guilak, F. (2001). Interleukin-1, tumor necrosis factor alpha, and interleukin-17 synergistically up-regulate nitric oxide and prostaglandin E2 production in explants of human osteoarthritic knee menisci. Arthritis. Rheum. 44, 2078–2083. Chow, J. W., Fox, S. W., Lean, J. M., and Chambers, T. J. (1998). Role of nitric oxide and prostaglandins in mechanically induced bone formation. J. Bone Miner. Res. 13, 1039–1044. Klein-Nulend, J., Burger, E. H., Semeins, C. M., Raisz, L. G., and Pilbeam, C. C. (1997). Pulsating fluid flow stimulates prostaglandin release and inducible prostaglandin G/H synthase mRNA expression in primary mouse bone cells. J. Bone Miner. Res. 12, 45–51. Forwood, M. R. (1996). Inducible cyclo-oxygenase (COX-2) mediates the induction of bone formation by mechanical loading in vivo. J. Bone Miner. Res. 11, 1688–1693. Li, J., Burr, D. B., and Turner, C. H. (2002). Suppression of prostaglandin synthesis with NS-398 has different effects on endocortical and periosteal bone formation induced by mechanical loading. Calcif. Tissue. Int. 70, 320–329. Wadhwa, S., Godwin, S. L., Peterson, D. R., Epstein, M. A., Raisz, L. G., and Pilbeam, C. C. (2002). Fluid flow induction of cyclo-oxygenase 2 gene expression in osteoblasts is dependent on an extracellular signalregulated kinase signaling pathway. J. Bone Miner. Res. 17, 266–274. Cherian, P. P., Cheng, B., Gu, S., Sprague, E., Bonewald, L. F., and Jiang, J. X. (2003). Effects of mechanical strain on the function of Gap junctions in osteocytes are mediated through the prostaglandin EP2 receptor. J. Biol. Chem. 278, 43146–43156. Jiang, J. X., and Cheng, B. (2001). Mechanical stimulation of gap junctions in bone osteocytes is mediated by prostaglandin E2. Cell Commun. Adhes. 8, 283–288. Saunders, M. M., You, J., Zhou, Z., Li, Z., Yellowley, C. E., Kunze, E. L., Jacobs, C. R., and Donahue, H. J. (2003). Fluid flow-induced prostaglandin E2 response of osteoblastic ROS 17/2.8 cells is gap junction-mediated and independent of cytosolic calcium. Bone 32, 350–356. Kanematsu, M., Sato, T., Takai, H., Watanabe, K., Ikeda, K., and Yamada, Y. (2000). Prostaglandin E2 induces expression of receptor activator of nuclear factor-kappaB ligand/osteoprotegrin ligand on pre-B cells: Implications for accelerated osteoclastogenesis in estrogen deficiency. J. Bone Miner. Res. 15, 1321–1329. Miyaura, C., Inada, M., Suzawa, T., Sugimoto, Y., Ushikubi, F., Ichikawa, A., Narumiya, S., and Suda, T. (2000). Impaired bone resorption to prostaglandin E2 in prostaglandin E receptor EP4-knockout mice. J. Biol. Chem. 275, 19819–19823. Sakuma, Y., Tanaka, K., Suda, M., Komatsu, Y., Yasoda, A., Miura, M., Ozasa, A., Narumiya, S., Sugimoto, Y., Ichikawa, A., Ushikubi, F., and Nakao, K. (2000). Impaired bone resorption by lipopolysaccharide in vivo in mice deficient in the prostaglandin E receptor EP4 subtype. Infect. Immun. 68, 6819–6825.

Lawrence G. Raisz and Joseph A. Lorenzo 34. Sakuma, Y., Tanaka, K., Suda, M., Yasoda, A., Natsui, K., Tanaka, I., Ushikubi, F., Narumiya, S., Segi, E., Sugimoto, Y., Ichikawa, A., and Nakao, K. (2000). Crucial involvement of the EP4 subtype of prostaglandin E receptor in osteoclast formation by proinflammatory cytokines and lipopolysaccharide. J. Bone Miner. Res. 15, 218–227. 35. Tomita, M., Li, X., Okada, Y., Woodiel, F. N., Young, R. N., Pilbeam, C. C., and Raisz, L. G. (2002). Effects of selective prostaglandin EP4 receptor antagonist on osteoclast formation and bone resorption in vitro. Bone 30, 159–163. 36. Li, X. D., Okada, Y., Pilbeam, C. C., Lorenzo, J. A., Kennedy, C. R. J., Breyer, R. M., and Raisz, L. G. (2000). Knockout of the murine prostaglandin EP2 receptor impairs osteoclastogenesis in vitro. Endocrinology 141, 2054–2061. 37. Raisz, L. G., and Woodiel, F. N. (2003). Effects of selective prostaglandin EP2 and EP4 receptor agonists on bone resorption and formation in fetal rat organ cultures. Prostaglandins Other Lipid Mediat. 71, 287–292. 38. Brandstrom, H., Jonsson, K. B., Ohlsson, C., Vidal, O., Ljunghall, S., and Ljunggren, O. (1998). Regulation of osteoprotegerin mRNA levels by prostaglandin E2 in human bone marrow stroma cells. Biochem. Biophys. Res. Commun. 247, 338–341. 39. Suda, K., Udagawa, N., Sato, N., Takami, M., Itoh, K., Woo, J. T., Takahashi, N., and Nagai, K. (2004). Suppression of osteoprotegerin expression by prostaglandin E(2) is crucially involved in lipopolysaccharide-induced osteoclast formation. J. Immunol. 172, 2504–2510. 40. Li, X., Pilbeam, C. C., Pan, L., Breyer, R. M., and Raisz, L. G. (2002). Effects of prostaglandin E(2) on gene expression in primary osteoblastic cells from prostaglandin receptor knockout mice. Bone 30, 567–573. 41. Wani, M. R., Fuller, K., Kim, N. S., Choi, Y. W., and Chambers, T. (1999). Prostaglandin E2 cooperates with TRANCE in osteoclast induction from hemopoietic precursors: Synergistic activation of differentiation, cell spreading, and fusion. Endocrinology 140, 1927–1935. 42. Granchi, D., Amato, I., Battistelli, L., Ciapetti, G., Pagani, S., Avnet, S., Baldini, N., and Giunti, A. (2005). Molecular basis of osteoclastogenesis induced by osteoblasts exposed to wear particles. Biomaterials 26, 2371–2379. 43. Sakuma, Y., Tanaka, K., Suda, M., Yasoda, A., Natsui, K., Tanaka, I., Ushikubi, F., Narumiya, S., Segi, E., Sugimoto, Y., Ichikawa, A., and Nakao, K. (2000). Crucial involvement of the EP4 subtype of prostaglandin E receptor in osteoclast formation by proinflammatory cytokines and lipopolysaccharide. J. Bone Miner. Res. 15, 218–227. 44. Cui, L., Ma, Y. F., Yao, W., Zhou, H., Setterberg, R. B., Liang, T. C., and Jee, W. S. (2001). Cancellous bone of aged rats maintains its capacity to respond vigorously to the anabolic effects of prostaglandin E2 by modeling-dependent bone gain. J. Bone Miner. Metab. 19, 29–37. 45. Zhou, H., Ma, Y. F., Yao, W., Cui, L., Setterberg, R., Liang, C. T., and Jee, W. S. (2001). Lumbar vertebral cancellous bone is capable of responding to PGE2 treatment by stimulating both modeling and remodeling-dependent bone gain in aged male rats. Calcif. Tissue Int. 68, 179–184. 46. Ho, M. L., Chang, J. K., Chuang, L. Y., Hsu, H. K., and Wang, G. J. (1999). Effects of nonsteroidal anti-inflammatory drugs and prostaglandins on osteoblastic functions. Biochem. Pharmacol. 58, 983–990. 47. Centrella, M., Casinghino, S., and McCarthy, T. L. (1994). Differential actions of prostaglandins in separate cell populations from fetal rat bone. Endocrinology 135, 1611–1620. 48. Keila, S., Kelner, A., and Weinreb, M. (2001). Systemic prostaglandin E2 increases cancellous bone formation and mass in aging rats and stimulates their bone marrow osteogenic capacity in vivo and in vitro. J. Endocrinol. 168, 131–139. 49. Kaneki, H., Takasugi, I., Fujieda, M., Kiriu, M., Mizuochi, S., and Ide, H. (1999). Prostaglandin E2 stimulates the formation of mineralized bone

Chapter 7 Prostaglandins and Proinflammatory Cytokines

123

nodules by a cAMP-independent mechanism in the culture of adult rat calvarial osteoblasts. J. Cell. Biochem. 73, 36–48. Conconi, M. T., Tommasini, M., Baiguera, S., De Coppi, P., Parnigotto, P. P., and Nussdorfer, G. G. (2002). Effects of prostaglandins E1 and E2 on the growth and differentiation of osteoblast-like cells cultured in vitro. Int. J. Mol. Med. 10, 451–456. Fall, P. M., Breault, D. T., and Raisz, L. G. (1994). Inhibition of collagen synthesis by prostaglandins in the immortalized rat osteoblastic cell line Py1a: Structure-activity relations and signal transduction mechanisms. J. Bone Miner. Res. 9; 1935–1943. Li, M., Ke, H. Z., Qi, H., Healy, D. R., Li, Y., Crawford, D. T., Paralkar, V. M., Owen, T. A., Cameron, K. O., Lefker, B. A., Brown, T. A., and Thompson, D. D. (2003). A novel, non-prostanoid EP2 receptorselective prostaglandin E2 agonist stimulates local bone formation and enhances fracture healing. J. Bone Miner. Res. 18, 2033–2042. Paralkar, V. M., Borovecki, F., Ke, H. Z., Cameron, K. O., Lefker, B., Grasser, W. A., Owen, T. A., Li, M., DaSilva-Jardine, P., Zhou, M., Dunn, R. L., Dumont, F., Korsmeyer, R., Krasney, P., Brown, T. A., Plowchalk, D., Vukicevic, S., and Thompson, D. D. (2003). An EP2 receptor-selective prostaglandin E2 agonist induces bone healing. Proc. Natl Acad. Sci. USA 100, 6736–6740. Machwate, M., Harada, S., Leu, C. T., Seedor, G., Labelle, M., Gallant, M., Hutchins, S., Lachance, N., Sawyer, N., Slipetz, D., Metters, K. M., Rodan, S. B., Young, R., and Rodan, G. A. (2001). Prostaglandin receptor EP(4) mediates the bone anabolic effects of PGE(2). Mol. Pharmacol. 60, 36–41. Shamir, D., Keila, S., and Weinreb, M. (2004). A selective EP4 receptor antagonist abrogates the stimulation of osteoblast recruitment from bone marrow stromal cells by prostaglandin E2 in vivo and in vitro. Bone 34, 157–162. Yoshida, K., Oida, H., Kobayashi, T., Maruyama, T., Tanaka, M., Katayama, T., Yamaguchi, K., Segi, E., Tsuboyama, T., Matsushita, M., Ito, K., Ito, Y., Sugimoto, Y., Ushikubi, F., Ohuchida, S., Kondo, K., Nakamura, T., and Narumiya, S. (2002). Stimulation of bone formation and prevention of bone loss by prostaglandin E EP4 receptor activation. Proc. Natl Acad. Sci. USA 99, 4580–4585. Soper, D. L., Wang, Y., De, B., deLong, M. A., Dirr, M. J., Soehner, M. E., Lundy, M. W., Mieling, G. E., and Wos, J. A. (2002). The design and synthesis of selective prostaglandin analogs as bone anabolic agents for the potential treatment of osteoporosis. Adv. Exp. Med. Biol. 507, 303–307. Raisz, L. G., Fall, P. M., Gabbitas, B. Y., McCarthy, T. L., Kream, B. E., and Canalis, E. (1993). Effects of prostaglandin E2 on bone formation in cultured fetal rat calvariae: Role of insulin-like growth factor- I. Endocrinology 133, 1504–1510. Harada, S., Nagy, J. A., Sullivan, K. A., Thomas, K. A., Endo, N., Rodan, G. A., and Rodan, S. B. (1994). Induction of vascular endothelial growth factor expression by prostaglandin E2 and E1 in osteoblasts. J. Clin. Invest. 93, 2490–2496. Arikawa, T., Omura, K., and Morita, I. (2004). Regulation of bone morphogenetic protein-2 expression by endogenous prostaglandin E2 in human mesenchymal stem cells. J. Cell. Physiol. 200, 400–406. Paralkar, V. M., Grasser, W. A., Mansolf, A. L., Baumann, A. P., Owen, T. A., Smock, S. L., Martinovic, S., Borovecki, F., Vukicevic, S., Ke, H. Z., and Thompson, D. D. (2002). Regulation of BMP-7 expression by retinoic acid and prostaglandin E(2). J. Cell. Physiol. 190, 207–217. Sabbieti, M. G., Marchetti, L., Abreu, C., Montero, A., Hand, A. R., Raisz, L. G., and Hurley, M. M. (1999). Prostaglandins regulate the expression of fibroblast growth factor-2 in bone. Endocrinology 140, 434–444. Chikazu, D., Katagiri, M., Ogasawara, T., Ogata, N., Shimoaka, T., Takato, T., Nakamura, K., and Kawaguchi, H. (2001). Regulation of osteoclast differentiation by fibroblast growth factor 2: stimulation of receptor activator of nuclear factor kappaB ligand/osteoclast

differentiation factor expression in osteoblasts and inhibition of macrophage colony-stimulating factor function in osteoclast precursors. J. Bone Miner. Res. 16, 2074–2081. Samoto, H., Shimizu, E., Matsuda-Honjyo. Y., Saito, R., Nakao, S., Yamazaki, M., Furuyama, S., Sugiya, H., Sodek, J., and Ogata, Y. (2003). Prostaglandin E2 stimulates bone sialoprotein (BSP) expression through cAMP and fibroblast growth factor 2 response elements in the proximal promoter of the rat BSP gene. J. Biol. Chem. 278, 28659–28667. Goldring, M. B., Berenbaum, F., and Buckwalter, J. (2004). The regulation of chondrocyte function by proinflammatory mediators: prostaglandins and nitric oxide. Clin. Orthop. 427, (Suppl) S37–S46. Laufer, S. (2003). Role of eicosanoids in structural degradation in osteoarthritis. Curr. Opin. Rheumatol. 15, 623–627. Notoya, K., Jovanovic, D. V., Reboul, P., Martel-Pelletier, J., Mineau, F., and Pelletier, J. P. (2000). The induction of cell death in human osteoarthritis chondrocytes by nitric oxide is related to the production of prostaglandin E2 via the induction of cyclooxygenase-2. J. Immunol. 165, 3402–3410. Geng, Y., Blanco, F. J., Cornelisson, M., and Lotz, M. (1995). Regulation of cyclooxygenase-2 expression in normal human articular chondrocytes. J. Immunol. 155, 796–801. Trebino, C. E., Stock, J. L., Gibbons, C. P., Naiman, B. M., Wachtmann, T. S., Umland, J. P., Pandher, K., Lapointe, J. M., Saha, S., Roach, M. L., Carter, D., Thomas, N. A., Durtschi, B. A., McNeish, J. D., Hambor, J. E., Jakobsson, P. J., Carty, T. J., Perez, J. R., and Audoly, L. P. (2003). Impaired inflammatory and pain responses in mice lacking an inducible prostaglandin E synthase. Proc. Natl Acad. Sci. USA 100, 9044–9049. McCoy, J. M., Wicks, J. R., and Audoly, L. P. (2002). The role of prostaglandin E2 receptors in the pathogenesis of rheumatoid arthritis. J. Clin. Invest. 110, 651–658. Li, T. F., Zuscik, M. J., Ionescu, A. M., Zhang, X., Rosier, R. N., Schwarz, E. M., Drissi, H., and O’Keefe, R. J. (2004). PGE2 inhibits chondrocyte differentiation through PKA and PKC signaling. Exp. Cell. Res. 300, 159–169. Boileau, C., Martel-Pelletier, J., Jouzeau, J. Y., Netter, P., Moldovan, F., Laufer, S., Tries, S., and Pelletier, J. P. (2002). Licofelone (ML-3000), a dual inhibitor of 5-lipoxygenase and cyclooxygenase, reduces the level of cartilage chondrocyte death in vivo in experimental dog osteoarthritis: inhibition of pro-apoptotic factors. J. Rheumatol. 29, 1446–1453. DiBattista, J. A., Dore, S., Morin, N., and Abribat, T. (1996). Prostaglandin E(2) up-regulates insulin-like growth factor binding protein-3 expression and synthesis in human articular chondrocytes by a c-AMP-independent pathway: Role of calcium and protein kinase A and C. J. Cell. Biochem. 63, 320–333. Hardy, M. M., Seibert, K., Manning, P. T., Currie, M. G., Woerner, B. M., Edwards, D., Koki, A., and Tripp, C. S. (2002). Cyclooxygenase 2-dependent prostaglandin E2 modulates cartilage proteoglycan degradation in human osteoarthritis explants. Arthritis. Rheum. 46, 1789–1803. Miyamoto, M., Ito, H., Mukai, S., Kobayashi, T., Yamamoto, H., Kobayashi, M., Maruyama, T., Akiyama, H., and Nakamura, T. (2003). Simultaneous stimulation of EP2 and EP4 is essential to the effect of prostaglandin E2 in chondrocyte differentiation. Osteoarthritis Cartilage 11, 644–652. Abramson, S. B. (1999). The role of COX-2 produced by cartilage in arthritis. Osteoarthritis Cartilage 7, 380–381. Raisz, L. G. (1999). Prostaglandins and bone: physiology and pathophysiology. Osteoarthritis Cartilage 7, 419–421. Raisz, L. G. (2001). Potential impact of selective cyclooxygenase-2 inhibitors on bone metabolism in health and disease. Am. J. Med. 110, 43–45. Crevenna, R., Quittan, M., Hulsmann, M., Wiesinger, G. F., Keilani, M. Y., Kainberger, F., Leitha, T., Fialka-Moser, V., and Pacher, R. (2002).

50.

51.

52.

53.

54.

55.

56.

57.

58.

59.

60.

61.

62.

63.

64.

65.

66. 67.

68.

69.

70.

71.

72.

73.

74.

75.

76. 77. 78.

79.

124

80.

81.

82.

83.

84. 85.

86.

87.

88.

89. 90.

91.

92.

93.

Hypertrophic osteoarthropathy caused by PGE1 in a patient with congestive heart failure during cardiac rehabilitation. Wien. Klin. Wochenschr. 114, 115–118. Thorsen, K., Kristoffersson, A. O., Lerner, U. H., and Lorentzon, R. P. (1996). In situ microdialysis in bone tissue – Stimulation of prostaglandin E2 release by weight-bearing mechanical loading. J. Clin. Invest. 98, 2446–2449. Morton, D. J., Barrett-Connor, E. L., and Schneider, D. L. (1998). Nonsteroidal anti-inflammatory drugs and bone mineral density in older women: the Rancho Bernardo study. J. Bone Miner. Res. 13, 1924–1931. Carbone, L. D., Tylavsky, F. A., Cauley, J. A., Harris, T. B., Lang, T. F., Bauer, D. C., Barrow, K. D., and Kritchevsky, S. B. (2003). Association between bone mineral density and the use of nonsteroidal antiinflammatory drugs and aspirin: impact of cyclooxygenase selectivity. J. Bone Miner. Res. 18, 1795–1802. Head, J. E., Bryant, B. J., Grills, B. L., and Ebeling, P. R. (2001). Effects of short-term use of ibuprofen or acetaminophen on bone resorption in healthy men: a double-blind, placebo-controlled pilot study. Bone 29, 437–441. Teitelbaum, S. L. (2000). Bone resorption by osteoclast. Science 289, 1504–1508. Lacey, D. L., Timms, E., Tan, H. L., Kelley, M. J., Dunstan, C. R., Burgess, T., Elliott, R., Colombero, A., Elliott, G., Scully, S., Hsu, H., Sullivan, J., Hawkins, N., Davy, E., Capparelli, C., Eli, A., Qian, Y. X., Kaufman, S., Sarosi, I., Shalhoub, V., Senaldi, G., Guo, J., Delaney, J., and Boyle, W. J. (1998). Osteoprotegerin ligand is a cytokine that regulates osteoclast differentiation and activation. Cell 93, 165–176. Yasuda, H., Shima, N., Nakagawa, N., Yamaguchi, K., Kinosaki, M., Mochizuki, S., Tomoyasu, A., Yano, K., Goto, M., Murakami, A., Tsuda, E., Morinaga, T., Higashio, K., Udagawa, N., Takahashi, N., and Suda, T. (1998). Osteoclast differentiation factor is a ligand for osteoprotegerin osteoclastogenesis-inhibitory factor and is identical to TRANCE/RANKL. Proc. Natl Acad. Sci. USA 95, 3597–3602. Anderson, D. M., Maraskovsky, E., Billingsley, W. L., Dougall, W. C., Tometsko, M. E., Roux, E. R., Teepe, M. C., DuBose, R. F., Cosman, D., and Galibert, L. (1997). A homologue of the TNF receptor and its ligand enhance T-cell growth and dendritic-cell function. Nature 390, 175–179. Wong, B. R., Rho, J., Arron, J., Robinson, E., Orlinick, J., Chao, M., Kalachikov, S., Cayani, E., Bartlett, F. S., III, Frankel, W. N., Lee, S. Y., and Choi, Y. (1997). TRANCE is a novel ligand of the tumor necrosis factor receptor family that activates c-Jun N-terminal kinase in T cells. J. Biol. Chem. 272, 25190–25194. Boyle, W. J., Simonet, W. S., and Lacey, D. L. (2003). Osteoclast differentiation and activation. Nature 423, 337–342. Simonet, W. S., Lacey, D. L., Dunstan, C. R., Kelley, M., Chang, M. S., Luthy, R., Nguyen, H. Q., Wooden, S., Bennett, L., Boone, T., Shimamoto, G., DeRose, M., Elliott, R., Colombero, A., Tan, H. L., Trail, G., Sullivan, J., Davy, E., Bucay, N., Renshaw-Gegg, L., Hughes, T. M., Hill, D., Pattison, W., Campbell, P., and Boyle, W. J. (1997). Osteoprotegerin: a novel secreted protein involved in the regulation of bone density. Cell 89, 309–319. Lee, S. K., and Lorenzo, J. A. (1999). Parathyroid hormone stimulates TRANCE and inhibits osteoprotegerin messenger ribonucleic acid expression in murine bone marrow cultures: correlation with osteoclastlike cell formation. Endocrinology 140, 3552–3561. Lee, S. K., Kalinowski, J., Jastrzebski, S., and Lorenzo, J. A. (2002). 1,25 (OH)(2) Vitamin D(3)-stimulated osteoclast formation in spleenosteoblast cocultures is mediated in part by enhanced IL-1alpha and receptor activator of NF-kappaB ligand production in osteoblasts. J. Immunol. 169, 2374–2380. Hofbauer, L. C., Lacey, D. L., Dunstan, C. R., Spelsberg, T. C., Riggs, B. L., and Khosla, S. (1999). Interleukin-1beta and tumor necrosis factor-alpha, but not interleukin-6, stimulate osteoprotegerin ligand gene expression in human osteoblastic cells. Bone 25, 255–259.

Lawrence G. Raisz and Joseph A. Lorenzo 94. Vidal, O. N., Sjögren, K., Eriksson, B. I., Ljunggren, Ö., and Ohlsson, C. (1998). Osteoprotegerin mRNA is increased by interleukin-1α in the human osteosarcoma cell line MG-63 and in human osteoblast-like cells. Biochem. Biophys. Res. Com. 248, 696–700. 95. Pixley, F. J., and Stanley, E. R. (2004). CSF-1 regulation of the wandering macrophage: complexity in action. Trends Cell. Biol. 14, 628–638. 96. Yoshida, H., Hayashi, S., Kunisada, T., Ogawa, M., Nishikawa, S., Okamura, H., Sudo, T., Shultz, L. D., and Nisikawa, S. (1990). The murine mutation osteopetrosis is in the coding region of the macrophage colony stimulating factor gene. Nature 345, 442–444. 97. Wiktor Jedrzejczak, W., Bartocci, A., Ferrante, A., Jr, Ahmed Ansari, A., Sell, K. W., Pollard, J. W., and Stanley, E. R. (1990). Total absence of colony-stimulating factor 1 in the macrophage-deficient osteopetrotic (op/op) mouse. Proc. Natl Acad. Sci. USA 87, 4828–4832. 98. Arai, F., Miyamoto, T., Ohneda, O., Inada, T., Sudo, T., Brasel, K., Miyata, T., Anderson, D. M., and Suda, T. (1999). Commitment and differentiation of osteoclast precursor cells by the sequential expression of c-Fms and receptor activator of nuclear factor kappaB (RANK) receptors. J. Exp. Med. 190, 1741–1754. 99. Lacey, D. L., Tan, H. L., Lu, J., Kaufman, S., Van, G., Qiu, W., Rattan, A., Scully, S., Fletcher, F., Juan, T., Kelley, M., Burgess, T. L., Boyle, W. J., and Polverino, A. J. (2000). Osteoprotegerin ligand modulates murine osteoclast survival in vitro and in vivo. Am. J. Pathol. 157, 435–448. 100. Dai, X. M., Ryan, G. R., Hapel, A. J., Dominguez, M. G., Russell, R. G., Kapp, S., Sylvestre, V., and Stanley, E. R. (2002). Targeted disruption of the mouse colony-stimulating factor 1 receptor gene results in osteopetrosis, mononuclear phagocyte deficiency, increased primitive progenitor cell frequencies, and reproductive defects. Blood 99, 111–120. 101. Cella, M., Buonsanti, C., Strader, C., Kondo, T., Salmaggi, A., and Colonna, M. (2003). impaired differentiation of osteoclast in TREM-2-deficient individuals. J. Exp. Med. 198, 645–651. 102. Kim, N., Takami, M., Rho, J., Josien, R., and Choi, Y. (2002). A novel member of the leukocyte receptor complex regulates osteoclast differentiation. J. Exp. Med. 195, 201–209. 103. Colonna, M. (2003). DAP12 signaling: from immune cells to bone modeling and brain myelination. J. Clin. Invest. 111, 313–314. 104. Kaifu, T., Nakahara, J., Inui, M., Mishima, K., Momiyama, T., Kaji, M., Sugahara, A., Koito, H., Ujike-Asai, A., Nakamura, A., Kanazawa, K., Tan-Takeuchi, K., Iwasaki, K., Yokoyama, W. M., Kudo, A., Fujiwara, M., Asou, H., and Takai, T. (2003). Osteopetrosis and thalamic hypomyelinosis with synaptic degeneration in DAP12-deficient mice. J. Clin. Invest. 111, 323–332. 105. Koga, T., Inui, M., Inoue, K., Kim, S., Suematsu, A., Kobayashi, E., Iwata, T., Ohnishi, H., Matozaki, T., Kodama, T., Taniguchi, T., Takayanagi, H., and Takai, T. (2004). Costimulatory signals mediated by the ITAM motif cooperate with RANKL for bone homeostasis. Nature 428, 758–763. 106. Humphrey, M. B., Ogasawara, K., Yao, W., Spusta, S. C., Daws, M. R., Lane, N. E., Lanier, L. L., and Nakamura, M. C. (2004). The signaling adapter protein DAP12 regulates multinucleation during osteoclast development. J. Bone Miner. Res. 19, 224–234. 107. Mocsai, A., Humphrey, M. B., Van Ziffle, J. A., Hu, Y., Burghardt, A., Spusta, S. C., Majumdar, S., Lanier, L. L., Lowell, C. A., and Nakamura, M. C. (2004). The immunomodulatory adapter proteins DAP12 and Fc receptor gamma-chain (FcRgamma) regulate development of functional osteoclast through the Syk tyrosine kinase. Proc. Natl Acad. Sci. USA 101, 6158–6163. 108. Dinarello, C. A. (1991). Interleukin-1 and interleukin-1 antagonism. Blood 77, 1627–1652. 109. Dewhirst, F. E., Stashenko, P. P., Mole, J. E., and Tsurumachi, T. (1985). Purification and partial sequence of human osteoclastactivating factor: Identity with interleukin 1 beta. J. Immunol. 135, 2562–2568.

Chapter 7 Prostaglandins and Proinflammatory Cytokines

125

110. Lorenzo, J. A., Sousa, S., Alander, C., Raisz, L. G., and Dinarello, C. A. (1987). Comparison of the bone-resorbing activity in the supernatants from phytohemagglutinin-stimulated human peripheral blood mononuclear cells with that of cytokines through the use of an antiserum to interleukin 1. Endocrinology 121, 1164–1170. 111. Min, Y. K., Rao, Y., Okada, Y., Raisz, L. G., and Pilbeam, C. C. (1998). Regulation of prostaglandin G/H synthase-2 expression by interleukin-1 in human osteoblast-like cells. J. Bone Miner. Res. 13, 1066–1075. 112. Hughes, F. J., Buttery, L. D. K., Hukkanen, M. V. J., O’Donnell, A., Maclouf, J., and Polak, J. M. (1999). Cytokine-induced prostaglandin E2 synthesis and cyclooxygenase-2 activity are regulated both by a nitric oxide-dependent and -independent mechanism in rat osteoblasts in vitro. J. Biol. Chem. 274, 1776–1782. 113. Boyce, B. F., Aufdemorte, T. B., Garrett, R., Yates, A. J. P., and Mundy, G. R. (1989). Effects of interleukin-1 on bone turnover in normal mice. Endocrinology 125, 1142. 114. Smith, D. D., Gowen, M., and Mundy, G. R. (1987). Effects of interferon-gamma and other cytokines on collagen synthesis in fetal rat bone cultures. Endocrinology 120, 2494–2499. 115. Canalis, E. (1986). Interleukin-1 has independent effects on deoxyribonucleic acid and collagen synthesis in cultures of rat calvariae. Endocrinology 118, 74–81. 116. Stashenko, P., Dewhirst, F. E., Rooney, M. L., Desjardins, L. A., and Heeley, J. D. (1987). Interleukin-1 Beta is a potent inhibitor of bone formation in vitro. J. Bone Miner. Res. 2, 559–565. 117. Gowen, M., Wood, D. D., and Russell, G. G. (1985). Stimulation of the proliferation of human bone cells in vitro by human monocyte products with interleukin-1 activity. J. Clin. Invest. 75, 1223–1229. 118. Lorenzo, J. A., Sousa, S. L., Van Den Brink-Webb, S. E., and Korn, J. H. (1990). Production of both interleukin-1 α and β by newborn mouse calvaria cultures. J. Bone Miner. Res. 5, 77–83. 119. Kawaguchi, H., Pilbeam, C. C., Vargas, S. J., Morse, E. E., Lorenzo, J. A., and Raisz, L. G. (1995). Ovariectomy enhances and estrogen replacement inhibits the activity of bone marrow factors that stimulate prostaglandin production in cultured mouse calvaria. J. Clin. Invest. 96, 539–548. 120. Miyaura, C., Kusano, K., Masuzawa, T., Chaki, O., Onoe, Y., Aoyagi, M., Sasaki, T., Tamura, T., Koishihara, Y., Ohsugi, Y., and Suda, T. (1995). Endogenous bone-resorbing factors in estrogen deficiency: Cooperative effects of IL-1 and IL-6. J. Bone Miner. Res. 10, 1365–1373. 121. Horowitz, M. C., Philbrick, W. M., and Jilka, R. L. (1989). IL-1 release from cultured calvarial cells is due to macrophages. J. Bone Miner. Res. (abstr)4, (S1)556. 122. Keeting, P. E., Rifas, L., Harris, S. A., Colvard, D. S., Spelsberg, T. C., Peck, W. A., and Riggs, B. L. (1991). Evidence for interleukin-1 beta production by cultured normal human osteoblast-like cells. J. Bone Miner. Res. 6, 827–833. 123. O’Keefe, R. J., Teot, L. A., Singh, D., Puzas, J. E., Rosier, R. N., and Hicks, D. G. (1997). Osteoclast constitutively express regulators of bone resorption: an immunohistochemical and in situ hybridization study. Lab. Invest. 76, 457–465. 124. Kobayashi, K., Takahashi, N., Jimi, E., Udagawa, N., Takami, M., Kotake, S., Nakagawa, N., Kinosaki, M., Yamaguchi, K., Shima, N., Yasuda, H., Morinaga, T., Higashio, K., Martin, T. J., and Suda, T. (2000). Tumor necrosis factor alpha stimulates osteoclast differentiation by a mechanism independent of the ODF/RANKL-RANK interaction. J. Exp. Med. 191, 275–286. 125. Eisenberg, S. P., Evans, R. J., Arend, W. P., Verderber, E., Brewer, M. T., Hannum, C. H., and Thompson, R. C. (1990). Primary structure and functional expression from complementary DNA of a human interlukin-1 receptor antagonist. Nature 343, 341–346. 126. Hannum, C. H., Wilcox, C. J., Arend, W. P., Joslin, F. G., Dripps, D. J., Heimdal, P. L., Armes, L. G., Sommer, A., Eisenberg, S. P., and

Thompson, R. C. (1990). Interleukin-1 receptor antagonist activity of a human interleukin-1 inhibitor. Nature 343, 336–340. Arend, W. P., Welgus, H. G., Thompson, C., and Eisenberg, S. P. (1990). Biological properties of recombinant human monocytederived interleukin 1 receptor antagonist. J. Clin. Invest. 85, 1694–1697. Seckinger, P., Klein-Nulend, J., Alander, C., Thompson, R. C., Dayer, J-M., and Raisz, L. G. (1990). Natural and recombinant human IL-1 receptor antagonists block the effects of IL-1 on bone resorption and prostaglandin production. J. Immunol. 145, 4181–4184. Pacifici, R., Vannice, J. L., Rifas, L., and Kimble, R. B. (1993). Monocytic secretion of interleukin-1 receptor antagonist in normal and osteoporotic women: Effects of menopause and estrogen/progesterone therapy. J. Clin. Endocrinol. Metab. 77, 1135–1141. Abrahamsen, B., Shalhoub, V., Larson, E., Eriksen, E. F., Beck-Nielsen, H., and Marks, S. C., Jr. (1996). IL-1 beta and IL-1RA gene expression in transiliac bone biopsies: correlation with bone loss and histomorphometry in healthy postmenopausal women. J. Bone Miner. Res. 11, S42 (Abstract). Dinarello, C. A. (1993). Blocking interleukin-1 in disease. Blood Purif. 11, 118–127. Sims, J. E., Gayle, M. A., Slack, J. L., Alderson, M. R., Bird, T. A., Giri, J. G., Colotta, F., Re, F., Mantovani, A., Shanebeck, K., Grabstein, K. H., and Dower, S. K. (1993). Interleukin 1 signaling occurs exclusively via the type I receptor. Proc. Natl Acad. Sci. USA 90, 6155–6159. Eder, J. (1997). Tumour necrosis factor alpha and interleukin 1 signalling: do MAPKK kinases connect it all? Trends Pharmacol. Sci. 18, 319–322. Martin, M. U., and Falk, W. (1997). The interleukin-1 receptor complex and interleukin-1 signal transduction. Eur. Cytokine Netw. 8, 5–17. Korherr, C., Hofmeister, R., Wesche, H., and Falk, W. (1997). A critical role for interleukin-1 receptor accessory protein in interleukin-1 signaling. Eur. J. Immunol. 27, 262–267. Wesche, H., Korherr, C., Kracht, M., Falk, W., Resch, K., and Martin, M. U. (1997). The interleukin-1 receptor accessory protein (IL-1RAcP) is essential for IL-1-induced activation of interleukin-1 receptor-associated kinase (IRAK) and stress-activated protein kinases (SAP kinases). J. Biol. Chem. 272, 7727–7731. Huang, J., Gao, X., Li, S., and Cao, Z. (1997). Recruitment of IRAK to the interleukin 1 receptor complex requires interleukin 1 receptor accessory protein. Proc. Natl Acad. Sci. USA 94, 12829–12832. Colotta, F., Re, F., Muzio, M., Bertini, R., Polentarutti, N., Sironi, M., Giri, J. G., Dower, S. K., Sims, J. E., and Mantovani, A. (1993). Interleukin-1 type II receptor: A decoy target for IL-1 that is regulated by IL-4. Science 261, 472–475. Burger, D., Chicheportiche, R., Giri, J. G., and Dayer, J-M. (1995). The inhibitory activity of human interleukin-1 receptor antagonist is enhanced by type II interleukin-1 soluble receptor and hindered by type 1 interleukin-1 soluble receptor. J. Clin. Invest. 96, 38–41. Kitazawa, R., Kimble, R. B., Vannice, J. L., Kung, V. T., and Pacifici, R. (1994). Interleukin-1 receptor antagonist and tumor necrosis factor binding protein decrease osteoclast formation and bone resorption in ovariectomized mice. J. Clin. Invest. 94, 2397–2406. Lorenzo, J. A., Naprta, A., Rao, Y., Alander, C., Glaccum, M., Widmer, M., Gronowicz, G., Kalinowski, J., and Pilbeam, C. C. (1998). Mice lacking the type I interleukin-1 receptor do not lose bone mass after ovariectomy. Endocrinology 139, 3022–3025. Fiore, C. E., Falcidia, E., Foti, R., Motta, M., and Tamburino, C. (1994). Differences in the time course of the effects of oophorectomy in women on parameters of bone metabolism and interleukin-1 levels in the circulation. Bone Miner. 20, 79–85. McKane, W. R., Khosla, S., Peterson, J. M., Egan, K., and Riggs, B. L. (1994). Circulating levels of cytokines that modulate

127.

128.

129.

130.

131. 132.

133.

134.

135.

136.

137.

138.

139.

140.

141.

142.

143.

126

144.

145.

146.

147.

148.

149.

150.

151.

152.

153.

154.

155.

156. 157. 158. 159. 160. 161.

162.

Lawrence G. Raisz and Joseph A. Lorenzo bone resorption: Effects of age and menopause in women. J. Bone Miner. Res. 9, 1313–1318. Khosla, S., Peterson, J. M., Egan, K., Jones, J. D., and Riggs, B. L. (1994). Circulating cytokine levels in osteoporotic and normal women. J. Clin. Endocrinol. Metab. 79, 707–711. Pilbeam, C., Rao, Y., Alander, C., Voznesensky, O., Okada, Y., Sims, J. E., Raisz, L., and Lorenzo, J. (1997). Downregulation of mRNA expression for the “Decoy” interleukin-1 receptor 2 by ovariectomy in mice. J. Bone Miner. Res. 12, S433 (Abstract). Sunyer, T., Lewis, J., Collin-Osdoby, P., and Osdoby, P. (1999). Estrogen’s bone-protective effects may involve differential IL-1 receptor regulation in human osteoclast-like cells. J. Clin. Invest. 103, 1409–1418. Iwamoto, M., Koike, T., Nakashima, K., Sato, K., and Kato, Y. (1989). Interleukin 1: a regulator of chondrocyte proliferation. Immunol. Lett. 21, 153–156. Morales, T. I., and Hascall, V. C. (1989). Effects of interleukin-1 and lipopolysaccharides on protein and carbohydrate metabolism in bovine articular cartilage organ cultures. Connect. Tissue Res. 19, 255–275. Grumbles, R. M., Shao, L., Jeffrey, J. J., and Howell, D. S. (1996). Regulation of rat interstitial collagenase gene expression in growth cartilage and chondrocytes by vitamin D3, interleukin-1 beta, and okadaic acid. J. Cell. Biochem. 63, 395–409. Goldring, M. B., Birkhead, J., Sandell, L. J., Kimura, T., and Krane, S. M. (1988). Interleukin 1 suppresses expression of cartilagespecific types II and IX collagens and increases types I and III collagens in human chondrocytes. J. Clin. Invest. 82, 2026–2037. Kato, Y., Nakashima, K., Iwamoto, M., Murakami, H., Hiranuma, H., Koike, T., Suzuki, F., Fuchihata, H., Ikehara, Y., and Noshiro, M. (1993). Effects of interleukin-1 on syntheses of alkaline phosphatase, type X collagen, and 1,25-dihydroxyvitamin D3 receptor, and matrix calcification in rabbit chondrocyte cultures. J. Clin. Invest. 92, 2323–2330. Horan, J., Dean, D. D., Kieswetter, K., Schwartz, Z., and Boyan, B. D. (1996). Evidence that interleukin-1, but not interleukin-6, affects costochondral chondrocyte proliferation, differentiation, and matrix synthesis through an autocrine pathway. J. Bone Miner. Res. 11, 1119–1129. Dean, D. D., Schwartz, Z., Muniz, O. E., Arsenis, C. H., Boyan, B. D., and Howell, D. S. (1997). Interleukin-1alpha and beta in growth plate cartilage are regulated by vitamin D metabolites in vivo. J. Bone Miner. Res. 12, 1560–1569. Vargas, S. J., Naprta, A., Glaccum, M., Lee, S. K., Kalinowski, J., and Lorenzo, J. A. (1996). Interleukin-6 expression and histomorphometry of bones from mice deficient for receptors for interleukin-1 or tumor necrosis factor. J. Bone Miner. Res. 11, 1736–1744. Martensson, K., Chrysis, D., and Savendahl, L. (2004). Interleukin1beta and TNF-alpha act in synergy to inhibit longitudinal growth in fetal rat metatarsal bones. J. Bone Miner. Res. 19, 1805–1812. Beutler, B., and Cerami, A. (1986). Cachectin and tumour necrosis factor as two sides of the same biological coin. Nature 320, 584–588. Oliff, A. (1988). The role of tumor necrosis factor (cachectin) in cachexia. Cell 54, 141–142. Beutler, B., and Cerami, A. (1987). Cachectin: More than a tumor necrosis factor. N. Eng. J. Med. 316, 379–385. Old, L. J. (1985). Tumor necrosis factor (TNF). Science 230, 630–632. Paul, N. L., and Ruddle, N. H. (1988). Lymphotoxin. Ann. Rev. Immunol. 6, 407–438. Bertolini, D. R., Nedwin, G. E., Bringman, T. S., Smith, D. D., and Mundy, G. R. (1986). Stimulation of bone resorption and inhibition of bone formation in vitro by human tumour necrosis factors. Nature 319, 516–518. Tashjian, A. H., Jr., Voelkel, E. F., Lazzaro, M., Goad, D., Bosma, T., and Levine, L. (1987). Tumor necrosis factor-alpha (Cachectin)

163.

164.

165.

166.

167.

168.

169.

170.

171.

172.

173.

174.

175.

176.

177.

stimulates bone resorption in mouse calvaria via a prostaglandinmediated mechanism. Endocrinology 120, 2029–2036. Pfeilschifter, J., Chenu, C., Bird, A., Mundy, G. R., and Roodman, G. D. (1989). Interleukin-1 and tumor necrosis factor stimulate the formation of human osteoclastlike cells in vitro. J. Bone Miner. Res. 4, 113–118. Gowen, M., Chapman, K., Littlewood, A., Hughes, D., Evans, D., and Russell, R. G. G. (1990). Production of tumor necrosis factor by human osteoblasts is modulated by other cytokines, but not by osteotropic hormones. Endocrinology 126, 1250–1255. Azuma, Y., Kaji, K., Katogi, R., Takeshita, S., and Kudo, A. (2000). Tumor necrosis factor-alpha induces differentiation of and bone resorption by osteoclast. J. Biol. Chem. 275, 4858–4864. Li, J., Sarosi, I., Yan, X. Q., Morony, S., Capparelli, C., Tan, H. L., McCabe, S., Elliott, R., Scully, S., Van, G., Kaufman, S., Juan, S. C., Sun, Y., Tarpley, J., Martin, L., Christensen, K., McCabe, J., Kostenuik, P., Hsu, H., Fletcher, F., Dunstan, C. R., Lacey, D. L., and Boyle, W. J. (2000). RANK is the intrinsic hematopoietic cell surface receptor that controls osteoclastogenesis and regulation of bone mass and calcium metabolism. Proc. Natl Acad. Sci. USA 97, 1566–1571. Fiers, W. (1993). Tumor necrosis factor. In (Sim, E., ed.), “The Natural Immune System: Humoral Factors.” IRL Press at Oxford University Press, Oxford, pp. 65–119. Rothe, J., Lesslauer, W., Lotscher, H., Lang, Y., Koebel, P., Kontgen, F., Althage, A., Zinkernagel, R., Steinmetz, M., and Bluethmann, H. (1993). Mice lacking the tumour necrosis factor receptor 1 are resistant to TNF-mediated toxicity but highly susceptible to infection by Listeria monocytogenes. Nature 364, 798–802. Pfeffer, K., Matsuyama, T., Kündig, T. M., Wakeham, A., Kishihara, K., Shahinian, A., Wiegmann, K., Ohashi, P. S., Krönke, M., and Mak, T. W. (1993). Mice deficient for the 55 kd tumor necrosis factor receptor are resistant to endotoxic shock, yet succumb to L. monocytogenes infection. Cell 73, 457–467. Erickson, S. L., de, S. F., Kikly, K., Carver-Moore, K., Pitts-Meek, S., Gillett, N., Sheehan, K. C., Schreiber, R. D., Goeddel, D. V., and Moore, M. W. (1994). Decreased sensitivity to tumour-necrosis factor but normal T-cell development in TNF receptor-2-deficient mice. Nature 372, 560–563. Roggia, C., Tamone, C., Cenci, S., Pacifici, R., and Isaia, G. C. (2004). Role of TNF-alpha producing T-cells in bone loss induced by estrogen deficiency. Minerva. Med. 95, 125–132. Roggia, C., Gao, Y., Cenci, S., Weitzmann, M. N., Toraldo, G., Isaia, G., and Pacifici, R. (2001). Up-regulation of TNF-producing T cells in the bone marrow: A key mechanism by which estrogen deficiency induces bone loss in vivo. Proc. Natl Acad. Sci. USA 98, 13960–13965. Lam, J., Takeshita, S., Barker, J. E., Kanagawa, O., Ross, F. P., and Teitelbaum, S. L. (2000). TNF-alpha induces osteoclastogenesis by direct stimulation of macrophages exposed to permissive levels of RANK ligand. J. Clin. Invest. 106, 1481–1488. Saklatvala, J. (1986). Tumour necrosis factor alpha stimulates resorption and inhibits synthesis of proteoglycan in cartilage. Nature 322, 547–549. Lefebvre, V., Peeters-Joris, C., and Vaes, G. (1991). Production of gelatin-degrading matrix metalloproteinases (‘type IV collagenases’) and inhibitors by articular chondrocytes during their dedifferentiation by serial subcultures and under stimulation by interleukin-1 and tumor necrosis factor α. Biochim. Biophys. Acta. Mol. Cell. Res. 1094, 8–18. Shinmei, M., Masuda, K., Kikuchi, T., and Shimomura, Y. (1989). Interleukin 1, tumor necrosis factor, and interleukin 6 as mediators of cartilage destruction. Semin. Arthritis Rheum. 18, 27–32. Enomoto, M., Pan, H. O., Kinoshita, A., Yutani, Y., Suzuki, F., and Takigawa, M. (1990). Effects of tumor necrosis factor alpha on proliferation and expression of differentiated phenotypes in rabbit costal chondrocytes in culture. Calcif. Tissue Int. 47, 145–151.

Chapter 7 Prostaglandins and Proinflammatory Cytokines

127

178. Hashimoto, J., Yoshikawa, H., Takaoka, K., Shimizu, N., Masuhara, K., Tsuda, T., Miyamoto, S., and Ono, K. (1989). Inhibitory effects of tumor necrosis factor alpha on fracture healing in rats. Bone 10, 453–457. 179. Akira, S., Hirano, T., Taga, T., and Kishimoto, T. (1990). Biology of multifunctional cytokines: IL-6 and related molecules (IL-1 and TNF). F.A.S.E.B. J. 4, 2860–2867. 180. Hirano, T., Akira, S., Taga, T., and Kishimoto, T. (1990). Biological and clinical aspects of interleukin 6. Immunol. Today 11, 443–449. 181. Kishimoto, T., Taga, T., and Akira, S. (1994). Cytokine signal transduction. Cell 76, 253–262. 182. Tamura, T., Udagawa, N., Takahashi, N., Miyaura, C., Tanaka, S., Yamada, Y., Koishihara, Y., Ohsugi, Y., Kumaki, K., Taga, T., Kishimoto, T., and Suda, T. (1993). Soluble interleukin-6 receptor triggers osteoclast formation by interleukin 6. Proc. Natl Acad. Sci. USA 90, 11924–11928. 183. Al-Humidan, A., Ralston, S. H., Hughes, D. E., Chapman, K., Aarden, L., Russell, R. G. G., and Gowen, M. (1991). Interleukin-6 does not stimulate bone resorption in neonatal mouse calvariae. J. Bone Miner. Res. 6, 3–8. 184. Lowik, C. W., Van der, P. G., Bloys, H., Hoekman, K., Bijvoet, O. L., Aarden, L. A., and Papapoulos, S. E. (1989). Parathyroid hormone (PTH) and PTH-like protein (PLP) stimulate interleukin-6 production by osteogenic cells: a possible role of interleukin-6 in osteoclastogeneis. Biochem. Biophys. Res. Commun. 162, 1546–1552. 185. Linkhart, T. A., Linkhart, S. G., MacCharles, D. C., Long, D. L., and Strong, D. D. (1991). Interleukin-6 messenger RNA expression and interleukin-6 protein secretion in cells isolated from normal human bone: regulation by interleukin-1. J. Bone Miner. Res. 6, 1285–1294. 186. Ishimi, Y., Miyaura, C., Jin, C. H., Akatsu, T., Abe, E., Nakamura, Y., Yamaguchi, A., Yoshiki, S., Matsuda, T., Hirano, T., Kishimoto, T., and Suda, T. (1990). IL-6 is produced by osteoblasts and induces bone resorption. J. Immunol. 145, 3297–3303. 187. Roodman, G. D. (1992). Interleukin-6: an osteotropic factor? J. Bone Miner. Res. 7, 475–478. 188. Manolagas, S. C., and Jilka, R. L. (1995). Mechanisms of disease: Bone marrow, cytokines, and bone remodeling—Emerging insights into the pathophysiology of osteoporosis. N. Eng. J. Med. 332, 305–311. 189. Kurihara, N., Bertolini, D., Suda, T., Akiyama, Y., and Roodman, G. D. (1990). IL-6 stimulates osteoclast-like multinucleated cell formation in long term human marrow cultures by inducing IL-1 release. J. Immunol. 144, 4226–4230. 190. Black, K., Garrett, I. R., and Mundy, G. R. (1991). Chinese hamster ovary cells transfected with the murine interleukin-6 gene cause hypercalcemia as well as cachexia, leukocytosis and thrombocytosis in tumor bearing nude mice. Endocrinology 128, 2657–2659. 191. De La Mata, J., Uy, H. L., Guise, T. A., Story, B., Boyce, B. F., Mundy, G. R., and Roodman, G. D. (1995). Interleukin-6 enhances hypercalcemia and bone resorption mediated by parathyroid hormone-related protein in vivo. J. Clin. Invest. 95, 2846–2852. 192. Roodman, G. D., Kurihara, N., Ohsaki, Y., Kukita, A., Hosking, D., Demulder, A., Smith, J. F., and Singer, F. R. (1992). Interleukin 6. A potential autocrine/paracrine factor in Paget’s disease of bone. J. Clin. Invest. 89, 46–52. 193. Roodman, G. D. (2001). Biology of osteoclast activation in cancer. J. Clin. Oncol. 19, 3562–3571. 194. Yamamoto, T., Ozono, K., Kasayama, S., Yoh, K., Hiroshima, K., Takagi, M., Matsumoto, S., Michigami, T., Yamaoka, K., Kishimoto, T., and Okada, S. (1996). Increased IL-6-production by cells isolated from the fibrous bone dysplasia tissues in patients with McCuneAlbright syndrome. J. Clin. Invest. 98, 30–35. 195. Reddy, S. V., Takahashi, S., Dallas, M., Williams, R. E., Neckers, L., and Roodman, G. D. (1994). Interleukin-6 antisense deoxyoligonucleotides inhibit bone resorption by giant cells from human giant cell tumors of bone. J. Bone Miner. Res. 9, 753–757.

196. Devlin, R. D., Bone, H. G., and Roodman, G. D. (1996). Interleukin-6: a potential mediator of the massive osteolysis in patients with Gorham-Stout disease. J. Clin. Endocrinol. Metab. 81, 1893–1897. 197. Jilka, R. L., Hangoc, G., Girasole, G., Passeri, G., Williams, D. C., Abrams, J. S., Boyce, B., Broxmeyer, H., and Manolagas, S. C. (1992). Increased osteoclast development after estrogen loss: mediation by interleukin-6. Science 257, 88–91. 198. Poli, V., Balena, R., Fattori, E., Markatos, A., Yamamoto, M., Tanaka, H., Ciliberto, G., Rodan, G. A., and Costantini, F. (1994). Interleukin-6 deficient mice are protected from bone loss caused by estrogen depletion. E.M.B.O. J. 13, 1189–1196. 199. Saggese, G., Federico, G., and Cinquanta, L. (1993). In vitro effects of growth hormone and other hormones on chondrocytes and osteoblast-like cells. Acta. Paediatr. Suppl. 82 Suppl 391, 54–9. 200. Guerne, P. A., Carson, D. A., and Lotz, M. (1990). IL-6 production by human articular chondrocytes. Modulation of its synthesis by cytokines, growth factors, and hormones in vitro. J. Immunol. 144, 499–505. 201. Pincus, S. H., Schooley, W. R., DiNapoli, A. M., and Broder, S. (1981). Metabolic heterogeneity of eosinophils from normal and hypereosinophilic patients. Blood 58, 1175–1181. 202. Shingu, M., Miyauchi, S., Nagai, Y., Yasutake, C., and Horie, K. (1995). The role of IL-4 and IL-6 in IL-1-dependent cartilage matrix degradation. Br. J. Rheumatol. 34, 101–106. 203. Lotz, M., Vaughan, J. H., and Carson, D. A. (1988). Effect of neuropeptides on production of inflammatory cytokines by human monocytes. Science 241, 1218–1221. 204. Manolagas, S. C., Bellido, T., and Jilka, R. L. (1995). New insights into the cellular, biochemical, and molecular basis of postmenopausal and senile osteoporosis: Roles of IL-6 and gp130. Int. J. Immunopharm. 17, 109–116. 205. Romas, E., Udagawa, N., Zhou, H., Tamura, T., Saito, M., Taga, T., Hilton, D. J., Suda, T., Ng, K. W., and Martin, T. J. (1996). The role of gp130-mediated signals in osteoclast development: Regulation of interleukin 11 production by osteoblasts and distribution of its receptor in bone marrow cultures. J. Exp. Med. 183, 2581–2591. 206. Bellido, T., Stahl, N., Farruggella, T. J., Borba, V., Yancopoulos, G. D., and Manolagas, S. C. (1996). Detection of receptors for interleukin-6, interleukin-11, leukemia inhibitory factor, oncostatin M, and ciliary neurotrophic factor in bone marrow stromal osteoblastic cells. J. Clin. Invest. 97, 431–437. 207. Marusic, A., Kalinowski, J. F., Jastrzebski, S., and Lorenzo, J. A. (1993). Production of leukemia inhibitory factor mRNA and protein by malignant and immortalized bone cells. J. Bone Miner. Res. 8, 617–624. 208. Greenfield, E. M., Horowitz, M. C., and Lavish, S. A. (1996). Stimulation by parathyroid hormone of interleukin-6 and leukemia inhibitory factor expression in osteoblasts is an immediate-early gene response induced by cAMP signal transduction. J. Biol. Chem. 271, 10984–10989. 209. Ishimi, Y., Abe, E., Jin, C. H., Miyaura, C., Hong, M. H., Oshida, M., Kurosawa, H., Yamaguchi, Y., Tomida, M., Hozumi, M., and Suda, T. (1992). Leukemia inhibitory factor/differentiation-stimulating factor (LIF/D-factor): Regulation of its production and possible roles in bone metabolism. J. Cell. Physiol. 152, 71–78. 210. Reid, I. R., Lowe, C., Cornish, J., Skinner, S. J., Hilton, D. J., Willson, T. A., Gearing, D. P., and Martin, T. J. (1990). Leukemia inhibitory factor: a novel bone-active cytokine. Endocrinology 126, 1416–1420. 211. Lorenzo, J. A., Sousa, S. L., and Leahy, C. L. (1990). Leukemia inhibitory factor (LIF) inhibits basal bone resorption in fetal rat long bone cultures. Cytokine 2, 266–271. 212. Van Beek, E., Van der Wee-Pals, L., van de Ruit, M., Nijweide, P., Papapoulos, S., and Lowik, C. (1993). Leukemia inhibitory factor

128 inhibits osteoclastic resorption, growth, mineralization, and alkaline phosphatase activity in fetal mouse metacarpal bones in culture. J. Bone Miner. Res. 8, 191–198. 213. Palmqvist, P., Persson, E., Conaway, H. H., and Lerner, U. H. (2002). IL-6, leukemia inhibitory factor, and oncostatin M stimulate bone resorption and regulate the expression of receptor activator of NFkappaB ligand, osteoprotegerin, and receptor activator of NF-kappaB in mouse calvariae. J. Immunol. 169, 3353–3362. 214. Ware, C. B., Horowitz, M. C., Renshaw, B. R., Hunt, J. S., Liggitt, D., Koblar, S. A., Gliniak, B. C., McKenna, H. J., Papayannopoulou, T., Thoma, B., Cheng, L., Donovan, P. J., Peschon, J. J., Bartlett, P. F., Willis, C. R., Wright, B. D., Carpenter, M. K., Davison, B. L., and

Lawrence G. Raisz and Joseph A. Lorenzo Gearing, D. P. (1995). Targeted disruption of the low-affinity leukemia inhibitory factor receptor gene causes placental, skeletal, neural and metabolic defects and results in perinatal death. Development 121, 1283–1299. 215. Kawasaki, K., Gao, Y. H., Yokose, S., Kaji, Y., Nakamura, T., Suda, T., Yoshida, K., Taga, T., Kishimoto, T., Kataoka, H., Yuasa, T., Norimatsu, H., and Yamaguchi, A. (1997). Osteoclast are present in gp130-deficient mice. Endocrinology 138, 4959–4965. 216. Sims, N. A., Jenkins, B. J., Quinn, J. M., Nakamura, A., Glatt, M., Gillespie, M. T., Ernst, M., and Martin, T. J. (2004). Glycoprotein 130 regulates bone turnover and bone size by distinct downstream signaling pathways. J. Clin. Invest. 113, 379–389.

Chapter 8

Integrins and Other Adhesion Molecules M. H. Helfrich M. A. Horton

Department of Medicine and Therapeutics, University of Aberdeen, Institute of Medical Sciences, Foresterhill, Aberdeen AB25 2ZD, United Kingdom Bone and Mineral Centre, Department of Medicine, The Rayne Institute, London WC1E 6JJ, United Kingdom

V. Adhesion Molecules in Osteoclasts VI. Adhesion Molecules in Chondrocytes VII. Conclusion References

I. Abstract II. Introduction III. Molecular Structure of Adhesion Molecules IV. Adhesion Molecules in Cells of the Osteoblast Lineage

I. ABSTRACT

extracellular matrix. Chondroblasts, osteoblasts, and, to a lesser extent, osteocytes are responsible for the synthesis of the majority of the organic components of this matrix, whereas osteoclasts mainly degrade the matrix. The function of bone and cartilage cells reflects the matrix components that surround them and, conversely, the composition of the matrix, i.e. the structure of cartilage and bone, is highly dependent upon the cellular function of chondroblasts, osteoblasts, and osteoclasts. Adhesion molecules mediate cell–matrix and cell–cell interactions and much information has become available regarding the expression and role of one class of adhesion molecules in bone, the integrins. The knowledge about other classes of adhesion molecules in bone is also increasing as it is becoming clear that nonintegrin adhesion molecules, especially cadherins and members of the immunoglobulin family, are expressed and have functional roles in bone and cartilage cells. Adhesion molecules not only play a part in the function of fully differentiated cells in the skeleton, but also are increasingly implicated in cell differentiation. In this

Skeletal tissues contain a wealth of adhesion molecules. Integrins are the best-studied and largest group of these, but there is increasing information about expression and function of other classes of adhesion molecules such as cadherins, Ig family members and cell surface proteoglycans. In the skeleton adhesion molecules play a role in a wide range of cellular activities, including cell differentiation, tissue organization, matrix production and breakdown, and in the response to mechanical strain. This chapter summarizes the published information on expression, functional role and possible therapeutic or diagnostic use of adhesion molecules in bone and cartilage.

II. INTRODUCTION Connective tissue cells in general, and bone and cartilage cells in particular, are surrounded by an abundance of Dynamics of Bone and Cartilage Metabolism

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M. H. HELFRICH AND M. A. HORTON

chapter, we discuss the expression and function of the major adhesion molecules in bone and cartilage, focusing on data obtained with human tissue. We will discuss data from rodents and other species where little or no information in human tissue is available and to illustrate essential roles of adhesion molecules from knockout or transgenic studies.

III. MOLECULAR STRUCTURE OF ADHESION MOLECULES The main classes of adhesion molecules expressed in bone and cartilage are given in Tables I–IV. Adhesion molecules are classified according to structural homologies [1] and below we briefly discuss the basic structure, ligands, general function, and signal transduction of the main groups.

A. Integrins The integrin family consists of receptors that share as basic structure two noncovalently linked transmembrane glycoproteins (α and β subunits). Both subunits have a single transmembrane domain and, with the exception of β4, short cytoplasmic regions [2]. α and β subunits may associate with more than one partner forming receptors with distinct ligand-binding capabilities. At present 24 integrins are known, formed from 18 different α and eight different β subunits. The β subunits have higher cDNA sequence homology than the α subunits. They contain

Table I.

Osteoblast Adhesion Molecules

Integrins α1, α2, α3, α4, α5, αv, β1, β3, β5, β6, β8 Main integrin is α5β1 (also in osteocytes), other β1 integrins expressed depending on differentiation stage and, when analyzed in vitro, on culture conditions Low levels of αvβ3 compared to osteoclasts Cadherins E-cadherin, cadherin-4, cadherin-11, and N-cadherin No evidence for cadherin expression in osteocytes, but adherens junctions present Ig family members ICAM-1, ICAM-2, VCAM-1, LFA-3, and NCAM In vivo ICAM-1 confined to lining cells No data in osteocytes Syndecans Syndecan-1, -2, -3, and -4 Syndecan-3 mainly during embryogenesis and bone regeneration No data in osteocytes CD44 CD44H Also strongly expressed in osteocytes

Table II.

Osteoclast Adhesion Molecules

Integrins αvβ3, αvβ5, αvβ1, α2β1 β5 on osteoclast precursors only, replaced by β3 in mature cells LFA-1 (αLβ2) on osteoclast precursors only Expression of all other integrins excluded in mature osteoclasts Cadherins ?E-cadherin Functional evidence for a role of type 1 cadherins, but no definitive data on type expressed Ig family members ICAM-1 CD44 CD44H

56 cysteine residues (except β4 which has 48), which are organized into four 40 amino acid-long segments that are internally disulfide bonded. Their molecular weight ranges from 90–110 kDa, except for β4 which has a much longer cytoplasmic domain and a molecular weight of 210 kDa. The α subunits contain seven homologous repeating domains, folded into a propeller domain with three or four of the “blades” containing divalent cation-binding sites. These cation-binding sites, together with one such site in the β subunit, are critically important for ligand binding – note how both the α and the β subunit contribute to the ligand binding site – and also play a role in receptor stability. Some subunits (the collagen-binding α1, α2, α10, and α11 chains, αE and the leukocyte integrins αD, αX, αM, and αL) contain an additional 200 amino acids inserted between repeats 2 and 3, known as I domain. The I domain contains a characteristic metal ion-dependent adhesion site (MIDAS) motif, which can bind Mn2+ and Mg2+ ions and is also critical in ligand binding (reviewed by [2]).

Table III.

Chondroblast/Chondrocyte Adhesion Molecules

Integrins α1, α2, α3, α5, α6, α10, α11, αv, β1, β3, β5 Main integrin is α5β1 α4 and β2 reported, but only in osteoarthritic cartilage β7–9 and CD11 not studied Cadherins Cadherin-11 and N-cadherin Cadherin expression mainly during early chondrogenesis, no evidence for expression in mature cells Ig family members ICAM-1, VCAM-1, and NCAM NCAM mainly during early chondrogenesis Syndecans Syndecan-3 CD44 CD44H

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Chapter 8 Integrins and Other Adhesion Molecules

Table IV.

Functional Role of Adhesion Molecules in Bone and Cartilage

Functional role Osteoblast Adhesion Molecules β1 integrins Adhesion to matrix

av integrins

Class 1 cadherins: N-cadherin E-cadherin Cadherin-4 and 11 Ig superfamily

Syndecans

CD44

Ig superfamily

Key references

Native collagens, fibronectin, osteopontin, bone sialoprotein, e.o. Most likely fibronectin, RGD

[25, 28, 59]

[59]

Type I collagen

[29, 52–56]

α5β1 α5β1

Fibronectin, RGD Fibronectin

[50] [60, 75]

αv integrins

Vitronectin, osteopontin, ? others Vitronectin, osteopontin ?others Class 1 cadherins

[49]

β1 integrins β1 integrin, probably α5β1 α2β1, α1β1

Osteoblast differentiation through facilitation of BMP-2 effects Osteoblast–osteoblast adhesion, differentiation, gene regulation, and bone formation

αvβ3, αvβ5, αvβ6, αvβ8 Class 1 cadherins

Mechanosensing Production of cytokines

Class 1 cadherins ICAM-1, VCAM-1

Osteoclast differentiation, through “juxtacrine stimulation” of RANKL expression Matrix adhesion, boundary formation during embryogenesis Osteoblast proliferation through regulation of growth factor signaling Osteoblast recruitment to areas of bone formation Hyaluronate degradation Matrix adhesion

ICAM-1

Osteoclast survival Osteoclast polarization and signal transduction Inhibition of osteoclast formation Osteoclast development and fusion ? Clear zone formation, bone resorption Osteoclast differentiation

?Osteoclast fusion

CD44

Ligand bound

Osteoblast and osteocyte organization, polarity and matrix secretion Mediation of differentiation signals in early osteoblasts Osteoblast proliferation Mechanosensing and mechanotransduction Adhesion to matrix

Regulation of osteoclast formation through up-regulation of ICAM-1 and V-CAM-1 Osteoclast Adhesion Molecules b1 integrins Adhesion to matrix, ?cessation of resorption av integrins Adhesion to matrix, ?cessation of resorption

Cadherins

Molecules involved

Osteoclast migration, fusion and bone resorption

[35] [85, 89, 90]

Class 1 cadherins LFA-1 (αLβ2) or α4 integrins on leukocytes LFA-1 on osteoclast precursors

[76] [105]

Syndecan-1, -2, and -4 Syndecan-2

Tenascin-C, type 1 collagen

[117]

GM-CSF

[121]

Syndecan-3

HB-GAM

[118]

CD44 CD44

[126] No definitive data

CD44

Hyaluronate Type I collagen, fibronectin, laminin, osteopontin ? Hyaluronate or other

α2β1

Native collagen

[138]

αvβ3

Vitronectin, osteopontin, bone sialoprotein, fibronectin, fibrinogen, denatured collagen, RGD As above As above

[260]

As for αvβ3, RGD E-cadherin Class 1 cadherins (HAV)

[161] [176] [178]

ICAM-1 on osteoblasts

[26, 107]

ICAM-1 on osteoclast precursors

[107]

Hyaluronate, osteopontin, ?others

[181, 182]

αvβ3 αvβ3 αvβ5 E-cadherin Class 1 cadherins (HAV) LFA-1 on osteoclast precursors LFA-1 on osteoclast precursors CD44

[26, 106]

[104]

[157] [163, 175, 261]

Continued

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M. H. HELFRICH AND M. A. HORTON Table IV.

Functional Role of Adhesion Molecules in Bone and Cartilage—cont’d

Functional role Chondroblast/Chondrocyte Adhesion Molecules b1 integrins Matrix adhesion

Cadherins

Chondroblast organization in the growth plate, cytokinesis Matrix degradation Cell survival Mechanosensing Cartilage development

Ig superfamily

Cartilage development Cartilage breakdown

Syndecans

Cartilage development Chondrocyte proliferation Pericellular matrix assembly Matrix degradation

CD44

Molecules involved

Ligand bound

Key references

α1β1, α2β1, α3β1, α5β1 β1 integrin(s)

Types I, II, and VI collagen and fibronectin Fibronectin (RGD)

[214, 221, 222]

α5β1 β1 integrins α5β1 Cadherin-11 and N-cadherin NCAM ICAM-1 and VCAM-1 Syndecan-3 Syndecan-3 CD44 CD44

Fibronectin (RGD) Collagens and fibronectin Fibronectin Cadherin-11 and N-cadherin

[218] [225, 226] [77] [235–237]

N-CAM LFA-1 and α4 integrins on leukocytes Tenascin-C FGF-2 Hyaluronate, others? Hyaluronate, osteopontin, collagen, fibronectin, ?others

[236] [242, 244, 245]

Recent studies suggest that the cation-binding site in β subunits may be homologous to a MIDAS motif (see reference [3]). The first electron microscopic studies to define the structure of the fibronectin receptor α5β1 [4] showed that the large extracellular domain is organized as a globular head supported by a stalk, composed of one α and one β subunit leg with overall dimensions of ~10 × 20 nm. This basic three-dimensional structure has now been confirmed in a number of integrins and more recently has been refined by detailed analysis of the crystal structure of the extracellular domain of the αvβ3 integrin, both with and without bound ligand [5, 6]. These studies revealed that the integrins not only exist as straight stalk structures, but can also bend over with the globular head facing towards the C-termini of the legs projecting from the plasma membrane. There has been extensive research to understand how the conformation of integrins changes upon cation binding. This generally results in “activation”, a conformation in which the receptor can bind ligand and is important especially for the β2- and β3-containing integrins. The majority of data suggests that the bent form of the integrin is the inactive, nonligand-bound state, whereas straightening is associated with activation and ligand binding. These switches between active and inactive conformations are comprehensively reviewed elsewhere [2, 3]. Post-translational modification of integrins include the cleavage of the α subunits lacking I domains, with the exception of α4, into heavy and light fragments that are disulfide linked, and glycolysation of α and β chains. Ligands for integrins include a wide range of extracellular matrix proteins and Ig family members such as

[209]

[248] [249] [254, 255] [256]

VCAM and ICAM, reviewed in [3]. Most integrins can bind more than one ligand and often this is through recognition of a common sequence. In many extracellular matrix molecules this is an RGD peptide sequence, although in other proteins sequences such as DGEA (in collagen), or LDV (in fibronectin) are recognized. Integrins are linked to the cytoskeleton via interaction of the β subunit with actin-binding proteins α-actinin, vinculin, and talin. The cytoplasmic domain of the β subunit also associates with a signaling complex comprising kinases and phosphatases and various adaptor proteins. Much information regarding signaling via integrins has come from studies in cells such as fibroblasts and osteosarcoma cell lines, which produce focal contacts in vitro containing focal adhesion kinase (FAK), which associates directly with the β subunit. Upon occupation of the integrin receptor and clustering of receptors, FAK is targeted to focal adhesions, where it associates with the cytoskeleton and is activated by autophosphorylation. Downstream signaling pathways include association of Src family kinases with phosphorylated FAK and engagement and activation of Rho-like GTPases, Erk, and Jnk signaling pathways, ultimately leading to cytoskeletal rearrangement [7].

B. Cadherins Cadherins are a rapidly growing family (>80 members) of calcium-dependent proteins that play prominent roles in morphogenesis and the maintenance of adhesive contacts in solid tissues. They are divided into five subsets of receptors: the classical cadherin types I and II, the latter

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directly linked to the actin cytoskeleton, desmocollins and desmogleins, protocadherins, and a number of other, more distantly related cadherins [8, 9]. Cadherins are calciumdependent single-chain single-pass transmembrane glycoproteins that share “cadherin domains”, which contain specific amino acid motifs, which have essential roles in calcium binding and dimerization of the receptors [8, 10]. In the context of this review the classical cadherins are the most important subgroup. The type I cadherins (members E-, N-, M-, P-, and R-cadherin) all share a common structure and have highest homology in the short cytoplasmic domain. They have an extracellular domain containing five repeats of around 110 amino acids which contain the negatively charged, calcium-binding motifs and conserved cysteine residues in the fifth repeat. They have molecular weights of around 100–130 kDa. The ligand-binding site of type I cadherins is the conserved HAV motif located at the N-terminal region of the molecule in the first conserved extracellular repeat. Type II cadherins (cadherin 5–12) share the basic cadherin structure, but have lower amino acid homology with type I cadherins. They also lack the cell adhesion HAV motif [8]. Cadherins are mediators of cell–cell adhesion and bind ligand mainly in a homophilic manner, although heterophilic binding between different cadherin molecules is possible, but appears restricted to cadherins from the same class [9]. As with integrins, cells often express a repertoire of different cadherins simultaneously, although a particular feature of cadherins is their restricted expression at specific stages of embryonic and cellular development. On the cell surface, cadherins tend to be concentrated at cell– cell junctions. The cytoplasmic tail of cadherins binds to catenins and this complex is important in gene transcription as well as regulation of adhesion. β Catenin is also the key player in the Wnt signaling pathway, a pathway that has recently been found to be important in the regulation of bone mass [11] and current work is trying to unravel interconnections between Wnt signaling and cadherinmediated cell adhesion through regulation of free β catenin levels in cells [12]. Signal transduction through cadherins is complex, dependent upon cell type and, in addition to phosphorylation of β-catenin, can include activation of small GTPases and tyrosine kinase pathways [13].

C. Immunoglobulin Superfamily The immunoglobulin (Ig) family of receptors all share a basic motif consisting of an Ig fold of between 70 and 110 amino acids (reviewed in [1, 14]) organized into two antiparallel β sheets which seem to serve as a scaffold on which unique determinants can be displayed. There is

considerable variation in the primary structure of the members of this family and hence in molecular weights, but their tertiary structure is well conserved. There are now over 700 human genes known which share Ig motifs, making this one of the largest superfamilies in the human genome [15]. Many of these are splice variants of cell adhesion molecules and Ig genes, but overall the cell adhesion molecules constitute a large part of this superfamily. Here we are concerned only with the cell adhesion molecules. Their functions are wide ranging with some members functioning as true signal-transducing receptors, whereas others have predominantly adhesive functions. Ligands for Ig family members include other Ig family members (identical, as well as nonidentical members), such as NCAM binding to itself, but also members of the integrin family (such as for the ICAMs, which bind β2 integrins) and components of the extracellular matrix, for example collagen binding for myelin-associated glycoprotein, a neuron-expressed Ig family member [1]. Signaling pathways activated by Ig family members include MAP kinase pathways [1, 16].

D. Syndecans Syndecans are a family of cell surface proteoglycans, varying in size from 20–45kDa. They are type I transmembrane proteins characterized by a shared heparan sulfate attachment sequence on the N-terminus of their single polypeptide chain. There is little homology in the rest of their extracellular domain, but their single transmembrane domain and short cytoplasmic domains are highly conserved. The cytoplasmic domain contains four conserved tyrosine residues, suggesting that phosphorylation of this domain could occur and be involved in signal transduction events. However, syndecans are thought to function predominantly as co-receptors for other receptors such as integrins, and members of the fibroblast growth factor family and vascular endothelial cell growth factor, which need heparan sulfate for signaling. In such situations the signaling is thought to occur via the cytoplasmic domains of associated receptors rather than the syndecan molecule [17]. There are four mammalian syndecans known, with syndecan-1 best studied so far. Syndecan-1 is the major syndecan of epithelia and can function as a cell–matrix receptor binding various matrix proteins (type I collagen, fibronectin, tenascin), and in addition can bind members of the FGF family. It appears that in different cell types syndecan-1 can have different patterns of glycosaminoglycans attached to its core protein and this influences the ligand-binding capabilities. Thus, where in one cell type syndecan-1 may contain heparan sulfate, as well as

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chondroitin sulfate side chains and bind collagen, this may not be the case in another cell type in which it has heparan sulfate side chains only. Syndecan-2 is the main form in mesenchymal cells and is present in neuronal tissues alongside syndecan-3. Syndecan-4 is present in many cells that form stable adhesions and in vitro is consistently present in focal adhesions [18]. The functions of syndecans are varied with roles in cell–matrix interaction and cell proliferation. Signal transduction, in addition to associated tyrosine kinsases, may also include cytoskeletal proteins through association with actin and tubulin.

E. CD44 CD44, also known as the hyaluronate receptor, is a family of transmembrane glycoproteins with molecular weights of 85–250 kDa. They share an N-terminal region that is related to the cartilage proteoglycan core and link proteins. Alternative splicing of ten exons and extensive post-translational modification such as glycosylation and addition of chondroitin sulfate produces the wide variety of CD44 proteins. Chondroitin sulfate containing variants can bind fibronectin, laminin, and collagen in addition to hyaluronate, and CD44 also binds to osteopontin [19]. Finally, CD44 may also bind homotypically. CD44 therefore functions in a variety of ways including in cell–cell interaction (homing of lymphocytes or cell clustering), but also in cell–matrix adhesion. Malignant transformation of cells leads to up-regulation of CD44 expression and metastatic tumors often express an altered repertoire of CD44 variants. More recently evidence has pointed to a more complex role of CD44 than simply as a cell–cell or cell–matrix adhesion molecule. It is becoming apparent that CD44 can be enzymatically cleaved and that functional fragments from the cytoplasmic domain can act as transcriptional regulators, whereas functional fragments from the ectodomain circulate in body fluids or can become incorporated in the matrix and thereby modulate cellular behavior [20].

74–240 kDa, with differently glycosylated forms expressed in different cell types. In general, the function of selectins is in leukocyte trafficking. They are thought to be involved in the earliest stages of leukocyte extravasation, where binding of the selectin ligand on the leukocyte to selectins expressed on the endothelial surface results in weak intercellular interactions and “rolling” of leukocytes over the endothelial surface. These functions have now been confirmed in selectin knockout mice for L- and P-selectin. In a double knockout for E- and P-selectin, it appeared that E-selectin also contributes to leukocyte homeostasis (reviewed in references [14, 21]). Other functions for selectins are in β2 integrin activation and O2− production by leukocytes. Selectin ligands are specific oligosaccharide sequences in sialated and often in sulfated glycans, such as sialyl-Lewisx. There remains some uncertainty about the natural ligands for selectins. Many have been identified in in vitro assays, using recombinant selectins presented in multimeric form or in clustered arrays that increase affinity for ligand, but these have no known in vivo equivalent (reviewed in reference [22]). However, some molecules seem to fulfill most of the criteria for true selectin ligands (i.e. with confirmation of a biological role), such as CD24 and P-selectin glycoprotein ligand-1 (PSGL-1), and when expressed in the neutrophil surface in a properly glycosylated and tyrosine sulfated form can bind to P-selectin in vascular endothelium and to L-selectin on other neutrophils to enable “rolling” over either surface. CD34, GlyCAM-1, and MAdCAM-1 are other candidate ligands for L-selectin and E-selectin ligand-1 for E-selectin (reviewed in reference [23]). The signal transduction pathways linked to selectins are only partially elucidated [24]. As with the other classes of adhesion molecules discussed above they include tyrosine phosphorylation cascades and increases in intracellular calcium.

IV. ADHESION MOLECULES IN CELLS OF THE OSTEOBLAST LINEAGE A. Integrins

F. Selectins Selectins are a family of three closely related glycoproteins (P- and E-selectin expressed in endothelial cells and L-selectin expressed in leukocytes). Their common structure consists of an N-terminal Ca2+-dependent lectintype domain, an EGF domain, and variable numbers of short repeats homologous to complement-binding sequences, a single transmembrane region and a short cytoplasmic domain. Their molecular weights range from

In vivo osteoblasts in normal human bone have been reported to express β1 and β5 integrins [25–33] and most recently reviewed by Bennett et al. [34], but there remain controversies regarding the α subunits associated with β1 and concerning expression of αvβ3 (Table I). β2 and β4 integrins have not been reported in osteoblastic cells. Recently cultured human primary osteoblasts were for the first time reported to also express αvβ6 and αvβ8 integrin [35]. There are considerable differences between various reports on the integrin repertoire expressed by

Chapter 8 Integrins and Other Adhesion Molecules

osteoblasts in situ, by human osteoblast cultures, or by murine osteoblastic cell lines and these are most likely due to the heterogeneity of the cell population under study and the different culture conditions employed. Moreover, it is becoming clear that osteoblast activation status (e.g. “real” osteoblasts, synthesizing bone matrix versus the synthetically inactive bone-lining cell), or anatomical site (e.g. osteoblast phenotype in endochondral versus intramembranous bone), or the presence of bone disease, influences integrin expression, but no definitive studies have been published. For example, it has been clearly demonstrated that in vitro, the substrate on which cells are cultured directly influences the pattern of integrin expression by osteoblasts [36, 37]. Generally, cultured osteoblasts express a wider integrin repertoire than osteoblasts in situ [25, 32] and in particular increased expression of the vitronectin receptor αvβ3 is seen. It is still unclear whether altered expression of integrins has an equivalent in bone pathology, since osteoblast integrin expression in bone disease is relatively unexplored, but bone diseases such as Paget’s disease and osteopetrosis are accompanied by major changes in the numbers of “real” osteoblasts, i.e. osteoid synthesizing cells [38], and hence differences in integrin expression in the overall osteoblastic population are likely. Also, recent studies are beginning to examine the effects of mechanical stimuli on integrin expression and suggest that external stimuli may influence the integrin expression in the osteoblastic population. Integrin expression in osteocytes has not been studied extensively, but there are some data from immunocytochemical analysis of human bone sections and from functional studies in the chick, where isolation procedures for osteocytes have been developed. Chick osteocytes and osteoblasts bind a comprehensive range of extracellular matrix proteins in vitro in a β1- and partially RGDdependent way [39]. The exact receptor involved was not determined, because of lack of α chain-specific antibodies for avian integrins. Expression of β1 integrin has been confirmed in mammalian osteocytes [31, 40], including the osteocytic cell line MLO-Y4 [41]. However, there are few definitive reports where integrin expression has been followed throughout osteoblastic differentiation to osteocytes within the same species. Difficulties in interpretation of immunocytochemical staining in bone sections, in particular for cells embedded within matrix, have been reported by many authors and this may well have contributed to the continuing controversies in integrin phenotype of osteoblasts/osteocytes. From the limited reports on isolated cells at different stages of differentiation, it has become clear that, at least in the mouse, α5β1 is the most abundantly expressed integrin throughout osteoblastic differentiation and that, maybe surprisingly,

135 α2β1 is expressed at much lower levels, in particular in more differentiated cells [41]. Expression of α1 and α6 integrins has been reported in mesenchymal stem cells [42] and Stewart et al. [43] recently showed that selection of α1 integrin-positive bone marrow cells markedly enriched the population of CFU-F, clonogenic precursors with osteogenic potential, confirming the importance of α1 early in the osteoblast lineage. Human osteogenic stem cells also express β1 integrin [44]. Current interest in differentiation of osteogenic cells from stem cells [44, 45] may reveal additional information on expression of integrins and other cell adhesion molecules during differentiation, an area which is still poorly understood (see also reference [34]). This will require comprehensive studies in well-defined osteoblastic populations (both in developmental stage and in synthetic activity) combining immunocytochemical, biochemical, and molecular techniques. Functional studies on osteoblast integrins initially concentrated on their role in adhesion. In keeping with their extensive repertoire of integrins, rodent and human osteoblasts were found to adhere to osteopontin, bone sialoprotein, vitronectin, and fibronectin in an RGD-dependent way (with fibronectin requiring much higher concentrations of peptide for inhibition), whereas binding to type I collagen and thrombospondin was less inhibited by RGD peptides [25, 28, 46]. In organ cultures of mineralizing fetal rat parietal bone, RGD peptides decreased bone formation accompanied by a decrease in α2 and β1 expression and disruption of the organization in the osteoblast layer [47]. Down-regulation of α2 and β1 integrin expression in osteoblasts by glucocorticoids has also been noted with similar disruption in osteoblast organization, whereas IGF-1 increased β1 expression in osteoblasts and increased calcified bone formation [40, 48]. Equally, it was shown that, in vitro at least, αvβ3 and αvβ5 integrins, which regulate adhesion to vitronectin and osteopontin, are down-regulated by long-term glucocorticoid exposure [49], a phenomenon that, together with the negative effect on β1 integrin, might help to explain the bone loss associated with long-term glucocorticoid usage. More recently it has become clear that the role of osteoblast integrins extends beyond adhesion. BMP-2, a potent stimulator of bone formation, increases expression of the whole repertoire of osteoblast αv integrins and the BMP-2 receptor colocalizes with integrins. The finding that blocking of the function of αv inhibited the BMP-2 effects in osteoblast, suggests that integrins may regulate BMP-2 effects in bone [35] and suggests that there is potentially a wider role for integrins in facilitating growth factor signaling in bone. Functional studies on osteoblast integrins also include reports on their role in osteoblast differentiation.

136 Moursi et al. [50] demonstrated that osteoblast–fibronectin interaction was a critical event in the differentiation of fetal rat osteoblasts in an in vitro model and the importance of the central cell-binding domain of fibronectin in this effect suggested that the α5β1 fibronectin receptor was implicated. Later studies by this group also suggested a role for α3β1 in osteoblast differentiation [51]. In addition to fibronectin, collagen is also implicated in osteoblast differentiation. Ganta et al. [29] showed that ascorbic acid deficiency, which leads to under-hydroxylation of type I collagen, resulted in down-regulation of α2β1 in osteoblasts and dysregulation of differentiation and mineralization in cultures of rat calvaria. Jikko et al. [52] showed that α1 and α2 integrins mediate differentiation signals, such as from BMP-2 in early osteoblastic cells. Minzuno et al. [53] demonstrated a role for α2β1 and type I collagen in osteoblast differentiation from early progenitors and Xiao et al. [54–56] provided evidence that binding of α2β1 to ligand leads to expression of differentiation-associated genes such as Runx2, alkaline phosphatase, and osteocalcin. These data correlated well with the higher levels of α1 and α2 collagen-binding integrins in early differentiation stages of the lineage and reduced expression of these integrins in more differentiated cells as reported above. Since β1 integrins have a critical role in embryogenesis, not much information has been gained from studies of β1 knockout mice, which are embryonic lethal at a time well before skeletal development begins [57, 58]. Zimmerman et al. [59], therefore generated a transgenic mouse in which a dominant negative β1 integrin is expressed under the control of the osteoblast-specific osteocalcin promoter, resulting in expression in mature osteoblasts and in osteocytes. In vitro, osteoblasts from these animals did not properly adhere to matrix, whereas in vivo, osteoblast and osteocyte morphology, polarity and matrix secretion was severely affected, resulting in decreased bone mass, especially in females. Anatomical differences were found in older animals [60]. The tibias of the transgenic animals were straighter, consistent with the possibility that in absence of functional β1 integrin, osteoblastic cells are not able to sense and/or respond adequately to load bearing, which in wild-type animals results in the characteristic curvature of this long bone. Viable knockouts have been generated for α1 and α2 integrins. Surprisingly, neither has a prominent bone phenotype [61–63], although detailed bone histology has not been reported. It is possible that compensation occurs in these single knockouts for collagenbinding integrins, since overall the absence of gross abnormalities in any tissue in the α2 null was largely unexpected [64]. The α1 null demonstrates abnormalities

M. H. HELFRICH AND M. A. HORTON

during fracture healing, which are, however, related mainly to slow cartilage proliferation, rather than abnormalities in osteoblasts [63]. The α3/α6 double knockout has severe developmental deficiencies in limb formation, but no data exist for a role in adult mice (see reference [65] for a review on all integrin deletion phenotypes). Single knockouts for β3 and β5 and αv and a double knockout for β3/β5 have been made (see reference [66]) and none of these appear to have osteoblast malfunction. The relatively subtle defects seen in the skeleton of the β3 knockout are caused by osteoclast malfunction and are discussed further below. An area of increasing interest is that of the possible role of adhesion molecules, especially integrins, in mechanosensing in bone. Osteocytes and osteoblasts are known to be highly responsive to mechanical effects on bone. In other cell systems it has been demonstrated that twisting or turning of integrin molecules directly affects gene transcription [67]. Stretching of cells, i.e. change of cell shape and dimension, has also been shown to affect gene transcription and cell survival [68] and cell shape is largely controlled by extracellular matrix [69]. It is therefore tempting to speculate that positive mechanical effects on bone cells resulting in bone formation may be mediated indirectly via adhesion receptors and/or cellular shape changes resulting from the bone deformation. In contrast, absence of mechanical stimuli which leads to a decrease in bone mass may be the result of lack of “cellular stretch” in particular in osteocytes, which may lead to apoptotic cell death [68, 70], a cellular phenomenon associated with induction of bone resorption [71]. Despite these plausible hypotheses, the mechanisms whereby bone cells sense and respond to mechanical strain remain largely unresolved, as it remains technically challenging to subject bone cells, in particular osteocytes to defined levels of strain and measure single cell responses. A further complication is that osteocytes/osteoblasts in vivo exist as a syncytium and these cell–cell interactions are not readily re-established in vitro. Despite these difficulties some progress has been made recently. Charras and Horton used atomic force microscopy to estimate the cellular strain necessary to elicit cellular responses in bone cells [72] and then went on, using finite element modeling, to determine the strain exerted on individual cells by a number of common techniques used for mechanical stimulation of cells in vitro [73], such as the magnetic bead twisting and stretch experiments described above. Interestingly, fluid shear stress, the mechanisms by which osteocytes have been thought to detect strain in bone [74], gives rise to the lowest cellular strain (deformation) and it is possible that cells may have different mechanisms for detecting different magnitudes

Chapter 8 Integrins and Other Adhesion Molecules

of strain [73]. Using paramagnetic beads coated with specific integrin antibodies, Pommerenke et al. [75] were able to show that drag forces (shear) applied to α2 and β1 integrin subunits resulted in a cellular response, in this case intracellular calcium increase. Other studies (further discussed below) indicate that cadherins [76] are also directly involved in the response to stain and there is strong evidence for integrin-mediated mechanosensing in chondrocytes (see reference [77]). Although further information is clearly needed, especially in osteocytes, the principle that integrins and other classes of adhesion molecules are involved in mechanosensing appears well established. Taken together, the available evidence from in vivo and in vitro studies confirms that β1 integrins are the main integrins in the osteoblastic lineage and have a variety of functions (Table IV). In addition to providing an anchor to the bone matrix, they are important in the differentiation of osteoblasts, in the expression of differentiation-associated genes, in the response to growth factors and ultimately in the maintenance of bone mass [78], possibly through a role in mechanosensing. Unsurprisingly, there is great interest in the orthopedics/biomaterials field in identification of bioactive surface coatings to enhance osteoblast attachment and matrix (reviewed in reference [79]). Coating of biomaterials with fibronectin has been shown to give the best adhesion of osteoblastic cells, which is completely in agreement with all data presented above. Many studies integrating adhesive proteins with micro- and nano-topography on biomaterials are underway and will increase our understanding of integrin–matrix interactions and applications in tissue engineering.

B. Cadherins Cell–cell interactions between osteoblastic cells themselves and osteoblasts and hemopoietic cells within the bone marrow compartment are known to be important in bone metabolism. Such interactions are likely to be governed by adhesion molecules such as cadherins. Indeed, osteoblasts have been found to express a number of cadherins, in particular E-cadherin, cadherin-4, cadherin-11 (OB-cadherin), and N-cadherin [80–82] and, in culture, osteoblasts form tight cellular junctions containing cadherins [83]. Some differences have been found between different species, skeletal site and differentiation stage of the osteoblasts, but it is clear from a range of immunocytochemical and molecular studies that N-cadherin and cadherin-11 are abundantly expressed in osteoblastic cells in many mammalian species (reviewed in reference [84]). Cadherin expression has been studied during the

137 differentiation of early mesenchymal cells into mature osteogenic cells both in vivo and in vitro. N-cadherin appears to be expressed widely in all mesenchymal lineage cells and remains expressed at all stages of bone formation with especially high levels of expression at late stages of differentiation [85]. Cadherin-11 seems to be more specifically associated with the osteoblast lineage [86]. Cadherin expression appears to be lost in osteocytes [86, 87]. It has been suggested that this is linked to osteoblast apoptosis at the end of a formation cycle and is necessary to allow a proportion of mature osteoblasts to enter into the osteoid and become osteocytes. However, it is somewhat surprising that cadherins might not contribute to maintaining the cellular contacts between osteocytes and surface osteoblasts and more information is required on the molecules that form the cell–cell contacts in this network. There is good functional evidence that cadherins are important for bone cell function (Table IV). HAV peptides inhibit cell–cell contacts in osteoblast cultures and matrix formation in vitro, suggesting a role for class I cadherins in osteoblast synthetic activity [85, 88]. Likewise, antibodies to E-cadherin inhibit cell–cell adhesion [89]. Expression of a dominant negative N-cadherin in vitro in committed pre-osteoblastic cells as well as more differentiated osteoblasts inhibited expression of genes associated with bone formation and mineralization [85, 90]. Recently, cadherins were implicated in the response to mechanical stimulation in cultured osteoblasts by the finding that fluid shear stress increased translocation of β-catenin to the nucleus and regulated COX-2 expression in osteoblasts [76]. In mice in which the gene for cadherin 11 is deleted, a reduction in bone density is seen, strongly implying a role for cadherin-11 in osteogenesis [91]. At present there is no in vivo information about the bone phenotype in absence of N-cadherin since the N-cadherin-null mice die at day E10, before mature osteoblasts are present [92]. However, using a dominant negative approach, Castro et al. [93] showed that a reduction in functional N-cadherin in osteoblasts in vivo leads to a reduction in osteoblast number and thus in peak bone mass, whilst the number of adipocytes was increased possibly via changes in mesenchymal cell lineage commitment. Osteoblast cadherins are up-regulated, through as yet unknown mechanisms, by a variety of cytokines and hormones including BMP-2, FGF-2, PTH, and downregulated by IL-1 and TNFα (reviewed in references [84, 94]), and it is plausible that some of the well-known effects of these hormones are mediated in part through modulation of cadherin function. Application of stretch to osteoblasts in vitro selectively up-regulated expression

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of N-cadherin [95], leading to increased cell–cell adhesion. The signaling pathways controlling cell–cell adhesion through osteoblast cadherins and the ways they regulate osteoblastic gene expression are as yet unclear. Finally, osteoblast cadherins are implicated in malignancy: N-cadherin expression is reduced and cadherin-11 is not normally displayed on the cell surface in osteosarcoma [96] and β-catenin mutations are found in malignant bone tumors [97]. Up-regulation of E- and N-cadherin is found in Apert syndrome (discussed in reference [84]), and since cadherin expression is differentially regulated in osteoblastic versus adipocytic cell populations [93, 98] cadherins may play a role in development of osteoporosis and osteoarthritis, diseases associated with changes in the differentiation of mesenchymal cells to osteogenic versus adipocytic cells.

C. Ig Family Members A number of Ig family members are expressed in osteoblasts. During skeletal development transient expression of NCAM is seen [99–101]. In mature osteoblasts expression of ICAM-1, and ICAM–2, VCAM-1, and LFA-3 have been reported [26, 99, 102–108] and VCAM-1 is also seen in early osteogenic cells [44]. There is circumstantial evidence for the expression of NCAM in human osteoblasts from studies in multiple myeloma, where NCAM antibodies blocked IL-6 production in osteoblast–myeloma co-cultures [109]. Similarly, crosslinking of ICAM-1 or VCAM-1 resulted in the production of bone resorbing cytokines by osteoblasts [105], suggesting that, in general, expression of Ig family members by osteoblasts may lead to direct cellular interaction with immune cells, which express their ligands (Table IV) and subsequently to activation of production of bone-resorbing cytokines. In addition, Ig family adhesion molecules expressed on osteoblasts are firmly implicated in osteoclast formation. VCAM-1 has been shown to be involved in the development of osteoclasts in vitro, an effect that could be mediated via osteoblastic cells [110]. Harada et al. [107] found evidence for involvement of ICAM-1 and its ligand LFA-1 in osteoclastogenesis, suggesting a role not only for osteoblast–osteoclast interaction, but also for osteoclast precursor fusion, enabled by transient expression of LFA-1 in pre-osteoclasts. The role of ICAM-1 has been studied in more detail recently. Tanaka et al. [103] found that ICAM-1 expression characterized a population of osteoblasts, induced by pro-inflammatory cytokines, that have arrested in G0 /G1 and are uniquely equipped to sustain osteoclast differentiation. Interestingly, Everts et al.

described that the only cells positive for ICAM-1 on the bone surface in resorbing mouse calvaria are bone-lining cells [111]. Linking roles for integrins and Ig family members, Nakayamada et al. [26] reported how, following β1-dependent osteoblast adhesion, signaling through FAK results in up-regulation of ICAM-1 and RANKL leading to osteoclast formation. Further molecules that have been found to induce ICAM-1 expression in rodent osteoblasts in vitro, leading to increased osteoclast formation, are PTH, IL1, TNFα, and 1,25-D3 [106, 112]. The same stimuli also independently induce expression of RANKL and it is becoming clear that efficient stimulation of osteoclast precursors by RANKL is dependent upon high-affinity adhesion between osteoblast and osteoclast precursors, a process coined as “juxtacrine stimulation” [106]. Furthermore, crosslinking of CD44 in osteoblasts results in strong up-regulation of ICAM-1 and VCAM-1 [104], adding yet another mechanism by which bone metabolism and in particular osteoclastogenesis may be regulated, in this case through a CD44 ligand such as, for example, hyaluronan or osteopontin. ICAM-1 expression on osteoblasts in vivo in pathologies has yet to be studied. Expression of human primary osteoblasts in vitro is highly variable, but has been associated with pathological status: increased expression was seen in cells derived from patients with osteoporosis and to a lesser extent patients with osteoarthritis, compared to normal individuals, and increased expression was related in particular to the presence of elevated levels of IL6 and PGE2 [113]. These data fit well with the established role of ICAM-1 in inflammatory diseases in general and especially in rheumatoid synovium, where levels are considerably increased during active periods of disease and where anti-ICAM-1 therapy has been considered [114, 115]. Taken together with the experimental data it seems likely that high osteoblastic ICAM-1 levels are involved in the mechanism of diseases associated with bone loss by increasing osteoclast recruitment and formation.

D. Syndecans The expression of these molecules in bone is best studied during differentiation. Syndecan-3 (also known as N-syndecan) expression is found in cartilage (see below) and periosteum during endochondral bone formation, where it interacts with tenascin to set boundaries in tissues [116], but low levels are still seen in osteoblasts and osteocytes close to periosteal surfaces [117] and higher levels are found in osteoblasts and precursors in areas of bone regeneration after damage [118]. Messenger RNA for the three cloned human forms of

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syndecan-1, -2, and -4 are found in primary human osteoblast cultures and a number of osteoblast cell lines [119, 120], where expression levels are regulated by cytokine-induced tissue regeneration [119]. Culture of osteoblastic cells under conditions that induce differentiation results in a decrease in syndecan-1 levels [120]. Syndecan-2 has been implicated in the mitogenic effect of GM-CSF in osteoblasts [121]. A role of syndecans in presentation of growth factors to their receptors has been suggested as their most prominent function in bone but this falls outside the scope of this chapter. In addition, syndecan-1 could also be an adhesion receptorregulating interaction of osteoblastic cells with the extracellular matrix. The heparin-binding growth-associated molecule (HB-GAM, or pleiotrophin), which is expressed by osteoblasts and osteocytes [118, 122], indeed seems to act in such a way by attracting N-syndecan-expressing osteoblasts, resulting in enhanced bone deposition [118], a process which may be important both during development as well as in response to bone injury.

E. CD44 CD44 has been found in humans and rodents to be expressed in osteoblasts and osteocytes at all stages of maturation; however, there appears to be a clear increase in expression levels with increasing maturation in this cell lineage with low levels expressed in pre-osteoblastic cells and very high levels expressed in osteocytes [123, 124]. Detailed localization studies have shown that CD44 expression in mature osteoblasts is confined to cytoplasmic processes only [125]. By a variety of immunological and molecular techniques only the standard form of CD44, CD44H, was found to be expressed in osteoblasts and osteocytes in rat bone [124, 126]; however, several isoforms of CD44 were described in the human osteoblastic cell line MG63 [127]. Expression of CD44 in cells of the osteoblastic lineage correlates well with the presence of hyaluronate in surrounding tissues [126], suggesting that the receptor may function as a hyaluronate-binding protein. In vitro, osteoblastic cells have indeed been shown to be able to bind to, and degrade, hyaluronate in the transition zone from cartilage to bone in the growth plate by a CD44dependent mechanism [126]. However, a variety of other known ligands for CD44, e.g. type I collagen, fibronectin, laminin, and osteopontin, are also produced by osteoblasts (see Chapters 1–4) and osteocytes [128] and co-localize with the receptor, indicating that a much wider range of functions for this molecule may exist. As described above, crosslinking of CD44 on osteoblasts

leads to up-regulation of ICAM-1 and VCAM-1 [104], which in turn can lead to increased osteoclast formation [106, 107].

F. Selectins There are as yet no published data on expression of selectins in osteoblasts or their precursors. As discussed for osteoclasts below, they may well play a role in extravasation of the mesenchymal precursors of osteoblasts. In the absence of definitive markers for osteoblast precursors, however, such information is as yet unavailable.

V. ADHESION MOLECULES IN OSTEOCLASTS A. Integrins The integrin repertoire expressed in mature osteoclasts is well defined and has been the subject of several reviews (see e.g. reference [129]). Immunocytochemical, biochemical, and functional assays have been used to confirm the expression of three major integrin receptors in human osteoclasts: αvβ3, α2β1, and αvβ1, e.g. [129–131], whereas expression of other subunits has been largely excluded (Tables II and IV). αvβ5 integrin is expressed in osteoclast precursors such as immature bone marrow macrophages, but no longer present in mature osteoclasts [132, 133]. Detailed localization studies have tried to answer the question whether a member of the integrin family is present in, and possibly responsible for, maintenance of the tight sealing zone in resorbing osteoclasts, but the data so far are conflicting. Although some studies have reported the presence of αv and/or β3 subunits at this site [134, 135], others have excluded the presence of αvβ3 (reviewed in references [136, 137]) and α2β1 [138] in the sealing zone, and recent data suggest that this structure is not as impermeable as originally thought [139]. It is clear that none of the integrin receptors are enriched in the clear zone of resorbing osteoclasts and that they are in fact most abundant on the basolateral membrane, where they may act as true receptors coupled to signal transduction pathways [138, 140, 141]. There is also high expression of integrins on the ruffled border membrane in resorbing osteoclasts [138, 140], suggesting that cell–matrix interactions at that site may be important in the formation of the ruffled border by facilitating penetration of the osteoclast membrane deep into the bone and possibly also in the cessation of bone resorption, since elevated levels of calcium in the

140 resorption lacunae may bind to cation-binding sites of αvβ3 integrin and inhibit adhesion (see reference [3]). The role of the vitronectin receptor αvβ3 in osteoclast biology is well established. In vitro mammalian osteoclasts adhere to a wide range of extracellular matrix proteins known to be expressed in bone and bone marrow using αvβ3 (Table IV) and an RGD peptide sequence forms the recognition site for the receptor in all these ligands. Possibly surprisingly, the question of which protein constitutes the natural ligand of osteoclasts in bone remains to be determined. Osteopontin is a candidate [142], since it was found to be enriched underneath the clear zones of resorbing osteoclasts. However, since osteoclasts actively synthesize osteopontin [143], this finding should be interpreted with some reservation. RGD-containing peptides and snake venoms (echistatin and kistrin) have been shown to inhibit osteoclast polarization, activity and formation in vitro [144–147] and they, as well as vitronectin receptor antibodies, also block bone resorption in various rodent models in vivo [148–150]. In vitro, occupation of the vitronectin receptor by antibodies or RGD peptides causes osteoclasts to retract and detach from matrix, similar to the shape changes observed after administration of the potent antiresorptive peptide hormone calcitonin. In mammalian osteoclasts this effect is preceded by a rise in intracellular calcium, localized predominantly to the nucleus [151, 152]. The impressive effects of nonpeptide RGD analogs developed by the pharmaceutical industry [153] on bone resorption (both in in vitro assays and more recently in in vivo animal models (see reference [129]) has led to their introduction into clinical trails for bone diseases associated with excessive bone resorption such as osteoporosis [261a]. Gene-knockout studies where β3 integrins have been deleted in the mouse have underscored the central role of αvβ3 integrin in osteoclast biology [154]. β3-null mice are relatively normal at birth, but develop a progressive, but mild, osteosclerosis by adulthood; this is associated with in vitro evidence of abnormal osteoclast adhesion and bone resorption. They are protected against bone loss after ovariectomy [155], a finding that, together with the reported down-regulation of β3 expression by estradiol in osteoclasts in vitro [156], suggests that the bone-sparing effects of estrogen may be in part through direct effects on osteoclasts. The critical role of β3 integrin in osteoclasts is also underscored by its role in promoting osteoclast survival and in regulating apoptosis [157]. αv knockout mice die perinatally due to vascular defects [158] and osteoclast defects have therefore not been studied extensively. However, mice from both deletions have shown normal skeletal development, contrary to the prediction

M. H. HELFRICH AND M. A. HORTON

that bone modeling would be significantly affected, suggesting that other integrins compensate in the process of bone recognition by osteoclasts. Indeed, this has been shown to be true in an equivalent study in humans using peripheral blood cultures from patients with Glanzmann thrombasthenia carrying an integrin β3-null mutation [159]; in the face of absent αvβ3 there is an up-regulation of α2β1 collagen-binding receptor in these osteoclasts that enables bone resorption to proceed, albeit to a reduced extent. αvβ5-null mice have a relatively normal phenotype [160]. However, αvβ5-null mice generate more osteoclast precursors and, in the absence of estrogen, osteoclast formation and bone resorption in vivo is markedly increased [161], indicating that αvβ5 is an inhibitor of osteoclast formation, contrary to the role of αvβ3 as described above. The function of α2β1 in mammalian osteoclasts is predominantly as a receptor for type I collagen. In contrast to other cell types, adhesion of osteoclasts to collagen appears to be RGD dependent [138, 144], although this could possibly be explained as a dominant-negative effect of RGD occupation of the abundant αvβ3 receptors on osteoclasts. Antibodies to α2 and β1 integrin inhibit resorption by human osteoclasts in vitro, but not to the same extent as antivitronectin receptor antibodies. Avian osteoclasts express abundant β1 integrin, but do not adhere to collagen [162] and probably use β1 integrin predominantly in association with α5 as a fibronectin-binding receptor. The role of αvβ1 on osteoclasts has not been explored in functional assays since no receptor complex-specific antibodies are available at present. This receptor is far less abundant than αvβ3 and α2β1 in osteoclasts [131]. By analogy with other cell types it is likely that αvβ1functions as a receptor for collagen or fibronectin in osteoclasts. Several groups have investigated the downstream signaling pathways associated with osteoclast polarization, which follows integrin-mediated adhesion to matrix and is essential for bone resorption. A number of candidate molecules, such as c-src, integrin-linked kinase (ILK), phosphatidylinositol 3-kinase (PI-3 kinase), FAK and Pyk2, are expressed at high levels in osteoclasts (reviewed in references [163, 164]). c-Src knockout mice have severe osteopetrosis due to lack of ruffled border formation in osteoclasts, clearly implicating this molecule in signaling cascade leading to polarization of osteoclasts [165]. Both PI-3 kinase and ILK have been shown to be associated with the cytoskeleton in osteoclasts when attached to bone matrix and ILK was also found associated with the β3 subunit in bone-adherent osteoclasts [153, 166], suggesting

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that these molecules may also play a role. The importance of FAK in osteoclast function has been queried since no osteoclast abnormalities are seen in the FAK knockout mouse [167]. However, Pyk2, another molecule of the FAK family with high homology to FAK and a similar role as adaptor protein orchestrating cytoskeletal architecture [168], is highly expressed in osteoclasts and seems to fulfill the role that FAK plays in less-motile cell types [169]. Pyk2 has been shown to interact with gelsolin, an actinbinding protein involved in the formation of adhesive structures in osteoclasts and in bone resorption [170, 171]. Following engagement of αvβ3 with ligand, clustering of the receptor occurs and association of its cytoplasmic domain with a complex containing Pyk2 and p130CAS. Autophosphorylation of Pyk2 then leads to recruitment of c-Src and ultimately to cytoskeletal reorganization and bone resorption [163, 172]. Activated c-Src also recruits and phosphorylates c-Cbl, which binds to the SH3 domain of c-Src and negatively regulates Src kinase activity. It is suggested that this may enable transient adhesion and allow for cell migration, a process critically dependent on turnover of adhesive structures [173]. In addition, the Src-dependent phosphorylation of c-Cbl has been shown to be critical for osteoclastic resorption [174]. In c-Srcnull osteoclasts, Pyk-2 has been shown to associate with PLC-γ, a process which can be induced not only by adhesion but also by M-CSF, and can lead to recruitment of integrin to adhesion contacts and cytoskeletal reorganization [175].

showed prominent staining in the clear zone, suggesting a possible role for cadherin in creating a close contact with bone matrix [178].

C. Immunoglobulin Family Members There is good evidence for expression of receptors of the Ig family in osteoclast precursors. Functional evidence implies ICAM-1 and VCAM-1 in osteoclast development in vitro [26, 107, 110] and as discussed in the osteoblast section, osteoclast precursors, i.e. monocytes, not only express ICAM-1 but also its ligand LFA-1, allowing for cell–cell adhesion and possibly cell fusion [107]. Now that osteoclasts can be generated in the absence of osteoblasts in defined cultures with RANKL and M-CSF, it will be possible to study in more detail the expression and functional role of this important class of adhesion molecule during osteoclast differentiation and in mature osteoclasts.

D. Syndecans There is no information about the expression of syndecans in mature osteoclasts or for a role during osteoclast development.

E. CD44 B. Cadherins There is limited information about cadherin expression in osteoclasts. Mbalaviele et al. [176] reported expression of E-cadherin, and absence of P- and N-cadherin, in mature human and mouse osteoclasts. This study also suggests that in the mouse in an in vitro system osteoclast development and in particular osteoclast fusion requires expression of E-cadherin, although it remains to be firmly established whether this is at the level of the osteoclast precursor, or whether E-cadherin expression is required in the accessory cells (osteoblasts and/or stromal cells, which are known to be crucially important in formation of multinucleated osteoclasts and express cadherins). In support of the former a recent study suggested a role for cadherinmediated interactions in early hemopoietic development [177]. In support of a role for cadherins in mature osteoclasts, Ilvesaro et al. [178] demonstrated that a cadherin dimer disrupting HAV peptide inhibited bone resorption. Interestingly pan-cadherin immunostaining in osteoclasts

CD44, for which several possible ligands are expressed in bone (Table IV), is highly expressed in osteoclasts in vivo [123, 125, 130] and generated in vitro [179]. Detailed studies of sites of expression have so far been confined to rodent osteoclasts, where expression is at the basolateral membrane, rather than the clear zone or ruffled border of resorbing osteoclasts [125]. In a functional study aimed at addressing the role of CD44 in mature osteoclasts and during osteoclast development, Kania et al. [180] described that mouse osteoclast formation in vitro was inhibited by CD44 antibodies, whereas the resorptive capacity of mature osteoclasts was not affected. More recently, these in vitro data were confirmed when it was demonstrated that the interaction between CD44 and osteopontin is involved in osteoclast migration, fusion, and bone resorption [181, 182]. However, two independently generated CD44-null mice show no differences in osteoclast formation, fusion, or bone resorption and only small differences in bone mass and anatomy [183], suggesting that

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there is an increased level of complexity and redundancy in the actions of CD44 in vivo. This has also been highlighted by the recent finding that the role of CD44 in osteoclast formation in vitro is dependent on the culture substrate and that it plays no role when cells are grown on bone [184].

F. Selectins So far there are no reports describing expression of selectins on mature osteoclasts. Selectins are likely, however, to have a role during osteoclast development. Osteoclast precursors are present in the circulation and mature osteoclasts can be generated in vitro from mononuclear cells in blood. It remains unclear how osteoclast precursors in vivo reach the sites in bone where they differentiate into fully active osteoclasts, but this must at some point include extravasation, a process that, for all lymphoid cells at least, initially involves selectin-mediated interaction with endothelial cells, followed by integrinmediated processes. Osteoclast–endothelial interactions have been studied [185, 186], but so far the nature of the adhesive molecular interactions between these cells remains unclear. This issue deserves further study, because it might offer therapeutic potential in diseases where osteoclast formation is increased.

VI. ADHESION MOLECULES IN CHONDROCYTES A. Integrins As for cells of the osteoblast lineage, the reported integrin phenotype of chondrocytes is complex (Table III), with additional inconsistency between publications (e.g. see references [187–192] and also reviewed in reference [193]). A synthesis of the literature suggests that human chondrocytes express the β1 integrins α1, α2, α3, α5, and α6, but not α4. β2, β4 and β6 are absent; analysis of β7–9 and CD11 has not been reported. Some studies have shown high expression of αv integrin. As in osteoblasts, this is mainly as αvβ5 and not the αvβ3 dimer; however, a subpopulation of superficial articular chondrocytes was found to be αvβ3 positive [191]. The most abundant chondrocyte integrin appears to be α5β1. These complexities could well relate in part to variation in sampling site, use of fetal versus adult material, species differences, artefacts induced in culture, or influences of disease on phenotypes. Indeed the first possibility is borne out by the study of Salter et al. [190] where the distribution of

integrin clearly differs by site (human articular, epiphyseal, and growth plate chondrocytes were studied) and variation in expression has been observed in cartilage when comparing fetal and adult samples but not during the endocrine-driven pubertal growth [194]. Likewise, changes have been reported in cultured chondrocytes [195–197]. Two relatively newly discovered integrins, α10 [198, 199] and α11 [200, 201], have been shown to be collagen receptors that are expressed in cartilage. α10 has a cellular distribution that differs from α1 and α2 and was dominant during embryonic development [199]. No knockout mice are available to study their cartilage phenotype. Some integrin receptors have been found in the cartilage matrix, in addition to expression in chondrocytes [202]. The role of cell adhesion molecules in cartilage is thus relatively unclear (though the current state of knowledge regarding the function of some cell adhesion molecules is summarized in Table IV). From first principles, these could include roles in cartilage differentiation during fetal development; response to mechanical forces (for example in articular cartilage, menisci); maintenance of tissue architecture and integrity including by matrix synthesis and assembly (e.g. via matrix integrins) and cell adhesion, and regulation of chondrocyte gene expression. Additionally, there is likely to be a role of cell adhesion molecules in responses in cartilage to injury and disease [203, 204]. For example, up-regulation of α2, α4, and β2 subunit expression in osteoarthritic versus normal and cartilage were reported by Ostergaard et al. [205], and altered physiological responses to mechanical stress mediated by α5β1 integrin in osteoarthritic cartilage [206]. The changes in distribution of both integrin and matrix proteins [190] in different zones of cartilage suggest a role in chondrocyte differentiation from mesenchymal precursors [207, 208] and in chondrocyte function. Recently, in vivo studies have underscored the important role of β1 integrin in chondrocytes where Aszódi and colleagues inactivated β1 selectively in chondrocytes [209]. The resulting mice had chondrodysplasia, failed to align chondrocytes into columns in the growth plate. Underlying these abnormalities was an inability of the cells to adhere to fibronectin and to progress normally through mitosis and cytokinesis, due to reduction in cyclin D1 expression. Evidence is accruing for a specialized function for integrins in responses of chondrocytes to mechanical stresses [77, 210–213] and most recently reviewed in reference [77]. Extensive studies with monolayers of isolated articular chondrocytes show that integrin, especially α5β1 interacting with fibronectin, is critical in the hyperpolarization response to mechanical load. Adhesion via integrin leads to formation of focal adhesions in chondrocytes and,

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following mechanical stimulus, phosphorylation of focal adhesion components, including signaling molecules in the MAP kinase pathway, is seen. Synthesis of autocrine/ paracrine factors known to be involved in transducing the biological effect from mechanosensor to effector cells (similar to osteocyte–osteoblast interactions described above) is stimulated by mechanical load on chondrocytes. In this context, IL-4 and substance P appear especially important, in addition to nitric oxide and prostaglandins. Extensive studies have though been performed to address the role of α5β1 in chondrocyte interactions with fibronectin [187, 189, 214–219]. Function-blocking antibodies and RGD peptides have been shown to inhibit cell adhesion to fibronectin and its fragments, thus modifying chondrocyte behavior and cartilage function. Likewise, chondrocyte recognition of collagen (including types I, II, and VI collagen) has been studied in vitro [187, 189, 214, 216, 219–221] and shown to be mediated via β1 integrins: α1β1 for adhesion to type I [196, 222] and type VI [222] and α3β1 [196, 197, 222] or α2β1 [221] for adhesion to type II. There is also increasing evidence for a connection between chondrocyte adhesion to extracellular matrix proteins, especially fibronectin, and chondrocyte–synovial cell interaction [223], chondrocyte cell signaling [224], and chondrocyte survival [225–227]. Regulation of integrin expression and function by cytokines such as IL1, TGFβ, and IGF1 has been linked to the release of matrix metalloproteinases [228] and hence cartilage breakdown [215, 218, 222, 229, 230]. Such events are likely to be involved in the pathogenesis of the cartilage destruction seen in osteoarthritis and rheumatoid arthritis. The key role for downstream signaling following integrin interaction with matrix ligands is underscored by the phenotype of integrin-linked kinase (ILK) knockout mice [231, 232]. ILK acts as a pleiotropic adapter interfacing β integrin cytoplasmic domains into the PKB/Akt signaling pathway; mice with ILK deletion in the chondrocyte lineage have chondrodysplasia, show dwarfism, and isolated chondrocytes have matrix adhesion defects [231, 232]. As discussed above for osteogenic cells, there is an increasing interest in articular cartilage engineering and adhesion processes, especially integrin–matrix interactions, which will need to be considered when designing appropriate scaffolds [233].

B. Cadherins Cadherin-11 is expressed in mesenchymal cells migrating from the neuroectodermal ridge which form presumptive cartilage (and in other morphogenetic events)

in the developing mouse [234]. Likewise, N-cadherin is expressed in prechondrocytic cells in avian and mammalian limb buds, but not mature cartilage [235–237]. In vitro culture of mesenchymal cells suggests a functional role for N-cadherin in early chondrogenesis [235, 236, 238] and its function may be regulated by calciotropic factors such as 1,25D3 and TGFβ [239].

C. Ig Family Members NCAM is similarly distributed to N-cadherin in early cartilage development [239, 240], with N-cadherin expression temporally preceding NCAM. It is therefore suggested that the role of NCAM is to stabilize cell–cell adhesions formed initially by N-cadherin [236]. Primary human articular chondrocytes constitutively express VCAM-1 and ICAM-1 and this expression is increased by cytokines such as IL1β and TNFα and reduced by TGFβ and γ interferon [241–243]. In vitro VCAM-1 and ICAM-1 contribute to T-cell adhesive processes, suggesting that in vivo these molecules may be important players in mediating T cell–chondrocyte interactions at sites of inflammatory joint destruction [242, 244, 245].

D. Syndecans Syndecan-3 expression has been analyzed in developing avian limb buds; there are little data in mammals or for other syndecan forms. Syndecan-3 is highly expressed in proliferating chondrocytes, below the tenascin C-rich layer of articular chondrocytes; decreased levels are found in hypertrophic cartilage [246]. High levels are also found in forming perichondrium (and later in periosteum) in the developing avian limb [247] and it has been suggested that, with tenascin C, syndecan-3 is involved in establishing or maintaining boundaries during skeletogenesis [248]. More recent data support a role for syndecan-3 in the regulation of chondrocyte proliferation [249, 250].

E. CD44 CD44 is expressed by cartilage and has been studied for a variety of sites and species [123, 126, 251], reviewed most recently in [252]. The predominant isoform detected is the standard CD44H variant; epithelial CD44E is not found [253]. There is some evidence from the use of function-blocking antibodies that CD44 is involved in chondrocyte pericellular matrix assembly [254, 255]. The full range of extracellular matrix molecules recognized

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by CD44 in cartilage is unclear, but interaction with hyaluronate clearly occurs [256]. CD44 is up-regulated during cartilage catabolism (for example on IL1α treatment of bovine articular cartilage) and chondrocytes have been shown to actively take up hyaluronate via CD44mediated endocytosis; thus it is reasonable to speculate that this molecule plays a regulatory role in cartilage matrix turnover in health and disease [257, 258]. In support of this, CD44 has been found to be up-regulated in chondrocytes from rheumatoid arthritis cartilage [259]. Recent evidence points to the presence of soluble CD44 in many tissues and especially in serum. Although mainly studied in the context of tumor biology, there is evidence for elevated levels of soluble CD44 in inflammation such as in rheumatoid arthritis [20].

References

F. Selectins There is no information on selectin expression in chondrocytes.

VII. CONCLUSION Bone and cartilage cells express a wide variety of adhesion molecules. Integrin expression has been studied extensively, and there is increasing information on expression of other adhesion molecule family members. There is still limited information on the expression and function of adhesion molecules of all classes during osteoblast and osteoclast development, but, with the ever-increasing knowledge of stage-specific markers, this information should become available. Adhesion molecules fulfill many functions in skeletal cells and are linked to a variety of intracellular signaling pathways, which are increasingly being elucidated. Although no unique osteoblast or osteoclast adhesion molecule has been identified to date, therapeutic strategies based on selectively inhibiting highly expressed receptors, such as αvβ3 in osteoclasts, have proved to be successful in the inhibition of excessive bone resorption. Better knowledge of the expression and roles of adhesion molecules in bone pathology may help in diagnosis, is essential to design appropriate biomaterials for use with skeletal tissue engineering, and may lead to therapeutic strategies for other bone cell types and for specific bone disorders.

Acknowledgments The authors acknowledge their longstanding support from the Arthritis Research Campaign, UK (MHH) and the Wellcome Trust (MAH) for their work in this area.

1. Isacke, C., and Horton, M. A. (2000). “The Adhesion Molecule Facts Book.” 2nd ed. Academic Press, San Diego. 2. Hynes, R. O. (2002). Integrins: bidirectional, allosteric signaling machines. Cell 110, 673–687. 3. Plow, E. F., Haas, T. A., Zhang, L., Loftus, J., and Smith, J. W. (2000). Ligand binding to integrins. J. Biol. Chem. 275, 21785–21788. 4. Nermut, M. V., Green, N. M., Eason, P., Yamada, S. S., and Yamada, K. M. (1988). Electron microscopy and structural model of human fibronectin receptor. EMBO J. 7, 4093–4099. 5. Xiong, J. P., Stehle, T., Diefenbach, B., Zhang, R., Dunker, R., Scott, D. L., Joachimiak, A., Goodman, S. L., and Arnaout, M. A. (2001). Crystal structure of the extracellular segment of integrin alpha Vbeta3. Science 294, 339–345. 6. Xiong, J. P., Stehle, T., Zhang, R., Joachimiak, A., Frech, M., Goodman, S. L., and Arnaout, M. A. (2002). Crystal structure of the extracellular segment of integrin alphaVbeta3 in complex with an Arg-Gly-Asp ligand. Science 296, 151–155. 7. van der Flier, A., and Sonnenberg, A. (2001). Function and interactions of integrins. Cell Tissue Res. 305, 285–298. 8. Potter, E., Bergwitz, C., and Brabant, G. (1999). The cadherin-catenin system: implications for growth and differentiation of endocrine tissues. Endocr. Rev. 20, 207–239. 9. Patel, S. D., Chen, C. P., Bahna, F., Honig, B., and Shapiro, L. (2003). Cadherin-mediated cell-cell adhesion: sticking together as a family. Curr. Opin. Struct. Biol. 13, 690–698. 10. Yagi, T., and Takeichi, M. (2000). Cadherin superfamily genes: functions, genomic organization, and neurologic diversity. Genes Dev. 14, 1169–1180. 11. Westendorf, J. J., Kahler, R. A., and Schroeder, T. M. (2004). Wnt signaling in osteoblasts and bone diseases. Gene 341, 19–39. 12. Nelson, W. J., and Nusse, R. (2004). Convergence of Wnt, beta-catenin, and cadherin pathways. Science 303, 1483–1487. 13. Wheelock, M. J., and Johnson, K. R. (2003). Cadherin-mediated cellular signaling. Curr. Opin. Cell Biol. 15, 509–514. 14. Barclay, A. N., Brown, M. H., Law, S. K. A., McKnight, A. J., Tomlinson, M. G., and van der Merwe, P. A. (1997). The Leukocyte Antigen Factsbook, 2nd ed. Academic Press, San Diego. 15. Brummendorf, T., and Lemmon, V. (2001). Immunoglobulin superfamily receptors: cis-interactions, intracellular adapters and alternative splicing regulate adhesion. Curr. Opin. Cell Biol. 13, 611–618. 16. Hubbard, A. K., and Rothlein, R. (2000). Intercellular adhesion molecule-1 (ICAM-1) expression and cell signaling cascades. Free Radic. Biol. Med. 28, 1379–1386. 17. Elenius, K., and Jalkanen, M. (1994). Function of the syndecans – a family of cell surface proteoglycans. J. Cell Sci. 107, 2975–2982. 18. Russell, R. G., Smith, R., Preston, C., Walton, R. J., and Woods, C. G. (1974). Diphosphonates in Paget’s disease. Lancet 1, 894–898. 19. Weber, G. F., Ashkar, S., Glimcher, M. J., and Cantor, H. (1996). Receptor-ligand interaction between CD44 and osteopontin (Eta-1). Science 271, 509–512. 20. Cichy, J., and Pure, E. (2003). The liberation of CD44. J Cell Biol. 161, 839–843. 21. Frenette, P. S., and Wagner, D. D. (1997). Insights into selectin function from knockout mice. Thromb. Haemost. 78, 60–64. 22. Varki, A. (1997). Selectin ligands: will the real ones please stand up? J. Clin. Invest. 99, 158–162. 23. Ehrhardt, C., Kneuer, C., and Bakowsky, U. (2004). Selectins – an emerging target for drug delivery. Adv. Drug Deliv. Rev. 56, 527–549. 24. Crockett-Torabi, E. (1998). Selectins and mechanisms of signal transduction. J. Leukocyte Biol. 63, 1–14.

Chapter 8 Integrins and Other Adhesion Molecules 25. Grzesik, W. J., and Robey, P. G. (1994). Bone matrix RGD glycoproteins: immunolocalization and interaction with human primary osteoblastic bone cells in vitro. J. Bone Miner. Res. 9, 487–496. 26. Nakayamada, S., Okada, Y., Saito, K., Tamura, M., and Tanaka, Y. (2003). Beta1 integrin/focal adhesion kinase-mediated signaling induces intercellular adhesion molecule 1 and receptor activator of nuclear factor kappaB ligand on osteoblasts and osteoclast maturation. J. Biol. Chem. 278, 45368–45374. 27. Brighton, C. T., and Albelda, S. M. (1992). Identification of integrin cell-substratum adhesion receptors on cultured rat bone cells. J. Orthop. Res. 10, 766–773. 28. Majeska, R. J., Port, M., and Einhorn, T. A. (1993). Attachment to extracellular matrix molecules by cells differing in the expression of osteoblastic traits. J. Bone Miner. Res. 8, 277–289. 29. Ganta, D. R., McCarthy, M. B., and Gronowicz, G. A. (1997). Ascorbic acid alters collagen integrins in bone culture. Endocrinology 138, 3606–3612. 30. Clover, J., and Gowen, M. (1994). Are MG-63 and HOS TE85 human osteosarcoma cell lines representative models of the osteoblastic phenotype? Bone 15, 585–591. 31. Hughes, D. E., Salter, D. M., Dedhar, S., and Simpson, R. (1993). Integrin expression in human bone. J. Bone Miner. Res. 8, 527–533. 32. Clover, J., Dodds, R. A., and Gowen, M. (1992). Integrin subunit expression by human osteoblasts and osteoclasts in situ and in culture. J. Cell Sci. 103, 267–271. 33. Bennett, J. H., Carter, D. H., Alavi, A. L., Beresford, J. N., and Walsh, S. (2001). Patterns of integrin expression in human mandibular explant model of osteoblast differentiation. Arch. Oral Biol. 46, 229–238. 34. Bennett, J. H., Moffatt, S., and Horton, M. (2001). Cell adhesion molecules in human osteoblasts: structure and function. Histol. Histopathol. 16, 603–611. 35. Lai, C. F., and Cheng, S. L. (2005). AlphaVbeta integrins play an essential role in BMP-2 induction of osteoblast differentiation. J Bone Miner. Res. 20, 330–340. 36. Sinha, R. K., and Tuan, R. S. (1996). Regulation of human osteoblast integrin expression by orthopedic implant materials. Bone 18, 451–457. 37. Gronowicz, G., and McCarthy, M. B. (1996). Response of human osteoblasts to implant materials: integrin-mediated adhesion. J. Orthop. Res. 14, 878–887. 38. Helfrich, M. H. (2003). Osteoclast diseases. Microsc. Res. Tech. 61, 514–532. 39. Aarden, E. M., Nijweide, P. J., van der Plas, A., Alblas, M. J., Mackie, E. J., Horton, M. A., and Helfrich, M. H. (1996). Adhesive properties of isolated chick osteocytes in vitro. Bone 18, 305–313. 40. Gohel, A. R., Hand, A. R., and Gronowicz, G. A. (1995). Immunogold localization of β1-integrin in bone: effect of glucocorticoids and insulin-like growth factor I on integrins and osteocyte formation. J. Histochem. Cytochem. 43, 1085–1096. 41. Helfrich, M., van’t Hof, R., and MacPhee, J. (2004). Expression and localization of integrins, NOS and caveolin-1 in osteoblasts and osteocytes. Calcif. Tissue Int. 74, S64–S65. 42. Bruder, S. P., Jaiswal, N., Ricalton, N. S., Mosca, J. D., Kraus, K. H., and Kadiyala, S. (1998). Mesenchymal stem cells in osteobiology and applied bone regeneration. Clin. Orthopaed. Relat. Res. 355(S), 247–256. 43. Stewart, K., Monk, P., Walsh, S., Jefferiss, C. M., Letchford, J., and Beresford, J. N. (2003). STRO-1, HOP-26 (CD63), CD49a and SB-10 (CD166) as markers of primitive human marrow stromal cells and their more differentiated progeny: a comparative investigation in vitro. Cell Tissue Res. 313, 281–290. 44. Kuznetsov, S. A., Mankani, M. H., Gronthos, S., Satomura, K., Bianco, P., and Robey, P. G. (2001). Circulating skeletal stem cells. J Cell Biol. 153, 1133–1140.

145 45. Caplan, A. I., and Bruder, S. P. (2001). Mesenchymal stem cells: building blocks for molecular medicine in the 21st century. Trends Mol. Med. 7, 259–264. 46. Puleo, D. A., and Bizios, R. (1991). RGDS tetrapeptide binds to osteoblasts and inhibits fibronectin-mediated adhesion. Bone 12, 271–276. 47. Gronowicz, G. A., and DeRome, M. E. (1994). Synthetic peptide containing Arg-Gly-Asp inhibits bone formation and resorption in a mineralizing organ culture system of fetal rat parietal bones. J. Bone Miner. Res. 9, 193–201. 48. Doherty, W. J., DeRome, M. E., McCarthy, M. B., and Gronowicz, G. A. (1995). The effect of glucocorticoids on osteoblast function. The effect of corticosterone on osteoblast expression of β1 integrins. J. Bone Joint Surg. Am. 77, 396–404. 49. Cheng, S. L., Lai, C. F., Fausto, A., Chellaiah, M., Feng, X., McHugh, K. P., Teitelbaum, S. L., Civitelli, R., Hruska, K. A., Ross, F. P., and Avioli, L. V. (2000). Regulation of alphaVbeta3 and alphaVbeta5 integrins by dexamethasone in normal human osteoblastic cells. J. Cell Biochem. 77, 265–276. 50. Moursi, A. M., Damsky, C. H., Lull, J., Zimmerman, D., Doty, S. B., Aota, S., and Globus, R. K. (1996). Fibronectin regulates calvarial osteoblast differentiation. J. Cell Sci. 109, 1369–1380. 51. Moursi, A. M., Globus, R. K., and Damsky, C. H. (1997). Interactions between integrin receptors and fibronectin are required for calvarial osteoblast differentiation in vitro. J. Cell Sci. 110, 2187–2196. 52. Jikko, A., Harris, S. E., Chen, D., Mendrick, D. L., and Damsky, C. H. (1999). Collagen integrin receptors regulate early osteoblast differentiation induced by BMP-2. J. Bone Miner. Res. 14, 1075–1083. 53. Mizuno, M., Fujisawa, R., and Kuboki, Y. (2000). Type I collageninduced osteoblastic differentiation of bone-marrow cells mediated by collagen-alpha2beta1 integrin interaction. J. Cell Physiol. 184, 207–213. 54. Xiao, G., Cui, Y., Ducy, P., Karsenty, G., and Franceschi, R. T. (1997). Ascorbic acid-dependent activation of the osteocalcin promoter in MC3T3-E1 preosteoblasts: requirement for collagen matrix synthesis and the presence of an intact OSE2 sequence. Mol. Endocrinol. 11, 1103–1113. 55. Xiao, G., Wang, D., Benson, M. D., Karsenty, G., and Franceschi, R. T. (1998). Role of the alpha2-integrin in osteoblast-specific gene expression and activation of the Osf2 transcription factor. J Biol. Chem. 273, 32988–32994. 56. Xiao, G., Gopalakrishnan, R., Jiang, D., Reith, E., Benson, M. D., and Franceschi, R. T. (2002). Bone morphogenetic proteins, extracellular matrix, and mitogen-activated protein kinase signaling pathways are required for osteoblast-specific gene expression and differentiation in MC3T3-E1 cells. J. Bone Miner. Res. 17, 101–110. 57. Fässler, R., and Meyer, M. (1995). Consequences of lack of beta 1 integrin gene expression in mice. Genes Dev. 9, 1896–1908. 58. Stephens, L. E., Sutherland, A. E., Klimanskaya, I. V., Andrieux, A., Meneses, J., Pedersen, R. A., and Damsky, C. H. (1995). Deletion of beta 1 integrins in mice results in inner cell mass failure and periimplantation lethality. Genes Dev. 9, 1883–1895. 59. Zimmerman, D., Jin, F., Leboy, P., Hardy, S., and Damsky, C. (2000). Impaired bone formation in transgenic mice resulting from altered integrin function in osteoblasts. Dev. Biol. 220, 2–15. 60. Globus, R. K., Amblard, D., Nishimura, Y., Iwaniec, U. T., Kim, J. B., Almeida, E. A., Damsky, C. D., Wronski, T. J., and van der Meulen, M. C. (2004). Skeletal phenotype of growing transgenic mice that express a function-perturbing form of beta1 Integrin in Osteoblasts. Calcif. Tissue Int. 76, 39–49. 61. Chen, J., Diacovo, T. G., Grenache, D. G., Santoro, S. A., and Zutter, M. M. (2002). The alpha(2) integrin subunit-deficient mouse: a multifaceted phenotype including defects of branching morphogenesis and hemostasis. Am. J. Pathol. 161, 337–344.

146 62. Holtkötter, O., Nieswandt, B., Smyth, N., Müller, W., Hafner, M., Schulte, V., Krieg, T., and Eckes, B. (2002). Integrin alpha 2-deficient mice develop normally, are fertile, but display partially defective platelet interaction with collagen. J Biol. Chem. 277, 10789–10794. 63. Ekholm, E., Hankenson, K. D., Uusitalo, H., Hiltunen, A., Gardner, H., Heino, J., and Penttinen, R. (2002). Diminished callus size and cartilage synthesis in alpha 1 beta 1 integrin-deficient mice during bone fracture healing. Am. J. Pathol. 160, 1779–1785. 64. Mercurio, A. M. (2002). Lessons from the alpha2 integrin knockout mouse. Am. J. Pathol. 161, 3–6. 65. Bouvard, D., Brakebusch, C., Gustafsson, E., Aszodi, A., Bengtsson, T., Berna, A., and Fassler, R. (2001). Functional consequences of integrin gene mutations in mice. Circ. Res. 89, 211–223. 66. Sheppard, D. (2000). In vivo functions of integrins: lessons from null mutations in mice. Matrix Biol. 19, 203–209. 67. Wang, N., and Ingber, D. E. (1994). Control of cytoskeletal mechanics by extracellular matrix, cell shape and mechanical tension. Biophys. J. 66, 2181–2189. 68. Ruoslahti, E. (1997). Stretching is good for a cell. Science 276, 1345–1346. 69. Chen, C. S., Mrksich, M., Huang, S., Whitesides, G. M., and Ingber, D. E. (1997). Geometric control of cell life and death. Science 276, 1425–1428. 70. Noble, B. S., and Reeve, J. (2000). Osteocyte function, osteocyte death and bone fracture resistance. Mol. Cell Endocrinol. 159, 7–13. 71. Bonewald, L. F. (2004). Osteocyte biology: its implications for osteoporosis. J. Musculoskelet. Neuronal. Interact. 4, 101–104. 72. Charras, G. T., and Horton, M. A. (2002). Single cell mechanotransduction and its modulation analyzed by atomic force microscope indentation. Biophys. J. 82, 2970–2981. 73. Charras, G. T., and Horton, M. A. (2002). Determination of cellular strains by combined atomic force microscopy and finite element modeling. Biophys. J. 83, 858–879. 74. Burger, E. H., and Klein-Nulend, J. (1999). Mechanotransduction in bone–role of the lacuno-canalicular network. FASEB J. 13 Suppl, S101–S112. 75. Pommerenke, H., Schmidt, C., Durr, F., Nebe, B., Luthen, F., Muller, P., and Rychly, J. (2002). The mode of mechanical integrin stressing controls intracellular signaling in osteoblasts. J. Bone Miner. Res. 17, 603–611. 76. Norvell, S. M., Alvarez, M., Bidwell, J. P., and Pavalko, F. M. (2004). Fluid shear stress induces beta-catenin signaling in osteoblasts. Calcif. Tissue Int. 75, 396–404. 77. Millward-Sadler, S. J., and Salter, D. M. (2004). Integrin-dependent signal cascades in chondrocyte mechanotransduction. Ann. Biomed. Eng. 32, 435–446. 78. Damsky, C. H. (1999). Extracellular matrix-integrin interactions in osteoblast function and tissue remodeling. Bone 25, 95–96. 79. Siebers, M. C., ter Brugge, P. J., Walboomers, X. F., and Jansen, J. A. (2005). Integrins as linker proteins between osteoblasts and bone replacing materials. A critical review. Biomaterials 26, 137–146. 80. Hay, E., Lemonnier, J., Modrowski, D., Lomri, A., Lasmoles, F., and Marie, P. J. (2000). N- and E-cadherin mediate early human calvaria osteoblast differentiation promoted by bone morphogenetic protein-2. J. Cell Physiol. 183, 117–128. 81. Chen, S.-L., Zhang, S.-F., Zhang, L.-M., Warlow, P. M., Davidson, M. K., and Civitelli, R. (1995). Changing pattern of cadherin expression during human osteoblast differentiation. J. Bone Miner. Res. 10, S316. 82. Okazaki, M., Takeshita, S., Kawai, S., Kikuno, R., Tsujimura, A., Kudo, A., and Amann, E. (1994). Molecular cloning and characterization of OB-cadherin, a new member of cadherin family expressed in osteoblasts. J. Biol. Chem. 269, 12092–12098. 83. Prêle, C. M., Horton, M. A., Caterina, P., and Stenbeck, G. (2003). Identification of the molecular mechanisms contributing to polarized trafficking in osteoblasts. Exp. Cell Res. 282, 24–34.

M. H. HELFRICH AND M. A. HORTON 84. Marie, P. J. (2002). Role of N-cadherin in bone formation. J. Cell Physiol. 190, 297–305. 85. Ferrari, S. L., Traianedes, K., Thorne, M., Lafage-Proust, M. H., Genever, P., Cecchini, M. G., Behar, V., Bisello, A., Chorev, M., Rosenblatt, M., and Suva, L. J. (2000). A role for N-cadherin in the development of the differentiated osteoblastic phenotype. J. Bone Miner. Res. 15, 198–208. 86. Kawaguchi, J., Kii, I., Sugiyama, Y., Takeshita, S., and Kudo, A. (2001). The transition of cadherin expression in osteoblast differentiation from mesenchymal cells: consistent expression of cadherin-11 in osteoblast lineage. J. Bone Miner. Res. 16, 260–269. 87. Monaghan, H., Bubb, V. J., Sirimujalin, R., Millward-Sadler, S. J., and Salter, D. M. (2001). Adenomatous polyposis coli (APC), betacatenin, and cadherin are expressed in human bone and cartilage. Histopathology 39, 611–619. 88. Cheng, S. L., Lecanda, F., Davidson, M. K., Warlow, M. P., Zhang, S. F., Zhang, L., Suzuki, S., St. John, T., and Civitelli, R. (1998). Human osteoblasts express a repertoire of cadherins, which are critical for BMP-2 induced osteogenic differentiation. J. Bone Miner. Res. 13, 633–644. 89. Babich, M., and Foti, L. R. P. (1994). E-Cadherins identified in osteoblastic cells effects of parathyroid hormone and extracellular calcium on localization. Life Sci. 54, 201–208. 90. Cheng, S. L., Shin, C. S., Towler, D. A., and Civitelli, R. (2000). A dominant negative cadherin inhibits osteoblast differentiation. J. Bone Miner. Res. 15, 2362–2370. 91. Kawaguchi, J., Azuma, Y., Hoshi, K., Kii, I., Takeshita, S., Ohta, T., Ozawa, H., Takeichi, M., Chisaka, O., and Kudo, A. (2001). Targeted disruption of cadherin-11 leads to a reduction in bone density in calvaria and long bone metaphyses. J. Bone Miner. Res. 16, 1265–1271. 92. Radice, G. L., Rayburn, H., Matsunami, H., Knudsen, K. A., Takeichi, M., and Hynes, R. O. (1997). Developmental defects in mouse embryos lacking N-cadherin. Dev. Biol. 181, 64–78. 93. Castro, C. H., Shin, C. S., Stains, J. P., Cheng, S. L., Sheikh, S., Mbalaviele, G., Szejnfeld, V. L., and Civitelli, R. (2004). Targeted expression of a dominant-negative N-cadherin in vivo delays peak bone mass and increases adipogenesis. J. Cell Sci. 117, 2853–2864. 94. Civitelli, R., Lecanda, F., Jørgensen, N. R., and Steinberg, T. H. (2002). Intercellular junctions and cell-cell communication in bone. In “Principles of Bone Biology”, 2nd ed (J. P. Bilezikan, L. R. Raisz and G. A. Rodan, Eds), Vol I, pp 287–302. Academic Press, San Diego. 95. Di Palma, F., Chamson, A., Lafage-Proust, M. H., Jouffray, P., Sabido, O., Peyroche, S., Vico, L., and Rattner, A. (2004). Physiological strains remodel extracellular matrix and cell-cell adhesion in osteoblastic cells cultured on alumina-coated titanium alloy. Biomaterials 25, 2565–2575. 96. Kashima, T., Kawaguchi, J., Takeshita, S., Kuroda, M., Takanashi, M., Horiuchi, H., Imamura, T., Ishikawa, Y., Ishida, T., Mori, S., Machinami, R., and Kudo, A. (1999). Anomalous cadherin expression in osteosarcoma. Am. J. Pathol. 155, 1549–1555. 97. Iwao, K., Miyoshi, Y., Nawa, G., Yoshikawa, H., Ochi, T., and Nakamura, Y. (1999). Frequent β catenin abnormalities in bone and soft tissue tumours. Jap. J. Cancer Res. 90, 205–209. 98. Shin, C. S., Lecanda, F., Sheikh, S., Weitzmann, L., Cheng, S. L., and Civitelli, R. (2000). Relative abundance of different cadherins defines differentiation of mesenchymal precursors into osteogenic, myogenic, or adipogenic pathways. J. Cell Biochem. 78, 566–577. 99. Lee, Y. S., and Chuong, C. M. (1992). Adhesion molecules in skeletogenesis: I. Transient expression of neural cell adhesion molecules (NCAM) in osteoblasts during endochondral and intramembranous ossification. J. Bone Miner. Res. 7, 1435–1446. 100. Lackie, P. M., Zuber, C., and Roth, J. (1994). Polysialic acid of the neural cell adhesion molecule (N-CAM) is widely expressed during organogenesis in mesodermal and endodermal derivatives. Differentiation 57, 119–131.

Chapter 8 Integrins and Other Adhesion Molecules 101. Chimal-Monroy, J., and Diaz, d. L. (1999). Expression of N-cadherin, N-CAM, fibronectin and tenascin is stimulated by TGF-beta1, beta2, beta3 and beta5 during the formation of precartilage condensations. Int. J. Dev. Biol. 43, 59–67. 102. Reyes-Botella, C., Montes, M. J., Vallecillo-Capilla, M. F., Olivares, E. G., and Ruiz, C. (2002). Antigenic phenotype of cultured human osteoblast-like cells. Cell Physiol. Biochem. 12, 359–364. 103. Tanaka, Y., Maruo, A., Fujii, K., Nomi, M., Nakamura, T., Eto, S., and Minami, Y. (2000). Intercellular adhesion molecule 1 discriminates functionally different populations of human osteoblasts: characteristic involvement of cell cycle regulators. J. Bone Miner. Res. 15, 1912–1923. 104. Fujii, Y., Fujii, K., Nakano, K., and Tanaka, Y. (2003). Crosslinking of CD44 on human osteoblastic cells upregulates ICAM-1 and VCAM-1. FEBS Lett. 539, 45–50. 105. Tanaka, Y., Morimoto, I., Nakano, Y., Okada, Y., Hirota, S., Nomura, S., Nakamura, T., and Eto, S. (1995). Osteoblasts are regulated by the cellular adhesion through ICAM-1 and VCAM-1. J. Bone Miner. Res. 10, 1462–1469. 106. Okada, Y., Morimoto, I., Ura, K., Watanabe, K., Eto, S., Kumegawa, M., Raisz, L., Pilbeam, C., and Tanaka, Y. (2002). Cell-to-Cell adhesion via intercellular adhesion molecule-1 and leukocyte function-associated antigen-1 pathway is involved in 1alpha,25(OH)2D3, PTH and IL-1alpha-induced osteoclast differentiation and bone resorption. Endocr. J. 49, 483–495. 107. Harada, H., Kukita, T., Kukita, A., Iwamoto, Y., and Iijima, T. (1998). Involvement of lymphocyte function-associated antigen-1 and intercellular adhesion molecule-1 in osteoclastogenesis: a possible role in direct interaction between osteoclast precursors. Endocrinology 139, 3967–3975. 108. Kurachi, T., Morita, I., and Murota, S. (1993). Involvement of adhesion molecules LFA-1 and ICAM-1 in osteoclast development. Biochim. Biophys. Acta 1178, 259–266. 109. Barillé, S., Collette, M., Bataille, R., and Amiot, M. (1995). Myeloma cells upregulate interleukin-6 secretion in osteoblastic cells through cell-to-cell contact but downregulate osteocalcin. Blood 86, 3151–3159. 110. Feuerbach, D., and Feyen, J. H. (1997). Expression of the cell-adhesion molecule VCAM-1 by stromal cells is necessary for osteoclastogenesis. FEBS Lett. 402, 21–24. 111. Everts, V., Delaisse, J. M., Korper, W., Jansen, D. C., Tigchelaar-Gutter, W., Saftig, P., and Beertsen, W. (2002). The bone lining cell: its role in cleaning Howship’s lacunae and initiating bone formation. J. Bone Miner. Res. 17, 77–90. 112. Kurokouchi, K., Kambe, F., Yasukawa, K., Izumi, R., Ishiguro, N., Iwata, H., and Seo, H. (1998). TNF-alpha increases expression of IL-6 and ICAM-1 genes through activation of NF-kappaB in osteoblast-like ROS17/2.8 cells. J. Bone Miner. Res. 13, 1290–1299. 113. Lavigne, P., Benderdour, M., Lajeunesse, D., Shi, Q., and Fernandes, J. C. (2004). Expression of ICAM-1 by osteoblasts in healthy individuals and in patients suffering from osteoarthritis and osteoporosis. Bone 35, 463–470. 114. Kavanaugh, A. F., Schulze-Koops, H., Davis, L. S., and Lipsky, P. E. (1997). Repeat treatment of rheumatoid arthritis patients with a murine anti-intercellular adhesion molecule 1 monoclonal antibody. Arthritis Rheum. 40, 849–853. 115. Kavanaugh, A. F., Davis, L. S., Jain, R. I., Nichols, L. A., Norris, S. H., and Lipsky, P. E. (1996). A phase I/II open label study of the safety and efficacy of an anti-ICAM-1 (intercellular adhesion molecule-1; CD54) monoclonal antibody in early rheumatoid arthritis. J. Rheumatol. 23, 1338–1344. 116. Hall, B. K., and Miyake, T. (2000). All for one and one for all: condensations and the initiation of skeletal development. Bioessays 22, 138–147.

147 117. Koyama, E., Shimazu, A., Leatherman, J. L., Golden, E. B., Nah, H. D., and Pacifici, M. (1996). Expression of syndecan-3 and tenascin-C: possible involvement in periosteum development. J. Orthop. Res. 14, 403–412. 118. Imai, S., Kaksonen, M., Raulo, E., Kinnunen, T., Fages, C., Meng, X., Lakso, M., and Rauvala, H. (1998). Osteoblast recruitment and bone formation enhanced by cell matrix-associated heparin-binding growth-associated molecule (HB-GAM). J. Cell Biol. 143, 1113–1128. 119. Worapamorn, W., Tam, S. P., Li, H., Haase, H. R., and Bartold, P. M. (2002). Cytokine regulation of syndecan-1 and -2 gene expression in human periodontal fibroblasts and osteoblasts. J. Periodontal Res. 37, 273–278. 120. Birch, M. A., and Skerry, T. M. (1999). Differential regulation of syndecan expression by osteosarcoma cell lines in response to cytokines but not osteotropic hormones. Bone 24, 571–578. 121. Modrowski, D., Basle, M., Lomri, A., and Marie, P. J. (2000). Syndecan-2 is involved in the mitogenic activity and signaling of granulocyte-macrophage colony-stimulating factor in osteoblasts. J. Biol. Chem. 275, 9178–9185. 122. Liedert, A., Augat, P., Ignatius, A., Hausser, H. J., and Claes, L. (2004). Mechanical regulation of HB-GAM expression in bone cells. Biochem. Biophys. Res. Commun. 319, 951–958. 123. Hughes, D. E., Salter, D. M., and Simpson, R. (1994). CD44 expression in human bone: a novel marker of osteocytic differentiation. J. Bone Miner. Res. 9, 39–44. 124. Jamal, H. H., and Aubin, J. E. (1996). CD44 expression in fetal rat bone: in vivo and in vitro analysis. Exp. Cell Res. 223, 467–477. 125. Nakamura, H., Kenmotsu, S., Sakai, H., and Ozawa, H. (1995). Localization of CD44, the hyaluronate receptor, on the plasma membrane of osteocytes and osteoclasts in rat tibiae. Cell Tissue Res. 280, 225–233. 126. Noonan, K. J., Stevens, J. W., Tammi, R., Tammi, M., Hernandez, J. A., and Midura, R. J. (1996). Spatial distribution of CD44 and hyaluronan in the proximal tibia of the growing rat. J. Orthop. Res. 14, 573–581. 127. Pavasant, P., Shizari, T. M., and Underhill, C. B. (1994). Distribution of hyaluronan in the epiphysial growth plate: turnover by CD44expressing osteoprogenitor cells. J. Cell Sci. 107, 2669–2677. 128. Aarden, E. M., Wassenaar, A. M., Alblas, M. J., and Nijweide, P. J. (1996). Immunocytochemical demonstration of extracellular matrix proteins in isolated osteocytes. Histochem. Cell Biol. 106, 495–501. 129. Horton, M. A., Nesbitt, S. A., Bennett, J. H., and Stenbeck, G. (2002). Integrins and other cell surface attachment molecules of bone cells. In “Principles of Bone Biology”, 2nd ed (J. P. Bilezikan, L. R. Raisz and G. A. Rodan, Eds), Vol I, pp 265–286. Academic Press, San Diego. 130. Athanasou, N. A., and Quinn, J. (1990). Immunophenotypic differences between osteoclasts and macrophage polykaryons: immunohistological distinction and implications for osteoclast ontogeny and function. J. Clin. Pathol. 43, 997–1003. 131. Nesbitt, S., Nesbit, A., Helfrich, M., and Horton, M. (1993). Biochemical characterization of human osteoclast integrins. Osteoclasts express αvβ3, α2β1, and αvβ1 integrins. J. Biol. Chem. 268, 16737–16745. 132. Inoue, M., Teitelbaum, S. L., Hurter, L., Hruska, K., Seftor, E., Hendrix, M., and Ross, F. P. (1995). GM-CSF regulates expression of the functional integrins αvβ3 and αvβ5 in a reciprocal manner during osteoclastogenesis. J. Bone Miner. Res. 10, S163. 133. Inoue, M., Ross, F. P., Erdmann, J. M., Abu-Amer, Y., Wei, S., and Teitelbaum, S. L. (2000). Tumor necrosis factor alpha regulates alpha(v)beta5 integrin expression by osteoclast precursors in vitro and in vivo. Endocrinology 141, 284–290. 134. Nakamura, I., Gailit, J., and Sasaki, T. (1996). Osteoclast integrin αvβ3is present in the clear zone and contributes to cellular polarization. Cell Tissue Res. 286, 507–515.

148 135. Hultenby, K., Reinholt, F. P., and Heinegård, D. (1993). Distribution of integrin subunits on rat metaphyseal osteoclasts and osteoblasts. Eur. J. Cell Biol. 62, 86–93. 136. Väänänen, H. K., and Horton, M. (1995). The osteoclast clear zone is a specialized cell-extracellular matrix adhesion structure. J. Cell Sci. 108, 2729–2732. 137. Stenbeck, G. (2002). Formation and function of the ruffled border in osteoclasts. Semin. Cell Dev. Biol. 13, 285–292. 138. Helfrich, M. H., Nesbitt, S. A., Lakkakorpi, P. T., Barnes, M. J., Bodary, S. C., Shankar, G., Mason, W. T., Mendrick, D. L., Väänänen, H. K., and Horton, M. A. (1996). Beta 1 integrins and osteoclast function: involvement in collagen recognition and bone resorption. Bone 19, 317–328. 139. Stenbeck, G., and Horton, M. A. (2000). A new specialized cell-matrix interaction in actively resorbing osteoclasts. J. Cell Sci. 113 (Pt 9), 1577–1587. 140. Lakkakorpi, P. T., Helfrich, M. H., Horton, M. A., and Väänänen, H. K. (1993). Spatial organization of microfilaments and vitronectin receptor, αvβ3, in osteoclasts. A study using confocal laser scanning microscopy. J. Cell Sci. 104, 663–670. 141. Lakkakorpi, P. T., Horton, M. A., Helfrich, M. H., Karhukorpi, E. K., and Väänänen, H. K. (1991). Vitronectin receptor has a role in bone resorption but does not mediate tight sealing zone attachment of osteoclasts to the bone surface. J. Cell Biol. 115, 1179–1186. 142. Reinholt, F. P., Hultenby, K., Oldberg, Å., and Heinegård, D. (1990). Osteopontin—a possible anchor of osteoclasts to bone. Proc. Natl Acad. Sci. USA 87, 4473–4475. 143. Dodds, R. A., Connor, J. R., James, I. E., Rykaczewski, E. L., Appelbaum, E., Dul, E., and Gowen, M. (1995). Human osteoclasts, not osteoblasts, deposit osteopontin onto resorption surfaces: an in vitro and ex vivo study of remodeling bone. J. Bone Miner. Res. 10, 1666–1680. 144. Nakamura, I., Takahashi, N., Sasaki, T., Jimi, E., Kurokawa, T., and Suda, T. (1996). Chemical and physical properties of the extracellular matrix are required for the actin ring formation in osteoclasts. J. Bone Miner. Res. 11, 1873–1879. 145. Horton, M. A., Taylor, M. L., Arnett, T. R., and Helfrich, M. H. (1991). Arg-Gly-Asp (RGD) peptides and the anti-vitronectin receptor antibody 23C6 inhibit dentine resorption and cell spreading by osteoclasts. Exp. Cell Res. 195, 368–375. 146. van der Pluijm, G., Mouthaan, H., Baas, C., de Groot, H., Papapoulos, S., Löwik, C. (1994). Integrins and osteoclastic resorption in three bone organ cultures: differential sensitivity to synthetic Arg-Gly-Asp peptides during osteoclast formation. J. Bone Miner. Res. 9, 1021–1028. 147. Sato, M., Sardana, M. K., Grasser, W. A., Garsky, V. M., Murray, J. M., and Gould, R. J. (1990). Echistatin is a potent inhibitor of bone resorption in culture. J. Cell Biol. 111, 1713–1723. 148. Fisher, J. E., Caulfield, M. P., Sato, M., Quartuccio, H. A., Gould, R. J., Garsky, V. M., Rodan, G. A., and Rosenblatt, M. (1993). Inhibition of osteoclastic bone resorption in vivo by echistatin, an “arginylglycyl-aspartyl” (RGD)-containing protein. Endocrinology 132, 1411–1413. 149. Crippes, B. A., Engleman, V. W., Settle, S. L., Delarco, J., Ornberg, R. L., Helfrich, M.H., Horton, M. A., and Nickols, G. A. (1996). Antibody to β3 integrin inhibits osteoclast-mediated bone resorption in the thyroparathyroidectomized rat. Endocrinology 137, 918–924. 150. Engleman, V. W., Nickols, G. A., Ross, F. P., Horton, M. A., Griggs, D. W., Settle, S. L., Ruminski, P. G., and Teitelbaum, S. L. (1997). A peptidomimetic antagonist of the αvβ3 integrin inhibits bone resorption in vitro and prevents osteoporosis in vivo. J. Clin. Invest. 99, 2284–2292. 151. Shankar, G., Davison, I., Helfrich, M. H., Mason, W. T., and Horton, M. A. (1993). Integrin receptor-mediated mobilisation of intranuclear calcium in rat osteoclasts. J. Cell Sci. 105, 61–68.

M. H. HELFRICH AND M. A. HORTON 152. Zimolo, Z., Wesolowski, G., Tanaka, H., Hyman, J. L., Hoyer, J. R., and Rodan, G. A. (1994). Soluble αvβ3-integrin ligands raise [Ca2+]i in rat osteoclasts and mouse-derived osteoclast-like cells. Am. J. Physiol. 266, C376–C381. 153. Lakkakorpi, P. T., Wesolowski, G., Zimolo, Z., Rodan, G. A., and Rodan, S. B. (1996). Phosphatidylinositol-3 kinase (PI3-kinase) involvement in osteoclast attachment and spreading on bone. J. Bone Miner. Res. 11, S287. 154. McHugh, K. P., Hodivala-Dilke, K., Zheng, M. H., Namba, N., Lam, J., Novack, D., Feng, X., Ross, F. P., Hynes, R. O., and Teitelbaum, S. L. (2000). Mice lacking beta3 integrins are osteosclerotic because of dysfunctional osteoclasts. J. Clin. Invest. 105, 433–440. 155. Zhao, H., Ross, F. P., Teitelbaum, S. L., and Novack, D. V. (2002). Prevention of ovariectomy-induced bone loss in β3 knockout mice. J. Bone Miner. Res. 17 suppl 1, S133. 156. Saintier, D., Burde, M. A., Rey, J. M., Maudelonde, T., de Vernejoul, M. C., and Cohen-Solal, M. E. (2004). 17beta-estradiol downregulates beta3-integrin expression in differentiating and mature human osteoclasts. J. Cell Physiol. 198, 269–276. 157. Zhao, H., Ross, F. P., and Teitelbaum, S. L. (2004). Unoccupied αvβ3 integrin regulates osteoclast apoptosis by transmitting a positive death signal. Mol. Endocrinol. 19, 771–780. 158. Bader, B. L., Rayburn, H., Crowley, D., and Hynes, R. O. (1998). Extensive vasculogenesis, angiogenesis, and organogenesis precede lethality in mice lacking all alpha v integrins. Cell 95, 507–519. 159. Horton, M. A., Massey, H. M., Rosenberg, N., Nicholls, B., Seligsohn, U., and Flanagan, A. M. (2003). Upregulation of osteoclast alpha2beta1 integrin compensates for lack of alphavbeta3 vitronectin receptor in Iraqi-Jewish-type Glanzmann thrombasthenia. Br. J. Haematol. 122, 950–957. 160. Huang, X., Griffiths, M., Wu, J., Farese, R. V., Jr., and Sheppard, D. (2000). Normal development, wound healing, and adenovirus susceptibility in beta5-deficient mice. Mol. Cell Biol. 20, 755–759. 161. Lane, N. E., Yao, W., Nakamura, M. C., Humphrey, M. B., Kimmel, D., Huang, X., Sheppard, D., Ross, F. P., and Teitelbaum, S. L. (2005). Mice lacking the integrin beta5 subunit have accelerated osteoclast maturation and increased activity in the estrogen-deficient state. J. Bone Miner. Res. 20, 58–66. 162. Ross, F. P., Chappel, J., Alvarez, J. I., Sander, D., Butler, W. T., Farach-Carson, M. C., Mintz, K. A., Robey, P. G., Teitelbaum, S. L., and Cheresh, D. A. (1993). Interactions between the bone matrix proteins osteopontin and bone sialoprotein and the osteoclast integrin αvβ3 potentiate bone resorption. J. Biol. Chem. 268, 9901–9907. 163. Duong, L. T., Lakkakorpi, P., Nakamura, I., and Rodan, G. A. (2000). Integrins and signaling in osteoclast function. Matrix Biol. 19, 97–105. 164. Golden, L. H., and Insogna, K. L. (2004). The expanding role of PI3kinase in bone. Bone 34, 3–12. 165. Soriano, P., Montgomery, C., Geske, R., and Bradley, A. (1991). Targeted disruption of the c-src proto-oncogene leads to osteopetrosis in mice. Cell 64, 693–702. 166. Lakkakorpi, P. T., Dedhar, S., Mostachfi-Vasseghi, H., Rodan, G. A., and Rodan, S. B. (1996). Association of integrin-linked kinase (ILK) with 3-integrin and its activation and translocation to the cytoskeleton upon osteoclast attachment to bone. J. Bone Miner. Res. 11, S287. 167. Ilic, D., Furuta, Y., Kanazawa, S., Takeda, N., Sobue, K., Nakatsuji, N., Nomura, S., Fujimoto, J., Okada, M., and Yamamoto, T. (1995). Reduced cell motility and enhanced focal adhesion contact formation in cells from FAK-deficient mice. Nature 377, 539–544. 168. Xiong, W. C., and Feng, X. (2003). PYK2 and FAK in osteoclasts. Front Biosci. 8, d1219–d1226. 169. Duong, L. T., Lakkakorpi, P. T., Nakamura, I., Machwate, M., Nagy, R. M., and Rodan, G. A. (1998). PYK2 in osteoclasts is an adhesion kinase, localized in the sealing zone, activated by ligation of

Chapter 8 Integrins and Other Adhesion Molecules

170.

171.

172.

173.

174.

175.

176.

177.

178.

179.

180. 181.

182.

183.

184.

185.

alpha(v)beta3 integrin, and phosphorylated by src kinase. J. Clin. Invest. 102, 881–892. Wang, Q., Xie, Y., Du, Q. S., Wu, X. J., Feng, X., Mei, L., McDonald, J. M., and Xiong, W. C. (2003). Regulation of the formation of osteoclastic actin rings by proline-rich tyrosine kinase 2 interacting with gelsolin. J. Cell Biol. 160, 565–575. Chellaiah, M., Kizer, N., Silva, M., Alvarez, U., Kwiatkowski, D., and Hruska, K. A. (2000). Gelsolin deficiency blocks podosome assembly and produces increased bone mass and strength. J. Cell Biol. 148, 665–678. Lakkakorpi, P. T., Bett, A. J., Lipfert, L., Rodan, G. A., and Duong, l. T. (2003). PYK2 autophosphorylation, but not kinase activity, is necessary for adhesion-induced association with c-Src, osteoclast spreading, and bone resorption. J. Biol. Chem. 278, 11502–11512. Sanjay, A., Houghton, A., Neff, L., DiDomenico, E., Bardelay, C., Antoine, E., Levy, J., Gailit, J., Bowtell, D., Horne, W. C., and Baron, R. (2001). Cbl associates with Pyk2 and Src to regulate Src kinase activity, alpha(v)beta(3) integrin-mediated signaling, cell adhesion, and osteoclast motility. J. Cell Biol. 152, 181–195. Miyazaki, T., Sanjay, A., Neff, L., Tanaka, S., Horne, W. C., and Baron, R. (2004). Src kinase activity is essential for osteoclast function. J. Biol. Chem. 279, 17660–17666. Nakamura, I., Lipfert, L., Rodan, G. A., and Le, T. D. (2001). Convergence of alpha(v)beta(3) integrin- and macrophage colony stimulating factor-mediated signals on phospholipase Cgamma in prefusion osteoclasts. J. Cell Biol. 152, 361–373. Mbalaviele, G., Chen, H., Boyce, B. F., Mundy, G., and Yoneda, T. (1995). The role of cadherin in the generation of multinucleated osteoclast from mononuclear precursors in murine marrow. J. Clin. Invest. 95, 2757–2765. Puch, S., Armeanu, S., Kibler, C., Johnson, K. R., Muller, C. A., Wheelock, M. J., and Klein, G. (2001). N-cadherin is developmentally regulated and functionally involved in early hematopoietic cell differentiation. J. Cell Sci. 114, 1567–1577. Ilvesaro, J. M., Lakkakorpi, P. T., and Vänäänen, H. K. (1998). Inhibition of bone resorption in vitro by a peptide containing the cadherin cell adhesion recognition sequence HAV is due to prevention of sealing zone formation. Exp. Cell Res. 242, 75–83. Flanagan, A. M., Sarma, U., Steward, C. G., Vellodi, A., and Horton, M. A. (2000). Study of the nonresorptive phenotype of osteoclast-like cells from patients with malignant osteopetrosis: a new approach to investigating pathogenesis. J. Bone Miner. Res. 15, 352–360. Kania, J. R., Kehat-Stadler, T., and Kupfer, S. R. (1997). CD44 antibodies inhibit osteoclast formation. J. Bone Miner. Res. 12, 1155–1164. Chellaiah, M. A., Kizer, N., Biswas, R., Alvarez, U., StraussSchoenberger, J., Rifas, L., Rittling, S. R., Denhardt, D. T., and Hruska, K. A. (2003). Osteopontin deficiency produces osteoclast dysfunction due to reduced CD44 surface expression. Mol. Biol. Cell 14, 173–189. Suzuki, K., Zhu, B., Rittling, S. R., Denhardt, D. T., Goldberg, H. A., McCulloch, C. A., and Sodek, J. (2002). Colocalization of intracellular osteopontin with CD44 is associated with migration, cell fusion, and resorption in osteoclasts. J. Bone Miner. Res. 17, 1486–1497. Cao, J. J., Singleton, P. A., Majumdar, S., Boudignon, B., Burghardt, A., Kurimoto, P., Wronski, T. J., Bourguignon, L. Y., and Halloran, B. P. (2005). Hyaluronan increases RANKL expression in bone marrow stromal cells through CD44. J. Bone Miner. Res. 20, 30–40. de Vries, T. J., Schoenmaker, T., Beertsen, W., van der, N. R., and Everts, V. (2004). Effect of CD44 deficiency on in vitro and in vivo osteoclast formation. J. Cell Biochem. 94, 954–966. Collin-Osdoby, P., Rothe, L., Anderson, F., Nelson, M., Maloney, W., and Osdoby, P. (2001). Receptor activator of NF-kappa B and osteoprotegerin expression by human microvascular endothelial cells, regulation by inflammatory cytokines, and role in human osteoclastogenesis. J. Biol. Chem. 276, 20659–20672.

149 186. McGowan, N. W., Walker, E. J., MacPherson, H., Ralston, S. H., and Helfrich, M. H. (2001). Cytokine-activated endothelium recruits osteoclast precursors. Endocrinology 142, 1678–1681. 187. Enomoto-Iwamoto, M., Iwamoto, M., Nakashima, K., Mukudai, Y., Boettinger, D., Pacifici, M., Kurisu, K., and Suzuki, F. (1997). Involvement of α5β1 integrin in matrix interactions and proliferation of chondrocytes. J. Bone Miner. Res. 12, 1124–1132. 188. Wang, H., and Kandel, R. A. (2004). Chondrocytes attach to hyaline or calcified cartilage and bone. Osteoarthritis. Cartilage. 12, 56–64. 189. Durr, J., Goodman, S., Potocnik, A., von der Mark, H., and von der Mark, K. (1993). Localization of β1-integrins in human cartilage and their role in chondrocyte adhesion to collagen and fibronectin. Exp. Cell Res. 207, 235–244. 190. Salter, D. M., Godolphin, J. L., and Gourlay, M. S. (1995). Chondrocyte heterogeneity: immunohistologically defined variation of integrin expression at different sites in human fetal knees. J. Histochem. Cytochem. 43, 447–457. 191. Woods, V. L., Jr., Schreck, P. J., Gesink, D. S., Pacheco, H. O., Amiel, D., Akeson, W. H., and Lotz, M. (1994). Integrin expression by human articular chondrocytes. Arthritis. Rheum. 37, 537–544. 192. Salter, D. M., Hughes, D. E., Simpson, R., and Gardner, D. L. (1992). Integrin expression by human articular chondrocytes. Brit. J. Rheumatol. 31, 231–234. 193. Loeser, R. F. (2000). Chondrocyte integrin expression and function. Biorheology 37, 109–116. 194. Hausler, G., Helmreich, M., Marlovits, S., and Egerbacher, M. (2002). Integrins and extracellular matrix proteins in the human childhood and adolescent growth plate. Calcif. Tissue Int. 71, 212–218. 195. Loeser, R. F., Carlson, C. S., and McGee, M. P. (1995). Expression of beta 1 integrins by cultured articular chondrocytes and in osteoarthritic cartilage. Exp. Cell Res. 217, 248–257. 196. Shakibaei, M. (1995). Integrin expression on epiphyseal mouse chondrocytes in monolayer culture. Histol. Histopathol. 10, 339–349. 197. Shakibaei, M., Abou-Rebyeh, H., and Merker, H. J. (1993). Integrins in ageing cartilage tissue in vitro. Histol. Histopathol. 8, 715–723. 198. Camper, L., Hellman, U., and Lundgren-Akerlund, E. (1998). Isolation, cloning, and sequence analysis of the integrin subunit alpha10, a beta1associated collagen binding integrin expressed on chondrocytes. J. Biol. Chem. 273, 20383–20389. 199. Camper, L., Holmvall, K., Wangnerud, C., Aszodi, A., and LundgrenAkerlund, E. (2001). Distribution of the collagen-binding integrin alpha10beta1 during mouse development. Cell Tissue Res. 306, 107–116. 200. Tiger, C. F., Fougerousse, F., Grundstrom, G., Velling, T., and Gullberg, D. (2001). alpha11beta1 integrin is a receptor for interstitial collagens involved in cell migration and collagen reorganization on mesenchymal nonmuscle cells. Dev. Biol. 237, 116–129. 201. Lehnert, K., Ni, J., Leung, E., Gough, S. M., Weaver, A., Yao, W. P., Liu, D., Wang, S. X., Morris, C. M., and Krissansen, G. W. (1999). Cloning, sequence analysis, and chromosomal localization of the novel human integrin alpha11 subunit (ITGA11). Genomics 60, 179–187. 202. Shakibaei, M., and Merker, H. J. (1999). Beta1-integrins in the cartilage matrix. Cell Tissue Res. 296, 565–573. 203. Lapadula, G., Iannone, F., Zuccaro, C., Grattagliano, V., Covelli, M., Patella, V., Lo Bianco, G., and Pipitone, V. (1997). Integrin expression on chondrocytes: correlations with the degree of cartilage damage in human osteoarthritis. Clin. Exp. Rheumatol. 15, 247–254. 204. Forster, C., Kociok, K., Shakibaei, M., Merker, H. J., Vormann, J., Gunther, T., and Stahlmann, R. (1996). Integrins on joint cartilage chondrocytes and alterations by ofloxacin or magnesium deficiency in immature rats. Arch. Toxicol. 70, 261–270. 205. Ostergaard, K., Salter, D. M., Petersen, J., Bendtzen, K., Hvolris, J., and Andersen, C. B. (1998). Expression of alpha and beta subunits of the integrin superfamily in articular cartilage from macroscopically

150

206.

207.

208.

209.

210.

211.

212.

213.

214.

215.

216.

217.

218.

219.

220.

221.

M. H. HELFRICH AND M. A. HORTON normal and osteoarthritic human femoral heads. Ann. Rheum. Dis. 57, 303–308. Millward-Sadler, S. J., Wright, M. O., Lee, H., Caldwell, H., Nuki, G., and Salter, D. M. (2000). Altered electrophysiological responses to mechanical stimulation and abnormal signalling through alpha5beta1 integrin in chondrocytes from osteoarthritic cartilage. Osteoarthritis. Cartilage. 8, 272–278. Hirsch, M. S., and Svoboda, K. K. (1996). Beta 1 integrin antibodies inhibit chondrocyte terminal differentiation in whole sterna. Ann. N. Y. Acad. Sci. 785, 267–270. Tavella, S., Bellesse, G., Castagnola, P., Martin, I., Piccini, D., Doliana, R., Colombatti, A., Cancedda, R., and Tacchetti, C. (1997). Regulated expression of fibronectin, laminin and related integrin receptors during the early chondrocyte differentiation. J. Cell Sci. 110, 2270–3361. Aszódi, A., Hunziker, E. B., Brakebusch, C., and Fassler, R. (2003). Beta1 integrins regulate chondrocyte rotation, G1 progression, and cytokinesis. Genes Dev. 17, 2465–2479. Lucchinetti, E., Bhargava, M. M., and Torzilli, P. A. (2004). The effect of mechanical load on integrin subunits alpha5 and beta1 in chondrocytes from mature and immature cartilage explants. Cell Tissue Res. 315, 385–391. Lee, H. S., Millward-Sadler, S. J., Wright, M. O., Nuki, G., and Salter, D. M. (2000). Integrin and mechanosensitive ion channeldependent tyrosine phosphorylation of focal adhesion proteins and beta-catenin in human articular chondrocytes after mechanical stimulation. J. Bone Miner. Res. 15, 1501–1509. Millward-Sadler, S. J., Wright, M. O., Lee, H., Nishida, K., Caldwell, H., Nuki, G., and Salter, D. M. (1999). Integrin-regulated secretion of interleukin 4: A novel pathway of mechanotransduction in human articular chondrocytes. J. Cell Biol. 145, 183–189. Wright, M. O., Nishida, K., Bavington, C., Godolphin, J. L., Dunne, E., Walmsley, S., Jobanputra, P., Nuki, G., and Salter, D. M. (1997). Hyperpolarisation of cultured human chondrocytes following cyclical pressure-induced strain: evidence of a role for alpha 5 beta 1 integrin as a chondrocyte mechanoreceptor. J. Orthop. Res. 15, 742–747. Enomoto, M., Leboy, P. S., Menko, A. S., and Boettiger, D. (1993). Beta 1 integrins mediate chondrocyte interaction with type I collagen, type II collagen, and fibronectin. Exp. Cell Res. 205, 276–285. Clancy, R. M., Rediske, J., Tang, X., Nijher, N., Frenkel, S., Philips, M., and Abrahamson, S. B. (1997). Outside-in signaling in the chondrocyte. Nitric oxide disrupts fibronectin-induced assembly of a subplasmalemmal actin/rho A/focal adhesion kinase signaling complex. J. Clin. Invest. 100, 1789–1796. Shimizu, M., Minakuchi, K., Kaji, S., and Koga, J. (1997). Chondrocyte migration to fibronectin, type I collagen, and type II collagen. Cell Struct. Funct. 22, 309–315. Homandberg, G. A., and Hui, F. (1994). Arg-Gly-Asp-Ser peptide analogs suppress cartilage chondrolytic activities of integrin-binding and nonbinding fibronectin fragments. Arch. Biochem. Biophys. 310, 40–48. Xie, D., and Homandberg, G. A. (1993). Fibronectin fragments bind to and penetrate cartilage tissue resulting in proteinase expression and cartilage damage. Biochem. Biophys. Acta 1182, 189–196. Loeser, R. F. (1993). Integrin-mediated attachment of articular chondrocytes to extracellular matrix proteins. Arthritis. Rheum. 36, 1103–1110. Camper, L., Heinegård, D., and Lundgren-Akerlund, E. (1997). Integrin α2β1 is a receptor for the cartilage matrix protein chondroadherin. J. Cell Biol. 138, 1159–1167. Holmvall, K., Camper, L., Johansson, S., Kimura, J. H., and Lundgren-Akerlund, E. (1995). Chondrocyte and chondrosarcoma cell integrins with affinity for collagen type II and their response to mechanical stress. Exp. Cell Res. 221, 496–503.

222. Loeser, R. F. (1997). Growth factor regulation of chondrocyte integrins. Differential effects of insulin-like growth factor 1 and transforming growth factor beta on α1β1 integrin expression and chondrocyte adhesion to type VI collagen. Arthritis. Rheum. 40, 270–276. 223. Ramachandrula, A., Tiku, K., and Tiku, M. L. (1992). Tripeptide RGD-dependent adhesion of articular chondrocytes to synovial fibroblasts. J. Cell Sci. 101, 859–871. 224. Loeser, R. F. (2002). Integrins and cell signaling in chondrocytes. Biorheology 39, 119–124. 225. Pulai, J. I., Del, C. M., Jr., and Loeser, R. F. (2002). The alpha5beta1 integrin provides matrix survival signals for normal and osteoarthritic human articular chondrocytes in vitro. Arthritis Rheum. 46, 1528–1535. 226. Cao, L., Lee, V., Adams, M. E., Kiani, C., Zhang, Y., Hu, W., and Yang, B. B. (1999). Beta-1 integrin-collagen interaction reduces chondrocyte apoptosis. Matrix Biol. 18, 343–355. 227. Hirsch, M. S., Lunsford, L. E., Trinkaus-Randall, V., and Svoboda, K. K. (1997). Chondrocyte survival and differentiation in situ are integrin mediated. Dev. Dyn. 210, 249–263. 228. Arner, E. C., and Tortorella, M. D. (1995). Signal transduction through chondrocyte integrin receptors induces matrix metalloproteinase synthesis and synergizes with interleukin-1. Arthritis Rheum. 38, 1304–1314. 229. Yonezawa, I., Kato, K., Yagita, H., Yamauchi, Y., and Okumura, K. (1996). VLA-5-mediated interaction with fibronectin induces cytokine production by human chondrocytes. Biochem. Biophys. Res. Commun. 219, 261–265. 230. Loeser, R. F. (1994). Modulation of integrin-mediated attachment of chondrocytes to extracellular matrix proteins by cations, retinoic acid, and transforming growth factor beta. Exp. Cell Res. 211, 17–23. 231. Grashoff, C., Aszodi, A., Sakai, T., Hunziker, E. B., and Fassler, R. (2003). Integrin-linked kinase regulates chondrocyte shape and proliferation. EMBO Rep. 4, 432–438. 232. Terpstra, L., Prud’homme, J., Arabian, A., Takeda, S., Karsenty, G., Dedhar, S., and St. Arnaud, R. (2003). Reduced chondrocyte proliferation and chondrodysplasia in mice lacking the integrin-linked kinase in chondrocytes. J. Cell Biol. 162, 139–148. 233. van der Kraan, P. M., Buma, P., van Kuppevelt, T., and van den Berg, W. B. (2002). Interaction of chondrocytes, extracellular matrix and growth factors: relevance for articular cartilage tissue engineering. Osteoarthritis. Cartilage. 10, 631–637. 234. Simonneau, L., Kitagawa, M., Suzuki, S., and Thiery, J. P. (1995). Cadherin 11 expression marks the mesenchymal phenotype: towards new functions for cadherins? Cell Adh. Commun. 3, 115–130. 235. Oberlender, S. A., and Tuan, R. S. (1994). Expression and functional involvement of N-cadherin in embryonic limb chondrogenesis. Development 120, 177–187. 236. Tavella, S., Raffo, P., Tacchetti, C., Cancedda, R., and Castagnola, P. (1994). N-CAM and N-cadherin expression during in vitro chondrogenesis. Exp. Cell Res. 215, 354–362. 237. DeLise, A. M., and Tuan, R. S. (2002). Alterations in the spatiotemporal expression pattern and function of N-cadherin inhibit cellular condensation and chondrogenesis of limb mesenchymal cells in vitro. J. Cell Biochem. 87, 342–359. 238. DeLise, A. M., and Tuan, R. S. (2002). Analysis of N-cadherin function in limb mesenchymal chondrogenesis in vitro. Dev. Dyn. 225, 195–204. 239. Tsonis, P. A., Del Rio-Tsonis, K., Millan, J. L., and Wheelock, M. J. (1994). Expression of N-cadherin and alkaline phosphatase in chick limb bud mesenchymal cells: regulation by 1,25-dihydroxyvitamin D3 or TGF-beta 1. Exp. Cell Res. 213, 433–437. 240. Hitselberger Kanitz, M. H., Ng, Y. K., and Iannaccone, P. M. (1993). Distribution of expression of cell adhesion molecules in the mid to late gestational mouse fetus. Pathobiol. 61, 13–18. 241. Bujia, J., Behrends, U., Rotter, N., Pitzke, P., Wilmes, E., and Hammer, C. (1996). Expression of ICAM-1 on intact cartilage and

Chapter 8 Integrins and Other Adhesion Molecules

242.

243.

244.

245.

246.

247.

248.

249.

250.

251.

isolated chondrocytes. In Vitro Cell. Develop. Biol. Animal. 32, 116–122. Kienzle, G., and von Kempis, J. (1998). Vascular cell adhesion molecule 1 (CD106) on primary human articular chondrocytes: functional regulation of expression by cytokines and comparison with intercellular adhesion molecule 1 (CD54) and very late activation antigen 2. Arthritis Rheum. 41, 1296–1305. Davies, M. E., Dingle, J. T., Pigott, R., Power, C., and Sharma, H. (1991). Expression of intercellular adhesion molecule 1 (ICAM-1) on human articular cartilage chondrocytes. Connect Tissue Res. 26, 207–216. Seidel, M. F., Keck, R., and Vetter, H. (1997). ICAM-1/LFA-1 expression in acute osteodestructive joint lesions in collagen-induced arthritis in rats. J. Histochem. Cytochem. 45, 1247–1253. Horner, A., Davies, M. E., and Franz, B. (1995). Chondrocyte-peripheral blood mononuclear cell interactions: the role of ICAM-1. Immunol. 86, 584–590. Shimazu, A., Nah, H. D., Kirsch, T., Koyama, E., Leatherman, J. L., Golden, E. B., Kosher, R. A., and Pacifici, M. (1996). Syndecan-3 and the control of chondrocyte proliferation during endochondral ossification. Exp. Cell Res. 229, 126–136. Seghatoleslami, M. R., and Kosher, R. A. (1996). Inhibition of in vitro limb cartilage differentiation by syndecan-3 antibodies. Dev. Dyn. 207, 114–119. Koyama, E., Leatherman, J. L., Shimazu, A., Nah, H. D., and Pacifici, M. (1995). Syndecan-3, tenascin-C, and the development of cartilaginous skeletal elements and joints in chick limbs. Dev. Dyn. 203, 152–162. Kirsch, T., Koyama, E., Liu, M., Golub, E. E., and Pacifici, M. (2002). Syndecan-3 is a selective regulator of chondrocyte proliferation. J. Biol. Chem. 277, 42171–42177. Pfander, D., Swoboda, B., and Kirsch, T. (2001). Expression of early and late differentiation markers (proliferating cell nuclear antigen, syndecan-3, annexin VI, and alkaline phosphatase) by human osteoarthritic chondrocytes. Am. J. Pathol. 159, 1777–1783. Stevens, J. W., Noonan, K. J., Bosch, P. P., Rapp, T. B., Martin, J. A., Kurriger, G. L., Maynard, J. A., Daniels, K. J., Solursh, M., Tammi, R., Tammi, M., and Midura, R. J. (1996). CD44 in growing normal and neoplastic rat cartilage. Ann. N. Y. Acad. Sci. 785, 333–336.

151 252. Knudson, W., and Loeser, R. F. (2002). CD44 and integrin matrix receptors participate in cartilage homeostasis. Cell Mol. Life Sci. 59, 36–44. 253. Salter, D. M., Godolphin, J. L., Gourlay, M. S., Lawson, M. F., Hughes, D. E., and Dunne, E. (1996). Analysis of human articular chondrocyte CD44 isoform expression and function in health and disease. J. Pathol. 179, 396–402. 254. Knudson, C. B. (1993). Hyaluronan receptor-directed assembly of chondrocyte pericellular matrix. J. Cell Biol. 120, 825–834. 255. Knudson, W., Aguiar, D. J., Hua, Q., and Knudson, C. B. (1996). CD44-anchored hyaluronan-rich pericellular matrices: an ultrastructural and biochemical analysis. Exp. Cell Res. 228, 216–228. 256. Knudson, C. B., and Knudson, W. (2004). Hyaluronan and CD44: modulators of chondrocyte metabolism. Clin. Orthop. S152–S162. 257. Chow, G., Knudson, C. B., Homandberg, G., and Knudson, W. (1995). Increased expression of CD44 in bovine articular chondrocytes by catabolic cellular mediators. J. Biol. Chem. 270, 27734–27741. 258. Hua, Q., Knudson, C. B., and Knudson, W. (1993). Internalization of hyaluronan by chondrocytes occurs via receptor-mediated endocytosis. J. Cell Sci. 106, 365–375. 259. Takagi, T., Okamoto, R., Suzuki, K., Hayashi, T., Sato, M., Sato, M., Kurosaka, N., and Koshino, T. (2001). Up-regulation of CD44 in rheumatoid chondrocytes. Scand. J. Rheumatol. 30, 110–113. 260. Helfrich, M. H., Nesbitt, S. A., Dorey, E. L., and Horton, M. A. (1992). Rat osteoclasts adhere to a wide range of RGD (Arg-Gly-Asp) peptide-containing proteins, including the bone sialoproteins and fibronectin, via a β3 integrin. J. Bone Miner. Res. 7, 335–343. 261. Nakamura, I., Pilkington, M. F., Lakkakorpi, P. T., Lipfert, L., Sims, S. M., Dixon, S. J., Rodan, G. A., and Duong, L. T. (1999). Role of alpha(v)beta(3) integrin in osteoclast migration and formation of the sealing zone. J. Cell Sci. 112, 3985–3993. 261a.Murphy, M. G., Cerchio, K., Stoch, S. A., Gottesdiener, K., Wu, M., Recker, R., and L-000845704 Study Group. (2005). Effect of L-000845704, an alphaVbeta3 integrin antagonist, on markers of bone turnover and bone mineral density in postmenopausal osteoporotic women. J. Clin. Endocrinol. Metab. 90, 2022–2228.

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Chapter 9

Alkaline Phosphatases José Luis Millán

The Burnham Institute, 10901 North Torrey Pines Road, La Jolla, CA 92037-1005, USA

I. Introduction II. Structure and Regulation of the TNAP Gene III. Protein Structure

IV. Function of TNAP V. Clinical Use References

I. INTRODUCTION

completion of the mouse genome project has revealed the presence of a yet uncharacterized mouse AP gene, which this author has named Akp6. The mouse Akp-ps1, Akp5, Akp6, and Akp3 loci, in this spatial order, are clustered in the same region of mouse chromosome 1 [1]. The mRNA molecules for all AP isozymes, human or mouse, are in the order of 2.5 kb in length and encode peptides ranging from 518 to 535 amino acids. While little new data have become available concerning the regulation of the AP genes in the last 5 years, a wealth of new information has been published regarding the protein structure and the function of mammalian APs. This chapter will focus particularly on the structure and function of TNAP, making reference to these new studies and mentioning briefly studies on other AP isozymes when there is paucity of data for TNAP or to exemplify structural features relevant to TNAP activity and function.

The alkaline phosphatase (E.C.3.1.3.1; AP) isozyme expressed in bone is one of several members of the mammalian AP gene family. In humans, APs are encoded by four genes traditionally named after the tissues where they are predominantly expressed, although the gene nomenclature is now gaining wider use (Table I). The tissuenonspecific AP (TNAP) gene (ALPL), located on chromosome 1, is expressed at its highest levels in liver, bone, and kidney (hence the alternative name “L/B/K” AP), in the placenta during the 1st trimester of pregnancy, and at lower levels in numerous other tissues [1]. The other three isozymes, i.e., placental (PLAP), placental-like or germ cell (GCAP), and intestinal AP (IAP), show a much more restricted tissue expression; hence, the general term tissuespecific APs. These isozymes are encoded by three genes (ALPP, ALPP2, and ALPI, respectively) clustered on human chromosome 2, bands q34–q37 [2, 3] and are closely related to one another, showing 90% and 87% identical nucleotide and amino acid sequences, respectively. The orthologous TNAP gene in mice is called Akp2 and is located on mouse chromosome 4 (Table I). The mouse Akp3 gene encodes the IAP isozyme and the mouse Akp5 gene encodes the embryonic AP (EAP) isozyme, that appears to be related to both the human PLAP and GCAP isozymes [4]. In addition to the existence of the inactive pseudoAP gene Akp-ps1 previously described [4], the recent Dynamics of Bone and Cartilage Metabolism

II. STRUCTURE AND REGULATION OF THE TNAP GENE The human TNAP gene (ALPL; NM_000478) is located on the distal short arm of chromosome 1, band 1p36.12 specifically at position chr1: 21581175-21650208, thus occupying a length of 69034 bp. The gene is composed of 13 exons; the first two exons (1a and 1b) are noncoding and are separated from one another and from the exon 153

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JOSÉ LUIS MILLA´ N Table I.

Genes Human genes ALPL

Protein names TNAP

ALPP

PLAP

ALPP2

GCAP

ALPI Mouse genes Akp2 Akp3 Akp5 Akp-ps1 Akp6*

Nomenclature of the Human and Mouse Alkaline Phosphatase Genes and Isozymes

IAP TNAP IAP EAP N/a uprIAP*

Common names

Chromosomal location

Tissue-nonspecific alkaline phosphatase; TNSALP; “liver-bone-kidney type” AP Placental alkaline phosphatase; PLALP; “Regan isozyme” Germ cell alkaline phosphatase, GCALP; “placentallike”; “Nagao isozyme” Intestinal alkaline phosphatase, IALP Tissue-nonspecific alkaline phosphatase; TNSALP; “liver-bone-kidney type” AP Intestinal alkaline phosphatase, IALP Embryonic alkaline phosphatase AP Pseudogene, pseudoAP RIKEN sequence, New AP locus

Accession numbers

chr1:21581174-21650208

NM_001631

chr2:233068964-233073097

NM_001632

Chr2:233097057-233100922

NM_031313

Chr2:233146369-233150245

NM_001631

chr4:136199753-136254338

NM_007431

chr1:87031694-87555136 chr1:86990248-86993641 chr1:86968828-86972484 chr1:87002298-87005230

NM_007432 NM_007433 NG 001240 AK008000

*Names tentatively assigned by the author.

that contains the ATG translation initiation site (exon 2) by relatively large introns, as first demonstrated in the rat [5, 6]. The last exon contains the termination codon and the 3′ untranslated region of the mRNA. The TNAP genes and promoter regions have been cloned and sequenced from human, rat, and mouse [5–10]. Exon 1a in humans and rats shares approximately 66% identical bases [5, 6], while sequence homology cannot be detected between the 1b exon in these two species. The major transcription start site and surrounding sequences of the 5′-most promoter (1a) upstream from the 1a exon of the TNAP gene has been determined for the human [5–7, 11] and mouse [9] TNAP gene. The 1b exon and its promoter (1b) were first recognized in rat [5, 6] and have also been identified and sequenced in the human and mouse genes [8, 9, 12–14]. Importantly, exons 1a and 1b are incorporated into the mRNA in a mutually exclusive fashion that results from the fact that each exon has its own promoter sequence. This results in two types of mRNA, each encoding an identical polypeptide, but having different 5′ untranslated sequences [5, 6, 10, 12, 15]. The promoter usage has primarily been inferred by examination of the structure of TNAP cDNAs isolated from various tissues. In humans and rats, the upstream promoter (1a) is preferentially utilized in osteoblasts and the downstream promoter (1b) preferred in kidney and liver. However, in mice, the upstream promoter is used in all tissues expressing significant amounts of TNAP [9, 10]. Although the 5′ flanking regions of human, rat, and mouse TNAP have been analyzed in transient transfection assays [16, 17], more work is needed to elucidate the mechanism(s) of tissue-specific regulation. Studies using agents that increase TNAP activity have provided evidence for both transcriptional and post-transcriptional regulation

of the enzyme [18, 19]. Retinoic acid, dexamethasone, and granulocyte colony-stimulating factor all promote transcription of TNAP from the upstream 1a promoter and there is evidence that TNAP transcription in nonexpressing tissues of mice is repressed in vivo by methylation at the 1a promoter [11, 20–22]. Regulation of TNAP expression by the forkhead transcription factor (FKHR) via a forkhead response element in the TNAP promoter has been demonstrated [23]. The TNAP promoter appears to be repressed by Smad-interacting protein 1 [24], while Smad3 promotes TNAP expression and mineralization in osteoblastic cell lines [25]. There is also evidence for regulation of TNAP at translational or post-translational stages [10, 19, 26–29]. Since the expression of TNAP in osteoblasts is linked to differentiation, some agents which cause elevated enzyme levels in these cells may do so by increasing expression of osteoblast-specific transcription factors, or relieving osteogenic suppression, rather than acting directly on the TNAP gene. Osteosarcoma/fibroblast cell hybrids regain elevated TNAP when treated with a combination of 1,25-dihydroxyvitamin D3 and TGF, suggesting that this combination diminishes repression of the TNAP gene [30]. Other reagents, such as ascorbic acid (vitamin C) and the bone morphogenetic proteins, may also act indirectly by stimulating osteogenic pathways. For example, ascorbic acid has been shown to increase TNAP mRNA in MC3T3-E1 cells by a slow mechanism requiring increased extracellular collagen synthesis which, in turn, promotes increased osteoblast differentiation [31]. It has been suggested that this mechanism involves activation of a promoter element by the osteogenic transcription factor OSF-2/CBFA-1 [32]. It is clear that detailed analyses of TNAP promoter function using cell transfection and

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trangenic techniques will be necessary to define the regulatory sequences controlling TNAP expression in bone and other tissues.

III. PROTEIN STRUCTURE APs catalyze the hydrolysis of monoesters of phosphoric acid and also catalyze a transphosphorylation reaction in the presence of large concentrations of phosphate acceptors [1]. APs occur widely in nature, and are found in many organisms from bacteria to man. Irrespective of their origin, APs are homodimeric enzymes and each catalytic site contains three metal ions, i.e., two Zn and one Mg, necessary for enzymatic activity. While the main features of the catalytic mechanism are conserved between bacterial and mammalian APs, mammalian APs have higher specific activity and Km values; have a more alkaline pH optimum; are membrane-bound; are inhibited by L-amino acids and peptides through an uncompetitive mechanism; and display lower heat stability. These properties, however, differ noticeably among isozymes. For many years E. coli AP [33] was the only source of structural information on APs, but now the threedimensional structure of the first mammalian AP, i.e., human PLAP (1EW2; http://pdbbeta.rcsb.org/pdb/; [34]) has been solved. The availability of the PLAP structure facilitated modeling the human GCAP, IAP, and TNAP isozymes, revealing that all the novel features discovered in PLAP are conserved in those human isozymes as well [35]. As had been predicted from sequence comparisons, the central core of PLAP, consisting of an extended β-sheet and flanking α-helices, is very similar to that of the E. coli enzyme. The same is true in the immediate vicinity of the three catalytic ions. However, a number of distinctive features, including a different positioning of the aminoterminal segment of the molecule and the expanded top-loop or “crown” domain are now very apparent (Fig. 1). Isozyme-specific properties, such as the characteristic uncompetitive inhibition [36–40], their variable heatstability [41], and their allosteric behavior [42], have been attributed to residues located on the top crown domain unique to mammalian APs (Fig. 1, upper panel). This domain is also responsible for the binding of TNAP to collagen [41, 43], and it may be involved in the function of this isozyme during bone mineralization. Both the amino terminal arm and the crown domain take part in stabilizing the AP dimeric structure. Modeling of the structure of TNAP, GCAP, and IAP allowed the examination of the residues that constitute the monomer–monomer interphase [35]. The overall surface buried at the interface varies between 4134 and 4244 Å2 per monomer, which

155 corresponds to about 25% of the overall protein surface and comprises about 90 residues per monomer. The comparison of the interfaces reveals high conservation in the tissuespecific APs compatible with the fact that PLAP/GCAP or PLAP/IAP heterodimers form readily in nature [44, 45]. In TNAP, however, a number of charged substitutions lead to repulsive forces at the interface, incompatible with the formation of heterodimers between TNAP and any of the tissue-specific APs [35]. An additional noncatalytic metal-binding site, not present in E. coli AP, was revealed in the PLAP structure that appears to be occupied by calcium [34, 46]. The ligands to the calcium site in PLAP include residues W248, F269, E270, D285, and Trp248 that interacts with the calcium through a very stable water molecule. This fourth metal site is conserved in all human and mouse APs and presumably represents a novel feature common to all mammalian APs. However, the structural and functional significance of this new metal site remains to be established. Recent structure-function studies comparing PLAP and the E. coli AP structure have found a conserved function for those residues that stabilize the active site Zn and Mg metal ions [39]. Mutations at residues that coordinate the Zn2 or Mg ions, i.e., D42, S155, E311, D357, and H358, had similar effects in PLAP and E. coli AP, but the environment of the Zn1 ion, i.e., D316, H320, and H432, in PLAP is less affected by substitutions than in E. coli AP. This is due to the identity and location of residue 429 (E429 in PLAP) that covers the entrance to the active site and prevents diffusion of the metal ions despite the introduction of destabilizing mutations at the coordinating residues (Fig. 1, bottom left panel). As will be seen below, residue 429 has crucial significance for the uncompetitive inhibition of other mammalian APs, including TNAP [36–40]. All mammalian APs have five cysteine residues (C101, C121, C183, C467, and C474 in PLAP) per subunit, not homologous to any of the four cysteines in E. coli AP. Disrupting the disulfide bond between C121 and C183 completely prevents the formation of the active enzyme while the C-terminally located C467–C474 bond plays a lesser structural role. The substitution of the free C101 does not significantly affect the properties of the enzyme [39]. Several isozyme-specific inhibitors of APs have been reported. They include L-aminoacids, such as L-phenylalanine, L-tryptophan, L-leucine, L-homoarginine [47, 48], as well as some nonrelated compounds, such as levamisole, the L-stereoisomer of tetramisole [49] and theophylline (a 1,3-dimethyl derivative of xanthine) [50]. The inhibition is of a rare uncompetitive type [51] and, while the biological implications of this inhibition are not known, the inhibitors have proven to be useful in the

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Figure 1 Three-dimensional structure of mammalian APs. Top panel: Modeled structure of human TNAP dimer [35] based on the 3D crystallographic coordinates of PLAP [34]. One subunit is colored magenta and the other white, both in backbone representation. The location of the top “crown domain” is indicated. The site of GP1 anchor attachment, the amino terminal arm and the active site, are only indicated for one subunit but are present in both. The active site metals and phosphate (CPK colors) are indicated. Bottom left panel: Active site environment of PLAP in the vicinity of the Zn1 and Zn2 metal sites and their ligands. Water molecules are shown as red spheres. Green dotted lines denote metal–ligand interactions and hydrogen bonds. Bottom right panel: Top view of the entrance to the PLAP active site, showing the position of the Y367 residue from one subunit (wireframe representation) in the immediate vicinity of the E429 residue of the other subunit (space filling representation). The lower left and right panels were initially published in [39] and are reproduced here with permission from the Journal of Biological Chemistry.

differential determination of AP isozymes in clinical chemistry [52, 53]. The selective uncompetitive inhibition mechanism of PLAP and GCAP by L-Phe and L-Leu is extraordinarily dependent on residue 429. A single E429G substitution in GCAP accounts for the sensitivity of GCAP for L-Leu and the reduced heat stability and decreased Km of this AP isozyme [36–38]. The inhibition occurs through three interaction points of the inhibitor, i.e., the carboxylic group of L-Phe or L-Leu attacks the

active site Arg166 during catalysis, the amino group of the inhibitor interacts with Zn1 in the active site, while the side chain of the inhibitor is stabilized by the loop containing residue 429 (Fig. 1, lower left panel) [36–39]. Another interesting novel structural feature of mammalian APs, also relevant to our understanding of AP inhibition, is Y367. This residue is part of the subunit interface in the PLAP dimer, where it protrudes from one subunit and is positioned within 5.6 Å of the catalytic Zn1

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ion in the active site of the other subunit (Fig. 1, lower right panel). Mutagenesis of this residue leads to PLAP mutants with significantly compromised heat and inhibition properties [39]. Recently, Kozlenkov et al. [40] clarified which amino acid residues are responsible for the marked differences in inhibition selectivity between TNAP and PLAP. In total, six TNAP residues were identified that cluster into two groups; the first group includes residues 433 and 434, and the second group includes residues 108, 109, 120, and 166 (using TNAP numbers). Residue 108 in TNAP largely determines the specificity of inhibition by L-homoarginine, while the conserved Tyr-371 (equivalent to Y367 in PLAP) contributes to a lesser degree to L-homoarginine positioning. In contrast, the binding of levamisole is mostly dependent on His-434 and Tyr-371, but not on residues 108 or 109, while the main determinant of sensitivity to theophylline is His-434 [40]. These studies open the door to drug design efforts aimed at developing more specific inhibitors of TNAP that could be used therapeutically to treat conditions of ectopic calcification where TNAP is known to play a role (see Section IV). As already mentioned, TNAP is also known as liver/ bone/kidney AP since these are tissues known to express high levels of TNAP. These different isoforms differ in their post-translational modifications of the carbohydrate side-chains. The amino acid sequence of the TNAP gene suggests five putative N-glycosylation sites in TNAP, i.e., N123, N213, N254, N286, and N413 [54]. Nosjean et al. [55] have confirmed that both the bone and liver TNAP isoforms are N-glycosylated; however, they did not determine the number of N-glycosylation sites that actually are linked to oligosaccharides. Their data further suggest that the main difference between the bone and liver isoforms is due to a difference in O-glycosylation, i.e., bone-derived TNAP has some O-linked oligosaccharides, whereas liverderived TNAP does not, but that the different immunoreactivity between the bone and liver isoforms is mainly due to differences in N-glycosylation [55]. It has also been reported that bone-derived TNAP contains more fucose [56] and sialic acid residues [55] compared with the liver isoform. The structures of the N-linked and O-linked oligosaccharides and other oligosaccharide differences of bone and liver TNAP have not been reported. Finally, while E. coli AP is located in the periplasmic space of the bacterium, mammalian APs are ectoenzymes bound to the plasma membrane via a glycosylphosphatidylinositol (GPI) anchor [57]. Asp484 was identified as the point of attachment of this anchor in PLAP [58]. Since Asp484 is perfectly conserved in GCAP and IAP we can predict the same attachment site for these human isozymes. Experimental evidence also indicates that the bovine IAPs may be attached via residue 480 [59].

However, the corresponding anchor residue for TNAP has not yet been identified (Fig. 1, upper panel). Pertinent to our discussion of skeletal mineralization, the bone isoform of TNAP functions as an ectoenzyme attached to the osteoblast cell membrane via its GPI anchor [60-63]. In vitro studies have demonstrated that bone AP is released from human osteoblast-line cells in an anchor-intact (insoluble) form attached to matrix vesicles [64, 65], where it participates in the initiation of bone matrix mineralization [66–69]. In vivo, bone AP circulates in an anchordepleted (soluble) homodimeric form, indicating the conversion of the insoluble to the soluble form found in serum by the actions of two endogenous circulating phospholipases, GPI-specific phospholipase C and GPI-specific phospholipase D. Both of these cleavage enzymes are present in humans; but phospholipase C is found in lower levels in serum compared with phospholipase D, which is abundant in the circulation [70–74].

IV. FUNCTION OF TNAP The identification and study of inborn-errors-ofmetabolism often provides invaluable clues as to the in vivo function of a molecule. In humans, only deficiencies in the ALPL gene have been observed. There are no reported cases of deficiencies in the ALPP, ALPP2, or ALPI genes, so the in vivo functions of the PLAP, GCAP, and IAP isozymes remain unclear. However, some clues may be derived from gene-targeting experiments in mice. Functional deletion of the Akp3 gene leads to an accelerated transport of fat through the intestinal epithelium suggesting that IAP may be involved in a rate-limiting step during fat absorption [75]. Deletion of the Akp5 gene leads to a 12-hour delay in pre-implantation development and increased embryo lethality in utero [76]. Whether similar functions could be attributable to the human orthologs remains to be determined. However, missense mutations in the human ALPL gene lead to the inborn-error-of metabolism known as hypophosphatasia [77] and studies of this disease, both in humans and in mouse models, have provided compelling evidence for an important role for TNAP during the development and mineralization of the human skeleton. Hypophosphatasia represents a rare form of rickets and osteomalacia in which neither calcium nor inorganic phosphate levels in serum are subnormal [78]. In fact hypercalcemia and hyperphosphatemia may exist and hypercalciuria is common in infantile hypophosphatasia. The clinical severity in hypophosphatasia patients varies widely. The different syndromes, listed from the most severe to the mildest forms, are: perinatal hypophosphatasia, infantile hypophosphatasia,

158 childhood hypophosphatasia, adult hypophosphatasia, odontohypophosphatasia, and pseudohypophosphatasia [78]. These phenotypes range from complete absence of bone mineralization and stillbirth to spontaneous fractures and loss of decidual teeth in adult life. The severity and expressivity of hypophosphatasia depends on the nature of the ALPL mutation [79–81]. The mapping of hypophosphatasia mutations to specific three-dimensional locations on the TNAP-molecule has provided clues as to the structural significance of these areas for enzyme structure and function [46]. In fact most mutations causing severe hypophosphatasia were mapped to five crucial regions on the TNAP-modeled structure, namely the active site and its vicinity, the active site valley, the homodimer interface, the crown domain, and the metal-binding site (see Section III). Inactivation of the mouse TNAP gene (Akp2) phenocopies the severe “infantile hypophosphatasia” phenotype. The Akp2−/− mice display rickets and osteomalacia at about 6–10 days after birth; they develop extensive epileptic seizures and suffer from apnea, increased apoptosis in the thymus, and abnormal lumbar nerve roots [82] and die around postnatal days 12–15. Biochemically, these animals show increased levels of inorganic pyrophosphate (PPi), pyridoxal-5′-phosphate (PLP, a hydropholic form of vitamin B6), and phosphoethanolamine (PEA) [82–84], molecules that have been proposed as natural substrates of TNAP [77]. Administration of pyridoxal (a hydrophobic form of vitamin B6 that can easily traverse biological membranes), temporarily suppresses the epileptic seizures [83], and reverses the apoptosis in the thymus and the morphology of the lumbar nerve roots [85]. However, the Akp2−/− mice still show a 100% mortality rate before weaning and their demise is often preceded by epileptic seizures. Vitamin B6 is an important coenzyme in several biochemical reactions, including the biosynthesis of the neurotransmitters γ-aminobutyric acid (GABA), dopamine, and serotonin, and is likely to be important for the normal perinatal development of the central nervous system [86]. It is known that vitamin B6 depletion causes epilepsy in rats accompanied by reduced PLP levels in the brain [87]. In general, the epilepsy is believed to involve the GABAergic synapses. Glutamic acid decarboxylase (GAD) catalyzes the synthesis of GABA requiring PLP as a cofactor, and knockout mice lacking one of the GAD isozymes, GAD65, developed spontaneous epilepsy in the adult stage [88]. Waymire et al. [83] showed that the Akp2−/− mice had reduced brain levels of GABA and hypothesized that the observed epileptic seizures result from GAD dysfunction due to shortage of PLP. Vitamin B6 deficiency in suckling rats has been found to result in a decrease in brain sphingolipids since sphingosine synthesis is also a PLP dependent reaction [89].

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We observed abnormal morphology of the lumbar nerve roots in both Akp2−/− and vitamin B6-deficient mice during the suckling stage when myelination is rapidly occurring [82, 85]. It is possible that impaired myelination of nerve cells in the brain and nerve roots descending within the dura may be a contributing factor to the development of epileptic attacks in these mice. While abnormalities in the metabolism of PLP explain many of the abnormalities of infantile hypophosphatasia, it is not the basis of the abnormal mineralization that characterizes this disease. In bone, TNAP is confined to the cell surface of osteoblasts and chondrocytes, including the membranes of their shed matrix vesicles (MVs) [90, 91]. In fact, by an unknown mechanism, MVs are markedly enriched in TNAP compared to both whole cells and the plasma membrane [91]. It has been proposed that the role of TNAP in the bone matrix is to generate the inorganic phosphate needed for hydroxyapatite crystallization [92–94]. However, TNAP has also been hypothesized to hydrolyze the mineralization inhibitor PPi to facilitate mineral precipitation and growth [95, 96]. Electron microscopy observations revealed that TNAP-deficient MVs contain apatite-like mineral crystals, but that extravesicular crystal propagation is retarded [97]. This growth retardation could be due to either the lack of TNAP’s pyrophosphatase function or the lack of inorganic phosphate generation. Recent studies have provided compelling proof that the function of TNAP in bone tissue consists in hydrolyzing PPi to maintain a proper concentration of this mineralization inhibitor to ensure normal bone mineralization [98–100]. The nucleotidetriphosphate pyrophosphohydrolase activity of NPP1 and the transmembrane PPi-channeling protein ANK are responsible for supplying the larger amount of PPi to the extracellular spaces. Mice deficient in NPP1 (Enpp1−/−) or ANK (ank/ank), have decreased levels of extracellular PPi and display soft tissue ossification. Enpp1−/− mice develop features essentially identical to the previously described phenotype of the tiptoe walking (ttw/ttw) mice [101, 102]. These include the development of hyperostosis, starting at approximately 3 weeks of age, in a progressive process that culminates in ossific intervertebral fusion and peripheral joint ankylosis, as well as Achilles tendon calcification. The ank/ank mice have also been characterized as a model of ankylosis [103, 104]. We surmised that affecting the function of either NPP1 or ANK would have beneficial consequences on hypophosphatasia by reducing the amounts of extracellular PPi in the Akp2−/− mice and, conversely, that affecting the function of TNAP should also ameliorate the soft tissue ossification in the Enpp1−/− and ank/ank mutant animals. Indeed, experiments to rescue these abnormalities by combining

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two mutations, i.e., [Akp2−/−; Enpp1−/−] and [Akp2−/−; ank/ank] revealed that both double mutants led to improvement of hypophosphatasia as well as of the ossification of the vertebral apophyses. We found that even carriers (heterozygotes) of Akp2, Enpp1, and Ank mutations were affected to some degree, demonstrating gene dosage effects for all three genes [99, 100]. Interestingly, the expression of yet another mineralization inhibitor, i.e., osteopontin (OPN), was highly elevated in Akp2−/− mice while it was decreased in both the Enpp1−/− and the ank/ank osteoblasts [102]. Importantly, both PPi and OPN levels were corrected in [Akp2−/−; Enpp1−/−] and [Akp2−/−; ank/ank] double knockout mice (Fig. 2). In vitro experiments on wt osteoblasts treated with exogenous PPi revealed an increase in OPN expression and decreased NPP1 and ANK expression [100]. These studies provided evidence for a direct regulation of OPN expression by NPP1 and ANK, mediated by PPi and enabled us to propose a model for the concerted action of the molecules that produce, transport, and degrade PPi. We proposed that under normal conditions the concerted action of TNAP, NPP1, and ANK regulate PPi and OPN levels and thus control hydroxyapatite deposition. Hypophosphatasia in the Akp2−/− mice arises from deficits in TNAP activity, resulting in an increase in PPi levels and a concomitant increase in OPN levels; the combined inhibitory effect of these molecules leads to hypomineralization. In contrast, an NPP1 or ANK deficiency leads to a decrease in the extracellular PPi and OPN

pools, thereby enabling ectopic soft tissue ossification (Fig. 3). The hypomineralization defects in Akp2−/− mice, along with elevated PPi and OPN levels are normalized by ablation of either the NPP1 or ANK gene. Conversely, ablating the function of TNAP causes normalization of the abnormalities in the Enpp1−/− and ank/ank mutant mice via the resulting increase in the concentrations of two inhibitors of mineralization, i.e., PPi and OPN. Thus, NPP1 and ANK represent possible therapeutic targets to correct abnormally elevated levels of PPi, such as those

Figure 3

Figure 2 Correlation between serum PPi and OPN levels. The elevated levels of PPi in Akp2−/− mice cause a secondary increase in OPN, whereas the decreased PPi concentrations in Enpp1−/− and ank/ank mice result in depressed OPN levels. Therefore there is a strong correlation between PPi concentration and serum OPN levels. The double knockout [Akp2−/−; Enpp1−/−] and [Akp2−/−; ank/ank] mice both show normalized PPi levels that also result in correction of OPN levels.

Concerted action of TNAP, NPP1, and ANK in regulating extracellular PPi and OPN levels. Both NPP1 and ANK raise extracellular levels of PPi while TNAP is required for depletion of the PPi pool. Both TNAP and NPP1 are functional in matrix vesicles whereas ANK is not; therefore, NPP1 plays a more crucial role in PPi production than ANK. As a result, the absence of NPP1 in Enpp1−/− mice results in a more severe phenotype than in ank/ank mice. A negative feedback loop exists in which PPi, produced by NPP1 and transported by the channeling action of ANK, inhibits expression of the Enpp1 and Ank genes. In addition, PPi induces expression of the Opn gene and production of OPN, which further inhibits mineralization. In the absence of TNAP, high levels of PPi inhibit mineral deposition directly and also via its induction of OPN expression. The combined action of increased concentrations of PPi and OPN causes hypomineralization. In the absence of NPP1 or ANK, low levels of PPi, in addition to a decrease in OPN levels, leads to hypermineralization. This model clearly points to NPP1 and ANK as therapeutic targets for the treatment of hypophosphatasia. Similarly, targeting TNAP function can be useful in the treatment of hypermineralization abnormalities caused by altered PPi metabolism. This figure was originally published in [100] and is reproduced here with permission from the American Journal of Pathology.

160 found in hypophosphatasia, and TNAP appears as a rational target molecule for the treatment of soft tissue ossification abnormalities due to decreased levels of extracellular PPi, including ankylosis, osteoarthritis, and arterial calcification, since the same molecules implicated in skeletal mineralization appear to also be involved in the pathological calcification of the arteries [105, 106]. It is clear that PLP and PPi are physiological substrates of TNAP and that abnormalities in their metabolism cause the epileptic seizures and hypomineralization, respectively, in hypophosphatasia. Interestingly, several hypophosphatasia mutations were found to affect PLP catalysis to a larger extent than PPi catalysis [81], providing a clue as to one mechanism to explain phenotypic variability in hypophosphatasia, i.e., some individuals suffer from epileptic seizures while displaying mild bone abnormalities, while others have the reverse manifestations [78]. While pyridoxal administration improves the lifespan of the Akp2−/− mice, they still succumb to the disease at about day 25–30. Even [Akp2−/−; Enpp1−/−] mice, that display corrected skeletal abnormalities, die at about this same time point. The ultimate cause of death in the mice has not yet been identified. At the time of death the animals have considerable respiratory distress and episodes of apnea. As discussed above it is possible that anatomic disturbances in the central nervous system resulting from lack of TNAP activity during embryonic development that are not correctable by vitamin B6 supplementation account for the late epilepsy and death [85]. Alternatively, a recent report has pointed to the potential role of TNAP in dephosphorylating AMP to produce adenosine, needed for proper mucociliary clearance and inflammatory responses in the airways that help prevent lung infections [107]. It is possible that abnormalities in adenosine production in the lungs of Akp2−/− mice contribute to their demise and this possibility should be tested experimentally. There is also some anecdotal evidence that PEA may be epileptogenic in hypophosphatasia patients [108]. The especially great elevations of endogenous PEA levels in the Akp2−/− mice could cause this complication as the mice age, which would then not be correctable by vitamin B6 administration. Meanwhile, the reasons for the increased excretion of PEA in hypophosphatasia remain unclear. One possibility is that indeed PEA is a natural substrate of TNAP [109], but the possible metabolic pathway has not been elucidated. An alternative explanation may relate to abnormalities in the function of O-phosphorylethanolamine phosphoylase (PEA-P-lyase) [110, 111]. PEA-P-lyase is an enzyme reported to require PLP as cofactor and, since TNAP is crucial for the normal metabolism of PLP [85], it is possible that it is the insufficient amount of PLP inside the cells that leads to suboptimal activity of

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PEA-P-lyase, which in turn leads to increased excretion of PEA in hypophosphatasia. While one study has addressed this issue [112] and did not find support for this hypothesis, in this author’s opinion there is merit in further exploring this possible metabolic pathway as being affected in hypophosphatasia.

V. CLINICAL USE Measurement of bone-derived TNAP activity in serum can provide an index of the rate of osteoblastic bone formation [50, 113] (see also Chapter 34). It has, however, been difficult to analyze the bone-specific fraction of AP in serum due to the co-existing liver-specific fraction of AP, which has similar physical and biochemical characteristics, e.g., the same primary structure. A vast number of techniques and methods have been developed for separation and quantification of the bone and liver isoforms, such as heat inactivation [114], various chemical inhibitors (e.g., L-homoarginine and levamisole) [49, 115], wheatgerm lectin precipitation [116], electrophoresis [117], isoelectric focusing [118], HPLC [119], and different immunoassays [120]. The development of commercial immunoassays for routine assessment of the bone-specific TNAP isoform [121–125] has indeed increased the availability and use of bone AP, particularly as a biochemical marker of bone turnover for monitoring therapy in osteoporotic patients [126–128]. In healthy adults, the bone and liver TNAP isoforms constitute approximately 95% of the total serum AP activity with similar quantitative levels [129]. At least six different AP isoform peaks can be separated and quantified by weak anion-exchange high-performance liquid chromatography (HPLC) in serum from healthy adults: three bone isoform peaks (B/I, B1, and B2) and three liver isoform peaks (L1, L2, and L3) isoforms [117, 128]. In healthy adults, the three bone isoforms, B/I, B1, and B2, account on average for 4, 16, and 37%, respectively, of the total serum AP activity [128]. In serum, the minor fraction B/I is not a pure bone isoform as it co-elutes with the IAP isozyme and is composed, on average, of 70% bone and 30% IAP. The circulating levels of these bone isoform peaks can vary independently during the pubertal growth spurt and in metabolic bone diseases [130, 131]. Some anatomical differences in the skeletal content of the bonespecific isoforms have also been reported; cortical bone had approximately two-fold higher activity of B1 compared with B2 and, conversely, B2 was approximately two-fold higher in trabecular bone compared with B1. However, the significance of these finding in terms of function and clinical utility remain to be determined.

Chapter 9 Alkaline Phosphatases

Acknowledgments This work was supported by grants AR47908 and DE12889 from the National Institutes of Health, USA.

References 1. Millán, J. L. (2006). Mammalian alkaline phosphatases. From biology to applications in medicine and bio-technology. Wiley-VCH Verlag GmbH & Co., Weinheim, Germany. 2. Smith, M., Weiss, M. J., Griffin, C. A., Murray, J. C., Buetow, K. H., Emanuel, B. S., Henthorn, P. S., and Harris, H. (1988). Regional assignment of the gene for human liver/bone/kidney alkaline phosphatase to chromosome 1p36.1-p34. Genomics 2, 139–143. 3. Martin, D., Tucker, D. F., Gorman, P. A., Sheer, D., Spurr, N. K., and Trowsdale, J. (1987). The human placental alkaline phosphatase gene and related sequences map to chromosome 2 band q37. Ann. Hum. Gen. 51, 145–152. 4. Manes, T., Glade, K., Ziomek, C. A., and Millán, J. L. (1990). Genomic structure and comparison of mouse tissue-specific alkaline phosphatase genes. Genomics 8, 541–554. 5. Toh, Y., Yamamoto, M., Endo, H., Fujiyata, A., Misumi, Y., and Ikehara, Y. (1989a). Sequence divergence of 5’ extremities in rat liver alkaline phosphatase mRNAs. J. Biochem. 105, 61–65. 6. Toh, Y., Yamamoto, M., Endo, H., Misumi, Y., and Ikehara, Y. (1989b). Isolation and characterization of a rat liver alkaline phosphatase gene: a single gene with two promoters. Eur. J. Biochem. 182, 231–237. 7. Weiss, M. J., Ray, K., Henthorn, P. S., Lamb, B., Kadesch, T., and Harris, H. (1988). Structure of the human liver/bone/kidney alkaline phosphatase gene. J. Biol. Chem. 263, 12002–12010. 8. Matsuura, S., Kishi, F., and Kajii, T. (1990). Characterization of a 5’flanking region of the human liver/bone/kidney alkaline phosphatase gene: two kinds of mRNA from a single gene. Biochem. Biophys. Res. Comm. 168, 993–1000. 9. Terao, M., Studer, M., Giannì, M., and Garattini, E. (1990). Isolation and characterization of the mouse liver/bone/kidney-type alkaline phosphatase gene. Biochem. J. 268, 641–648. 10. Studer, M., Terao, M., Giannì, M., and Garattini, E. (1991). Characterization of a second promoter for the mouse liver/bone/kidneytype alkaline phosphatase gene: Cell and tissue specific expression. Biochem. Biophys. Res. Comm. 179, 1352–1360. 11. Zernik, J., Thiede, M. A., Twarog, K., Stover, M. L., Rodan, G. A., Upholt, W. B., and Rowe, D. W. (1990). Cloning and analysis of the 5’ region of the rat bone/liver/kidney/placenta alkaline phosphatase gene – a dual function promoter. Matrix 10, 38–47. 12. Kishi, F., Matsuura, S., and Kaji, T. (1989). Nucleotide sequence of the human liver-type alkaline phosphatase cDNA. Nucl. Acids Res. 17, 2129. 13. Uhler, M.D., and Stofko, R.E. (1990). Induction of alkaline phosphatase in mouse L cells by overexpression of the catalytic subunit of cAMP-dependent protein kinase. J. Biol. Chem. 265, 13181–13189. 14. Terao, M., and Mintz, B. (1987). Cloning and characterization of a cDNA coding for mouose placental alkaline phosphatase. Proc. Natl Acad. Sci. USA 84, 7051–7055. 15. Zernik, J., Kream, B., and Twarog, K. (1991). Tissue-specific and dexamethasone-inducible expression of alkaline phosphatase from alternative promoters of the rat bone/liver/kidney/placenta gene. Biochem. Biophys. Res. Commun. 176, 1149–1156. 16. Kiledjian, M., and Kadesch, T. (1990). Analysis of the human liver/bone/kidney alkaline phosphatase promoter in vivo and in vitro. Nucl. Acids Res. 18, 957–961. 17. Kiledjian, M., and Kadesch, T. (1991). Post-transcriptional regulation of the human liver/bone/kidney alkaline phosphatase gene. J. Biol. Chem. 266, 4207–4213.

161 18. Rodan, G.A., and Rodan, S.B. (1983). Expression of the osteoblastic phenotype. In: Bone and Mineral Research, ed. W.A. Peck, Vol 2, pp. 244–285. Elsevier, Amsterdam. 19. Sato, N., Takahashi, Y., and Asano, S. (1994). Preferential usage of the bone-type leader sequence for the transcripts of liver/bone/kidney-type alkaline phosphatase gene in neutrophilic granulocytes. Blood 83, 1093–1101. 20. Heath, J. K., Suva, L. J., Yoon, K., Kiledjian, M., Martin, T. J., and Rodan, G. A. (1992). Retinoic acid stimulates transcriptional activity from the alkaline phosphatase promoter in the immortalized rat calvarial cell line, RCT-1. Mol. Endo. 6, 636–646. 21. Weiss, M. J., Ray, K., Fallon, M. D., Whyte, M. P., Fedde, K. N., Lafferty, M. A., Mulivor, R., and Harris, H. (1989). Analysis of liver/ bone/kidney alkaline phosphatase mRNA, DNA, and enzymatic activity in cultured skin fibroblasts from 14 unrelated patients with severe hypophosphatasia. Am. J. Hum. Genet. 44, 686–694. 22. Escalante-Alcalde, D., Recillas-Targa, F., Hernández-García, D., Castro-Obregón, S., Terao, M., Garattini, E., and Covarrubias, L. (1996). Retinoic acid and methylation cis-regulatory elements control the mouse tissue non-specific alkaline phosphatase gene expression. Mech. Dev. 57, 21–32. 23. Hatta, M., Daitoku, H., Matsuzaki, H., Deyama, Y., Yoshimura, Y., Suzuki, K., Matsumoto, A., and Fukamizu, A. (2002). Regulation of alkaline phosphatase promoter by forkhead transcription factor FKHR. Int. J. Mol. Med. 9, 147–152. 24. Tylzanowski, P., Verschueren, K., Huylebroeck, D., and Luyten, F. P. (2001). Smad-interacting protein 1 is a repressor of liver/bone/kidney alkaline phosphatase transcription in bone morphogenetic proteininduced osteogenic differentiation of C2C12 cells. J. Biol. Chem. 276, 40001–40007. 25. Sowa, H., Kaji, H., Yamaguchi, T., Sugimoto, T., and Chihara, K. (2002). Smad3 promotes alkaline phosphatase activity and mineralization of osteoblastic MC3T3-E1 cells. J. Bone Min. Res. 17, 1190–1199. 26. Leboy, P. S., Beresford, J. N., Devlin, C., and Owen, M. E. (1991). Dexamethasone induction of osteoblast mRNAs in rat marrow stromal cell cultures. J. Cell. Physiol. 146, 370–378. 27. Kyeyune-Nyombi, E., Nicolas, V., Strong, D. D., and Farley, J. (1995). Paradoxical effects of phosphate to directly regulate the level of skeletal alkaline phosphatase activity in human osteosarcoma (SaOS-2) cells and inversely regulate the level of skeletal alkaline phosphatase mRNA. Calcif. Tissue Int. 56, 154–159. 28. Kobayashi, T., and Robinson, J. M. (1991). A novel intracellular compartment with unusual secretory properties in human neutrophils. J. Cell. Biol. 113, 743–756. 29. Cain, T. J., Liu, Y., Takizawa, T., and Robinson, J. M. (1995). Solubilization of glycosyl-phosphatidylinositol-anchored proteins in quiescent and stimulated neutrophils. Biochim. Biophys. Acta 1235, 69–78. 30. Johnson-Pais, T. L., and Leach, R. J. (1996). 1,25-dihydroxyvitamin D3 and transforming growth factor-β act synergistically to override extinction of liver/bone/kidney alkaline phosphatase in osteosarcoma hybrid cells. Exp. Cell Res. 226, 67–74. 31. Franceschi, R. T., Iyer, B. S., and Cui, Y. (1994). Effects of ascorbic acid on collagen matrix formation and osteoblast differentiation in murine MC3T3-E1 cells. J. Bone Min. Res. 9, 843–854. 32. Xiao, G., Cui, Y., Ducy, P., Karsenty, G., and Franceschi, R. T. (1997). Ascorbic-acid-dependent activation of the osteocalcin promoter in MC3T3-E1 preosteoblasts: Requirement for collagen matrix synthesis and the presence of an intact OSE2 sequence. Mol. Endo. 11, 1103–1113. 33. Kim, E. E., and Wyckoff, H. W. (1991). Reaction mechanism of alkaline phosphatase based on crystal structures: two metal ions catalysis. J. Mol. Biol. 218, 449–464. 34. Le Du, M. H., Stigbrand, T., Taussig, M. J., Menez, A., and Stura, E. A. (2001). Crystal structure of alkaline phosphatase from human placenta

162

35.

36.

37.

38.

39.

40.

41.

42. 43.

44.

45.

46.

47.

48.

49.

50. 51. 52. 53.

54.

at 1.8 A resolution. Implication for a substrate specificity. J. Biol. Chem. 276, 9158–9165. Le Du, M-H., and Millán, J. L. (2002). Structural evidence of functional divergence in human alkaline phosphatases. J. Biol. Chem. 277, 49808–49814. Hummer, C., and Millán, J. L. (1991). Gly429 is the major determinant of uncompetitive inhibition of human germ cell alkaline phosphatase by L-leucine. Biochem. J. 274, 91–95. Hoylaerts, M. F., and Millán, J. L. (1991). Site-directed mutagenesis and epitope-mapped monoclonal antibodies define a catallytically important conformational difference between human placental and germ cell alkaline phosphatase. Eur. J. Biochem. 202, 605–616. Hoylaerts, M. F., Manes, T., and Millan, J. L. (1992). Molecular mechanism of uncompetitive inhibition of human placental and germ cell alkaline phosphatase. Biochem. J. 286, 23–30. Kozlenkov, A., Manes, T., Hoylaerts, M. F., and Millán, J. L. (2002). Function assignment to conserved residues in mammalian alkaline phosphatases. J. Biol. Chem. 277, 22992–22999. Kozlenkov, A., Hoylaerts, M. F., Ny, T., Le Du, M-H., and Millán, J. L. (2004). Residues determining the binding specificity of uncompetitive inhibitors to tissue-nonspecific alkaline phosphatase. J. Bone Min. Res. 19, 1862–1872. Bossi, M., Hoylaerts, M. F., and Millán, J. L. (1993). Modifications in a flexible surface loop modulate the isozyme-specific properties of mammalian alkaline phosphatases. J. Biol. Chem. 268, 25409–25416. Hoylaerts, M. F., Manes, T., and Millán, J. L. (1997). Mammalian alkaline phosphatases are allosteric enzymes. J. Biol. Chem. 272, 22781–22787. Wu, L. N. Y., Genge, B. R., Lloyd, G. C., and Wuthier, R. E. (1991). Collagen binding proteins in collagenase-released matrix vesicles from cartilage. J. Biol. Chem. 266, 1195–1203. Behrens, C. M., Enns, C. A., and Sussman, H. H. (1983). Characterization of human foetal intestinal alkaline phosphatase. Comparison with the isoenzymes from the adult intestine and human tumour cell lines. Biochem. J. 211, 553–558. Imanishi, H., Hada, T., Muratani, K., Hirano, K., and Higashino, K. (1990). Expression of a hybrid form of alkaline phosphatase isoenzyme in a newly established cell line (HuG-1) from a gastric cancer patient. Cancer Res. 50, 3408–3412. Mornet, E., Stura, E., Lia-Baldini, A. S., Stigbrand, T., Menez, A., and Le Du, M. H. (2001). Structural evidence for a functional role of human tissue-nonspecific alkaline phosphatase in bone mineralization. J. Biol. Chem. 276, 31171–31178. Fishman, W. H., and Sie, H.-G. (1971). Organ-specific inhibition of human alkaline phosphatase isozymes of liver, bone, intestine, and placenta: L-phenylalanine, L-tryptophan and L-homoarginine. Enzymologia 41, 140–167. Doellgast, G. J., and Fishman, W. H. (1976). L-Leucine, a specific inhibitor of a rare human placental alkaline phosphatase phenotype. Nature 259, 49–51. Van Belle, H. (1976). Alkaline phosphatase. I. Kinetics and inhibition by levamisole of purified isoenzymes from humans. Clin. Chem. 22, 972–976. Farley, J. R., and Baylink, D. J. (1986). Skeletal alkaline phosphatase activity as a bone formation index in vitro. Metabolism 35, 563–571. Cornish-Bowden, A. (1995). Fundamentals of Enzyme Kinetics, revised Edition, Chap. 5, p 100. Portland Press Ltd, London. Fishman, W. H. (1974). Perspectives on alkaline phosphatase isoenzymes. Am. J. Med. 56, 617–650. Mulivor, R. A., Plotkin, L. I., and Harris, H. (1978). Differential inhibition of the products of the human alkaline phosphatase loci. Ann. Hum. Genet. 42, 1–13. Weiss, M. J., Henthorn, P. S., Lafferty, M. A., Slaughter, C., Raducha, M., and Harris, H. (1986). Isolation and characterization of a cDNA encoding a human liver/bone/kidney-type alkaline phosphatase. Proc. Natl Acad. Sci. USA 83, 7182–7186.

JOSÉ LUIS MILLA´ N 55. Nosjean, O., Koyama, I., Goseki, M., Roux, B., and Komoda, T. (1997). Human tissue non-specific alkaline phosphatases: sugar-moietyinduced enzymic and antigenic modulations and genetic aspects. Biochem. J. 321, 297–303. 56. Kuwana, T., Sugita, O., and Yakata, M. (1991). Sugar chain heterogeneity of bone and liver alkaline phosphatase in serum. Enzyme 45, 63–66. 57. Low, M. G., and Saltiel, A. R. (1988). Structural and functional roles of glycosylphosphatidylinositol in membranes. Science 239, 268–275. 58. Micanovic, R., Bailey, C. A., Brink, L., Gerber, L., Pan, Y. C. E., Hulmes, J. D., and Udenfriend, S. (1988). Aspartic acid-484 of nascent placental alkaline phosphatase condenses with a phosphatidylinositol glycan to become the carboxyl terminus of the mature enzyme. Proc. Natl Acad. Sci. USA 85, 1398–1402. 59. Manes, T., Hoylaerts, M. F., Muller, R., Lottspeich, F., Holke, W., and Millán, J. L. (1998). Genetic complexity, structure, and characterization of highly active bovine intestinal alkaline phosphatases. J. Biol. Chem. 273, 23353–23360. 60. Hawrylak, K., and Stinson, R. A. (1988). The solubilization of tetrameric alkaline phosphatase from human liver and its conversion into various forms by phosphatidylinositol phospholipase C or proteolysis. J. Biol. Chem. 263, 14368–14373. 61. Hamilton, B. A., McPhee, J. L., Hawrylak, K., and Stinson, R.A. (1990). Alkaline phosphatase releasing activity in human tissues. Clin. Chim. Acta. 186, 249–254. 62. Fedde, K. N., Lane, C. C., and Whyte, M. P. (1988). Alkaline phosphatase is an ectoenzyme that acts on micromolar concentrations of natural substrates at physiologic pH in human osteosarcoma cells. Arch. Biochem. Biophys. 264, 400–409. 63. Hooper, N. M. (1997). Glycosyl-phosphatidylinositol anchored membrane enzymes. Clin. Chim. Acta. 266, 3–12. 64. Fedde, K. N. (1992). Human osteosarcoma cells spontaneously release matrix-vesicle-like structures with the capacity to mineralize. Bone Min. 17, 145–151. 65. Anh, D. J., Dimai, H. P., Hall, S. L., and Farley, J. R. (1998). Skeletal alkaline phosphatase activity is primarily released from human osteoblasts in an insoluble form, and the net release is inhibited by calcium and skeletal growth factors. Calcif. Tissue Int. 62, 332–340. 66. Tenenbaum, H. C. (1987). Levamisole and inorganic pyrophosphate inhibit β-glycerophosphate induced mineralization of bone formed in vitro. Bone Min. 3, 13–26. 67. Bellows, C. G., Aubin, J. E., and Heersche, J. N. M. (1991). Initiation and progression of mineralization of bone nodules formed in vitro: the role of alkaline phosphatase and organic phosphate. Bone Min. 14, 27–40. 68. Barling, P. M., Gupta, D. K., and Lim, C. E. L. (1999). Involvement of phosphodiesterase I in mineralization: histochemical studies using antler from red deer (Cervus elaphus) as a model. Calcif. Tissue Int. 65, 384–389. 69. Wennberg, C., Hessle, L., Lundberg, P., Mauro, S., Narisawa, S., Lerner, U. H., and Millán, J. L. (2000). Functional characterization of osteoblasts and osteoclasts from alkaline phosphatase knockout mice. J. Bone Min. Res. 15, 1879–1888. 70. Low, M. G., Carroll, R. C., and Cox, A. C. (1986). Characterization of multiple forms of phosphoinositide-specific phospholipase C purified from human platelets. Biochem. J. 237, 139–145. 71. Davitz, M. A., Hereld, D., Shak, S., Krakow, J., Englund, P. T., and Nussenzweig, V. (1987). A glycan-phosphatidylinositol-specific phospholipase D in human serum. Science 238, 81–84. 72. Low, M. G., and Prasad, A. R. S. (1988). A phospholipase D specific for the phosphatidylinositol anchor of cell-surface proteins is abundant in plasma. Proc. Natl Acad. Sci. USA 85, 980–984. 73. Miller, J. L. (1998). Release and extracellular transit of glycosylphosphatidylinositol proteins. J. Lab. Clin. Med. 131, 115–123.

Chapter 9 Alkaline Phosphatases 74. Anh, D. J., Eden, A., and Farley, J. R. (2001). Quantitation of soluble and skeletal alkaline phosphatase, and insoluble alkaline phosphatase anchor-hydrolase activities in human serum. Clin. Chim. Acta. 311, 137–148. 75. Narisawa, S., Huang, L., Iwasaki, A., Hasegawa, H., Alpers, D. H., and Millán, J. L. (2003). Accelerated fat absorption in intestinal alkaline phosphatase knockout mice. Mol. Cell. Biol. 23, 7525–7530. 76. Dehghani, H., Narisawa, S., Millán, J. L., and Hahnel, A. C. (2000). Effects of disruption of the embryonic alkaline phosphatase gene on the preimplantation development of the mouse. Dev. Dyn. 217, 440–448. 77. Whyte, M. P. (1994). Hypophosphatasia and the role of alkaline phosphatase in skeletal mineralization. Endo. Rev. 15, 439–461. 78. Whyte, M. P. (1995). Hypophosphatasia. In: C. R. Scriver, A. L. Beaudet, W. S. Sly, D.Valle (eds), The Metabolic and Molecular Bases of Inherited Disease, 7th ed. McGraw-Hill, New York, pp. 4095–4112. 79. Henthorn, P. S., and Whyte, M. P. (1992). Missense mutations of the tissue-nonspecific alkaline phosphatase gene in hypophosphatasia. Clin. Chem. 38, 2501–2505. 80. Zurutuza, L., Muller, F., Gibrat, J. F., Taillandier, A., Simon-Buoy, B., Serre, J. L., and Mornet, E. (1999). Correlations of genotype and phenotype in hypophosphatasia. Hum. Mol. Genet. 8, 1039–1046. 81. Di Mauro, S., Manes, T., Hessle, L., Kozlenkov, A., Pizauro, J. M., Hoylaerts, M. F., and Millán, J. L. (2002). Kinetic characterization of hypophosphatasia mutations with physiological substrates. J. Bone Min. Res. 17, 1383–1391. 82. Narisawa, S., Fröhlander, N., and Millán, J. L. (1997). Inactivation of two mouse alkaline phosphatase genes and establishment of a model of infantile hypophosphatasia. Dev. Dyn. 208, 432–446. 83. Waymire, K. G., Mahuren, J. D., Jaje, J. M., Guilarte, T. R., Coburn, S. P., and MacGregor, G. R. (1995). Mice lacking tissue non-specific alkaline phosphatase die from seizures due to defective metabolism of vitamin B-6. Nat. Genet. 11, 45–51. 84. Fedde, K. N., Blair, L., Silverstein, J., Coburn, S. P., Ryan, L. M., Weinstein, R. S., Waymire, K., Narisawa, S., Millán, J. L., MacGregor, G. R., and Whyte, M. P. (1999). Alkaline phosphatase knock-out mice recapitulate the metabolic and skeletal defects of infantile hypophosphatasia. J. Bone Min. Res. 14, 2015–2026. 85. Narisawa, S., Wennberg, C., and Millán, J. L. (2001). Abnormal vitamin B6 metabolism causes multiple abnormalities, but not impaired bone mineralization in alkaline phosphatase knockout mice. J. Pathol. 193, 125–133. 86. Spector, R. (1978). Vitamin B6 transport in the central nervous system: in vivo studies. J. Neurochem. 30, 881–887. 87. Stephens, M. C., Havlicek, V., and Dakshinamurti, K. (1971). Pyridoxine deficiency and development of the central nervous system in the rat. J. Neurochem. 18, 2407–2416. 88. Kash, S. F., Johnson, R. S., Tecott, L. H., Noebels, J. L., Mayfield, R. D., Hanahan, D., and Baekkeskov, S. (1997). Epilepsy in mice deficient in the 65-kDa isoform of glutamic acid decarboxylase. Proc. Natl Acad. Sci. USA 94, 14060–14065. 89. Stephens, M. C., and Dakshinamurti, K. (1975). Brain lipids in pyridoxine-deficient young rats. Neurobiology 5, 262–269. 90. Bernard, G. W. (1978). Ultrastructural localization of alkaline phosphatase in initial membranous osteogenesis. Clin. Orthop. 135, 218–225. 91. Morris, D. C., Masuhara, K., Takaoka, K., Ono, K., and Anderson, H. C. (1992). Immunolocalization of alkaline phosphatase in osteoblasts and matrix vesicles of human fetal bone. Bone Min. 19, 287–298. 92. Robison, R. (1923). The possible significance of hexosephosphoric esters in ossification. Biochem. J. 17, 286–293. 93. Majeska, R. J., and Wuthier, R. E. (1975). Studies on matrix vesicles isolated from chick epiphyseal cartilage. Association of pyrophosphatase and ATPase activities with alkaline phosphatase. Biochim. Biophys. Acta. 391, 51–60.

163 94. Fallon, M. D., Whyte, M. P., and Teitelbaum, S. L. (1980). Stereospecific inhibition of alkaline phosphatase by L-tetramisole prevents in vitro cartilage calcification. Lab. Invest. 43, 489–494. 95. Moss, D. W., Eaton, R. H., Smith, J. K., and Whitby, L. G. (1967). Association of inorganic-pyrophosphatase activity with human alkaline-phosphatase preparations. Biochem. J. 102, 53–57. 96. Rezende, A., Pizauro, J., Ciancaglini, P., and Leone, F. (1994). Phosphodiesterase activity is a novel property of alkaline phosphatase from osseous plate. Biochem. J. 301, 517–522. 97. Anderson, H. C., Hsu, H. H., Morris, D. C., Fedde, K. N., and Whyte, P. W. (1997). Matrix vesicles in osteomalacic hypophosphatasia bone contain apatite-like mineral crystals. Am. J. Path. 151, 1555–1561. 98. Johnson, K. A., Hessle, L., Vaingankar, S., Wennberg, C., Mauro, S., Narisawa, S., Goding, J. W., Sano, K., Millán, J. L., and Terkeltaub, R. (2000). Osteoblast tissue-nonspecific alkaline phosphatase antagonizes and regulates PC-1. Am. J. Physiol. Reg. Integr. Comp. Physiol. 279, R1365–R1377. 99. Hessle, L., Johnsson, K. A., Anderson, H. C., Narisawa, S., Sali, A., Goding, J. W., Terkeltaub, R., and Millán, J. L. (2002). Tissuenonspecific alkaline phosphatase and plasma cell membrane glycoprotein-1 are central antagonistic regulators of bone mineralization. Proc. Natl Acad. Sci. USA 99, 9445–9449. 100. Harmey, D., Hessle, L., Narisawa, S., Johnson, K., Terkeltaub, R., and Millán, J. L. (2004). Concerted regulation of inorganic pyrophosphate and osteopontin by Akp2, Enpp1 and Ank. An integrated model of the pathogenesis of mineralization disorders. Am. J. Pathol. 164, 1199–1209. 101. Okawa, A., Nakamura, I., Goto, S., Moriya, H., Nakamura, Y., and Ikegawa, S. (1998). Mutation in Npps in a mouse model of ossification of the posterior longitudinal ligament of the spine. Nat. Genet. 19, 271–273. 102. Johnson, K., Goding, J., Van Etten, D, Sali, A., Hu, S.-I., Farley, D., Krug, H., Hessle, L., Millán, J. L., and Terkeltaub, R. (2003). Linked deficiencies in extracellular inorganic pyrophosphate (ppi) and osteopontin expression mediate pathologic ossification in PC-1 null mice. J. Bone Min. Res. 18, 994–1004. 103. Ho, A. M., Johnson, M. D., and Kingsley, D. M. (2000). Role of the mouse ank gene in control of tissue calcification and arthritis. Science 289, 265–270. 104. Nürnberg, P., Theile, H., Chandler, D., Höhne, W., Cunningham, M. L., and Ritter, H. (2001). Heterozygous mutations in ANKH, the human ortholog of the mouse progressive ankylosis gene, result in craniometaphyseal dysplasia. Nat. Genet. 28, 37–41. 105. Bostrom, K. (2001). Insights into the mechanism of vascular calcification. Am. J. Cardiol. 88, 20E–22E. 106. Bostrom, K., and Demer, L. L. (2000). Regulatory mechanisms in vascular calcification. Crit. Rev. Eukaryotic Gene Expression 10, 151–158. 107. Picher, M., Burch, L. H., Hirsh, A. J., Spychala J., and Boucher, R. C. (2003). Ecto 5′-nucleotidase and non-specific alkaline phosphatase: two AMP-hydrolyzing ectoenzymes with distinct roles in human airways. J. Biol. Chem. 278, 13468–13479. 108. Takahashi, T., Iwantanti, T., Mizuno, A. S., Morishita, Y., Nishio, H., Kodama, S., and Matsuo, T. (1984). The relationship between phosphoethanolamine level in serum and intractable seizure on hypophosphatasia infantile form. In D.V. Cohn, T. Fugita, J.T. Potts, Jr., R.V. Talmage (eds.), Endocrine Control of Bone and Calcium Metabolism, Vol 8B. Excerpta Medica, Amsterdam, The Netherlands, pp. 93–94. 109. Whyte, M. P., Landt, M., Ryan, L. M., Mulivor, R. A., Henthorn, P. S., Fedde, K. N., Mahuren, J. D., and Coburn, S. P. (1995). Alkaline phosphatase: placental and tissue-nonspecific isoenzymes hydrolyze phosphoethanolamine, inorganic pyrophosphate, and pyridoxal 5’-phosphate. Substrate accumulation in carriers of hypophosphatasia corrects during pregnancy. J. Clin. Invest. 95, 1440–1445.

164 110. Fleshood, H. L., and Pitot, H. C. (1969). O-phosphorylethanolamine ammonia lyase, a new pyridoxal phosphate-dependent enzyme. Biochem. Biophys. Res. Comm. 36, 110–118. 111. Fleshood, H. L., and Pitot, H. C. (1970). The metabolism of O-phosphorylethanolamine in animal tissues. II. Metabolic regulation of O-phosphorylethanolamine phospho-lyase in vivo. Arch. Biochem. Biophysics. 141, 423–429. 112. Grøn, I. H. (1978). Mammalian O-phosphorylethanolamine phospho-lyase activity and its inhibition. Scand. J. Clin. Lab. Invest. 38, 107–112. 113. Van Straalen, J. P., Sanders, E., Prummel, M. F., and Sanders, G. T. B. (1991). Bone-alkaline phosphatase as indicator of bone formation. Clin. Chim. Acta. 201, 27–34. 114. Moss, D. W., and Whitby, L. G. (1975). A simplified heat-inactivation method for investigating alkaline phosphatase isoenzymes in serum. Clin. Chim. Acta. 61, 63–71. 115. Lin, C. W., and Fishman, W. H. (1972). L-Homoarginine. An organspecific, uncompetitive inhibitor of human liver and bone alkaline phosphohydrolases. J. Biol. Chem. 247, 3082–3087. 116. Rosalki, S. B., and Foo, A. Y. (1984). Two new methods for separating and quantifying bone and liver alkaline phosphatase isoenzymes in plasma. Clin. Chem. 30, 1182–1186. 117. Van Hoof, V. O., Lepoutre, L. G., Hoylaerts, M. F., Chevigné, R., and De Broe, M. E. (1988). Improved agarose electrophoretic method for separating alkaline phosphatase isoenzymes in serum. Clin. Chem. 34, 1857–1862. 118. Griffiths, J., and Black, J. (1987). Separation and identification of alkaline phosphatase isoenzymes and isoforms in serum of healthy persons by isoelectric focusing. Clin. Chem. 33, 2171–2177. 119. Magnusson, P., Löfman, O., and Larsson, L. (1992). Determination of alkaline phosphatase isoenzymes in serum by high-performance liquid chromatography with post-column reaction detection. J. Chromat. 576, 79–86. 120. Bailyes, E. M., Seabrook, R. N., Calvin, J., Maguire, G. A., Price, C. P., Siddle, K., and Luzio, J. P. (1987). The preparation of monoclonal antibodies to human bone and liver alkaline phosphatase and their use in immunoaffinity purification and in studying these enzymes when present in serum. Biochem. J. 244, 725–733. 121. Hill, C. S., and Wolfert, R. L. (1990). The preparation of monoclonal antibodies which react preferentially with human bone alkaline

JOSÉ LUIS MILLA´ N

122.

123.

124.

125.

126. 127.

128.

129.

130.

131.

phosphatase and not liver alkaline phosphatase. Clin. Chim. Acta. 186, 315–320. Panigrahi, K., Delmas, P. D., Singer, F., Ryan, W., Reiss, O., Fisher, R., Miller, P. D., Mizrahi, I., Darte, C., Kress, B. C., and Christenson, R. H. (1994). Characteristics of a two-site immunoradiometric assay for human skeletal alkaline phosphatase in serum. Clin. Chem. 40, 822–828. Gomez, Jr., B., Ardakani, S., Ju, J., Jenkins, D., Cerelli, M. J., Daniloff, G. Y., and Kung, V. T. (1995). Monoclonal antibody assay for measuring bone-specific alkaline phosphatase activity in serum. Clin. Chem. 41, 1560–1566. Broyles, D. L., Nielsen, R. G., Bussett, E. M., Lu, W. D., Mizrahi, I. A., Nunnelly, P. A., Ngo, T. A., Noell, J., Christenson, R. H., and Kress, B. C. (1998). Analytical and clinical performance characteristics of Tandem-MP Ostase, a new immunoassay for serum bone alkaline phosphatase. Clin. Chem. 44, 2139–2147. Magnusson, P., Ärlestig, L., Paus, E., Di Mauro, S., Testa, M. P., Stigbrand, T., Farley, J. R., Knustad, K., and Millán, J. L. (2002). Monoclonal antibodies against bone alkaline phosphatase: The ISOBM TD-9 workshop. Tumor Biol. 23, 228–248. Kleerekoper, M. (1996). Biochemical markers of bone remodeling. Am. J. Med. Sci. 312, 270–277. Miller, P. D., Baran, D. T., Bilezikian, J. P., Greenspan, S. L., Lindsay, R., Riggs, B. L., and Watts, N. B. (1999). Practical clinical application of biochemical markers of bone turnover. J. Clin. Densitometry 2, 323–342. Riggs, B.L. (2000). Are biochemical markers for bone turnover clinically useful for monitoring therapy in individual osteoporotic patients? Bone 26, 551–552. Magnusson, P., Larsson, L., Magnusson, M., Davie, M. W. J., and Sharp, C. A. (1999). Isoforms of bone alkaline phosphatase: characterization and origin in human trabecular and cortical bone. J. Bone Min. Res. 14, 1926–1933. Magnusson, P., Larsson, L., Englund, G., Larsson, B., Strang, P., and Selin-Sjögren, L. (1998). Differences of bone alkaline phosphatase isoforms in metastatic bone disease and discrepant effects of clodronate on different skeletal sites indicated by the location of pain. Clin. Chem. 44, 1621–1628. Magnusson, P., Sharp, C. A., Magnusson, M., Risteli, J., Davie, M. W. J., and Larsson, L. (2001). Effect of chronic renal failure on bone turnover and bone alkaline phosphatase isoforms. Kidney Int. 60, 257–265.

Chapter 10

Acid Phosphatases Helena Kaija, and Lila O.T. Patrikainen Sari L. Alatalo H. Kalervo Väänänen Pirkko T. Vihko

I. Acid Phosphatases II. Tartrate-Resistant Acid Phosphatase (TRACP)

Finnish Red Cross Blood Service, Helsinki, Finland Institute of Biomedicine, Department of Anatomy, University of Turku, Turku, Finland Department of Biological and Environmental Sciences, Division of Biochemistry, University of Helsinki, Finland

III. Prostatic Acid Phosphatase (PAP) References

I. ACID PHOSPHATASES

for catalysis. One obvious difference among phosphatases is the presence or absence of metal ion cofactors. Acid phosphatases catalyze the hydrolysis of phosphate ester bond in the following reaction at an optimal pH below 7:

The phosphate ester bond functions as an extremely important linkage within the living cell. It participates in storage and transfer of the genetic information, carries chemical energy, and regulates the activity of enzymes and signaling molecules in the cell. Enzymes capable of acting on ester bonds (Enzyme Commission classification number EC 3.1) and catalyzing the cleavage of phosphate esters (EC 3.1.3) constitute the subclass of phosphohydrolases, i.e. phosphatases. Phosphatases can be classified according to several frameworks [1]. They can be divided into groups based on their substrate type. Nonspecific phosphatases catalyze the hydrolysis of almost any phosphate ester, whereas protein phosphatases prefer phosphoproteins or phosphopeptides as substrates. Nonspecific phosphatases can be divided into alkaline phosphatases (EC 3.1.3.1) and acid phosphatases (EC 3.1.3.2) based on their optimal pH Dynamics of Bone and Cartilage Metabolism

Research Center for Molecular Endocrinology, University of Oulu, Finland

orthophosphoric monoester + H2O = alcohol + phosphate Acid phosphatases have rather wide substrate specificity and some of them catalyze transphosphorylations between phosphoesters and alcohols. Nonspecific acid phosphatases recycle phosphate in metabolic reactions [2]. Protein phosphatases are a structurally miscellaneous group of enzymes that remove phosphate groups that have been attached to amino acid residues of proteins by protein kinases. Protein phosphatases have different mechanisms of action, subcellular localization, and substrate specificity with varied demands for optimal pH. Protein phosphatases comprise two groups based upon their substrate specificity. Protein-tyrosine phosphatases (PTPases, EC 3.1.3.48) 165

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prefer to remove phosphate from tyrosine residue and serine/threonine protein phosphatases (PPases, EC 3.1.3.16) from serine or threonine residue: phosphoprotein + H2O = protein + phosphate The PTPase group includes dual-specificity phosphatases, which are capable of hydrolyzing phosphorylated tyrosine, serine, and threonine, and phosphorylated lipids. Protein phosphatases regulate signal-transduction pathways [2]. Approximately 30% of intracellular proteins are subject to reversible protein phosphorylation [3].

A. Mammalian Acid Phosphatases Li and co-workers demonstrated, in 1970, seven different acid phosphatase isoenzymes in human leukocytes by acrylamide gel electrophoresis and ion exchange chromatography [4]. The heterogeneity of acid phosphatases within tissues had been reported previously, but the relationship between acid phosphatase isoenzymes and tissues and different cells in tissues were not clarified until later due to technical limitations. Based on the differences at the structural level of the gene, acid phosphatases can be divided into five isoenzymes, namely prostatic (AcPP, PAP, human chromosomal location 3q21–q23), lysosomal (AcP2, LAP, human chromosomal location 11p12–p11), erythrocytic (AcP1, soluble acid phosphatase, human chromosomal location 2p25), testicular (AcPT, human chromosomal localization 19q13), and macrophagic acid phosphatases (AcP5, tartrate-resistant acid phosphatase, chromosomal location 19p13.3–p13.1). Acid phosphatases can also be distinguished based on their molecular weight into low-molecular-weight acid phosphatases and high-molecular-weight acid phosphatases, or on the basis of their resistance to inhibition by tartrate into tartrate-sensitive and tartrate-resistant acid phosphatases. Both protein tyrosine phosphatase group and protein serine/threonine phosphatase group include phosphatases with acidic optimal pH.

B. Biochemical and Physiological Significance of Phosphatase Activity The ubiquity of phosphatases in nature makes them versatile regulators of metabolic reactions and cell signaling. Phosphoregulation is involved in many biological events and often occurs as a network-like cascade, in which activity of one phosphatase or kinase is dependent on the upstream activity of another. Many protein kinases are

regulated by phosphorylation constituting an important substrate group for protein phosphatases. Regulation of enzyme activities is most frequently performed by sequential cycles of phosphorylations and dephosphorylation. There seem to be fewer phosphatases than kinases, and phosphatases are not as well characterized as kinases. Activators, inhibitors as well as substrates, affect the rate at which the enzyme is phosphorylated/ dephosphorylated. Regulation of glucose and lipid metabolism at the cellular as well as molecular level is mainly carried out by alternating phosphorylations and dephosphorylations. Activation of the insulin receptor requires the tyrosine autophosphorylation [5]. Dephosphorylation of any one of the activating phosphotyrosine residues dramatically reduces the receptor kinase activity, thus functioning as negative regulators of the insulin signaling cascade [6]. Alterations in the extracellular environment through hormones, growth factors, or cytokines, induce signal transduction pathways to target transcription factors and transcriptional coregulators leading to their phosphorylation or dephosphorylation. Regulation of transcriptional activity by kinases and phosphatases is a rapid and readily reversible mechanism with great versatility and flexibility [7]. Phosphorylation/dephosphorylation plays also a central role in cell-cycle progression by regulating the activity of CDK/cyclin complexes. Although the concentration of human acid phosphatases is normally low, significant changes in their expression occur in certain diseases combined with specific tissue location. This observation suggests that acid phosphatases may be usable as cell-organelle markers as well as diagnostic markers.

II. TARTRATE-RESISTANT ACID PHOSPHATASE (TRACP) Tartrate-resistant acid phosphatase (TRACP) belongs to the acid phosphatases (EC 3.1.3.2), sharing catalytic activity towards phosphoesters in an acidic environment. In 1970, Li and colleagues discovered at least seven distinct isoenzymes 0, 1, 2, 3a, 3b, 4, and 5 from human leukocytes, numbered in order of their increasing electrophoretic mobility towards the cathode [4, 8]. “Band 5” acid phosphatase (AcP 5) or type-5 AcP was the only resistant to L(+)-tartrate inhibition, from which the name tartrateresistant acid phosphatase derives. Also, platelets and red blood cells contain tartrate-resistant AcPs, but these are clearly different from type-5 AcP based on their molecular weights, substrate specificities, electrophoretic mobilities, and antigenic properties. Hence, TRACP specifically refers

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to type-5 AcP demonstrated on acidic acrylamide gels. Previously, the commonly used abbreviation for type-5 AcP was TRAP, but following a recommendation from the nomenclature committee an additional ‘C’ was added to TRACP as a separation from alkaline phosphatases (APs). The members of TRACP family are also known as purple acid phosphatases because of the bound metal ions in their active sites giving an intense purple color.

A. Isolation of Mammalian TRACP Enzymes TRACP enzymes have been purified from many mammalian sources, including pig uterine fluid, bovine bone [9], the spleens of patients affected with hairy cell leukemia [10, 11], and rat [12, 13] bone. When TRACP enzymes were purified from distinct sources they were shown to exist either in monomeric, approximately 30–35 kDa proteins, or in two subunits, sized approximately 16 and 20–25 kDa, connected by a covalent bond. For a long time this discrepancy remained enigmatic, until in the 1990s several research groups were able to demonstrate an exposed, highly antigenic, protease-sensitive loop structure in the sequence of TRACP [14, 15]. After cleavage of this loop, the enzyme was turned into two subunits connected by a disulfide bridge between two conserved cysteine residues.

B. Expression of TRACP Physiologically, TRACP expression is found in cells of the mononuclear phagocyte system, most abundantly in bone-resorbing osteoclasts, alveolar macrophages, and dendritic cells. High TRACP activity has been demonstrated in bone, spleen, liver, lung, and placenta, with lower amounts in kidney, skin, colon, stomach, ileum, testis, and brain. TRACP activity in other than bone tissue is primarily due to the wide distribution of TRACP-positive dendritic cells and activated macrophages. Unstimulated monocytes in bone marrow or peripheral blood are TRACP negative. Pathologically, TRACP expression is elevated in diseases such as hairy cell leukemia (HCL) and Gaucher’s disease [16]; hence, TRACP is clinically used as a marker enzyme for these diseases. Both hairy cells in HCL and spleen cells of patients with Gaucher’s disease stain strongly for TRACP. Elevated TRACP activities in serum have been demonstrated in situations with increased osteoclastic activity such as hyperparathyroidism, Paget’s disease, inflamed synovial joints, postmenopausal women, osteoporosis, osteoclastoma, and various cancers with bone metastases.

C. Subcellular Localization of TRACP TRACP activity in osteoclasts was originally detected adjacent to the bone-resorbing organ, ruffled border membrane, with nonspecific substrates such as β-glycerophosphate and adenosine triphosphate (ATP). However, TRACP is not able to hydrolyze β-glycerophosphate, and the activity detected with this substrate was most probably tartrate-sensitive lysosomal AcP (LAP) activity. Also, ATP is a better substrate for vacuolar-type proton pump, V-ATPase, located in the ruffled border membrane than for TRACP [17]. Immunohistochemical detection revealed more reliable results, showing intracellular vesicular staining for TRACP with no positive reaction at the ruffled border region [18, 19]. However, contradictory data have been observed using an immunogold technique demonstrating intensive TRACP staining both at the ruffled border membrane and intracellularly in vesicular structures [20]. In studies using electron microscopy or separation of subcellular granules by density gradient centrifugation, followed by cytochemical or immunocytochemical staining, TRACP was localized to lysosomes, lysosome-like organelles of the Golgi complex, and microsomes in spleen cells of HCL, rat liver macrophages, rat bone osteoclasts, and bovine spleen histiocytes. Within the lysosome-like organelles, TRACP activity was restricted to the inner membrane surface [19], which is consistent with the sequence data, suggesting a putative N-terminal lysosomal leader sequence in the TRACP gene. Nevertheless, confocal microscopy studies revealed no or only partial colocalization with TRACP and known lysosomal markers, lysosome-associated membrane protein 1 and the cationindependent mannose-6-phosphate receptor in osteoclasts and in alveolar macrophages [21]. Instead, TRACP was shown to colocalize with bone degradation products at the transcytotic route in resorbing osteoclasts [19] and with internalized S. aureus in alveolar macrophages [21]. The transcytotic route of osteoclasts is analogous to the antigen presentation route of activated macrophages, as both are a transport route from a late endosomal/lysosomal compartment to a cell membrane. These data indicate that TRACP may be localized into some new, still unidentified, vesicular compartment both in osteoclasts and in macrophages. Some researchers have also found TRACP in other bone cells, namely osteoblasts and osteocytes, but the intensity of the staining has been lower than in osteoclasts. However, in other studies osteoblasts and osteocytes have been totally negative for TRACP [18, 20]. It has been speculated that osteoblasts would endocytose TRACP secreted by osteoclasts, and TRACP would be as a coupling factor between bone resorption and formation [22].

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Nevertheless, researchers are in agreement that TRACP is mainly produced by osteoclasts and secreted from them during bone resorption [23]. Active enzyme can be detected in the serum of mammals as a complex with α2-macroglobulin [24] and used as a marker of bone resorption [25, 26].

D. Protein Structure Purple acid phosphatase from the red kidney bean (Protein Data Bank code, 1KBP) was the first member of the TRACP family for which the crystal structure became available at 2.9 Å resolution in 1995 [27]. In 1999, the three-dimensional structures for three mammalian TRACP enzymes were published almost simultaneously: for pig TRACP (1UTE) at 1.55 Å resolution [28], for rat TRACP from bone (1QHW] at 2.2 Å resolution [29], and for recombinant rat TRACP (1QFC) at 2.7 Å resolution [30]. Despite less than 15% nucleotide sequence homology between distinct mammalian TRACP enzymes, the amino acid sequences exhibit over 80% identity and the protein

folds are very similar. The structure of mammalian TRACP enzymes is approximately spherical with dimensions of 45 × 45 × 40 Å. The enzymes are formed of two sandwiched β-sheets flanked by α-helical segments facing the solvent. The enzymes show internal symmetry, with the metal ions bound at the interface between the two halves comprising the active site (Fig. 1). Mammalian enzymes are approximately 35 kDa monomeric proteins, whereas plant enzymes are typically dimers of 55 kDa subunits. Mammalian TRACPs contain two iron atoms in their active site, while plant enzymes have one iron atom and either zinc or manganese [27]. The antiferromagnetically spin-coupled binuclear iron center of the mammalian TRACP enzymes exists in two stable interconvertible states: pink, reduced, and enzymatically active, with a mixedvalent Fe3+–Fe2+ cluster; and purple, oxidized, and catalytically inactive, with a binuclear pair as Fe3+–Fe3+ [31]. Another of the iron atoms in the active site of mammalian TRACP enzymes is stabilized into the ferric (Fe3+) form by a tyrosine residue (Tyr55; numbering according to human sequence), accounting for the characteristic purple color of TRACPs. A histidine (His223) and an

Figure 1 The three-dimensional structure of rat bone TRACP determined at 2.2 Å resolution (PDB code 1QHW). The core of the enzyme was formed by two seven-stranded β-sheets which, packed together, making up a β-sandwich. The β-sandwich was surrounded by three solvent-exposed α-helices on both sides. The di-iron center was located on the other edge of the β-sandwich. Two N-acetylglucosamine (NAG) molecules were linked to the side chain of Asn118. One zinc and one sulfate ion were found in the active site and one further zinc and sulfate ion were involved in crystal contact. The protease sensitive loop, seen at the upper left corner of the structure, is located close to the active site [29].

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aspartate (Asp14) residue coordinate the ferric iron among tyrosine residue. A µ-hydroxo bridge and an aspartate carboxylate (Asp52) connect the ferric iron to the redox active iron (Fe2+/3+), which is in turn coordinated by two histidines (His186, His221) and an asparagine (Asn91) residue [27]. The residues coordinating the binuclear active site of TRACP enzymes are essentially identical to those of plant TRACPs and the regulatory Ser/Thr protein phosphatases (PPs) [32]. However, both plant TRACPs and PPs have a ferric iron together with a zinc or manganese in their active site instead of two iron atoms. Additionally, PPs differ essentially from the TRACP family by having a water molecule instead of the tyrosine residue coordinating the ferric iron, accounting for the loss of purple color [27]. Near the active site pocket is an exposed, proteasesensitive loop. The loop structure of mammalian enzymes can be cleaved by trypsin or cathepsins, yielding two fragments connected by a disulfide bridge. Proteolytic cleavage shifts the pH optimum approximately from 5 to 6 and activates the acid phosphatase activity of TRACP by 3–8fold, also causing changes in the EPR spectrum [32, 15]. Enzymatic activation was shown to be not due to the removal of steric hindrance or conformational changes, but rather due to changes in the molecular interaction between residue Asp146 of the loop region and residue Asn91 coordinating the redox active iron in the active site [15]. From the two putative N-linked glycosylation sites Asn97 and Asn128, the previous one was shown to be glycosylated [29, 30]. In uteroferrin, the carbohydrate side chain consists of phosphorylated, high-mannose-type oligosaccharide composed of six mannose residues and two N-acetylglucosamine residues [34]. Similar to uteroferrin, human bone TRACP also contains only N-linked highmannose carbohydrate [11]. Rat, mouse, and human TRACP, but not uteroferrin, contain a consensus motif for protein tyrosine phosphatases (PTPase); Cys(X)5Arg [35]. The conserved cysteine residue in the motif for PTPases forms a covalent bond with phosphate during catalysis and is essential for enzyme activity. However, in TRACPs the conserved cysteine forms a disulfide bridge, being unable to participate in dephosphorylation.

shown to be due to additional sialic acid in the carbohydrate side chain of TRACP 5a. After sialidase treatment TRACP 5a is converted to TRACP 5b. TRACP 5a has a pH optimum of 5.2 and relatively low specific activity, whereas TRACP 5b has a pH optimum of 5.8 with substantially higher specific activity. In addition to a different pH optimum and carbohydrate content, TRACP 5a and 5b are also distinguishable by antigenic properties (Table I) [36]. Based on current knowledge, osteoclasts contain and secrete only TRACP 5b isoform, while activated macrophages express both TRACP 5a and 5b, but secrete only TRACP 5a into the circulation [37]. The osteoclastic origin of TRACP 5b makes it a specific marker for osteoclasts and bone resorption [38, 39].

F. TRACP Gene The mammalian TRACP genes have been cloned from several species. The exon–intron structure of the TRACP gene is highly conserved, consisting of five exons with the translation initial signal (A+1TG) at the beginning of exon 2. The TRACP gene in mammals is 975–1020 bp long, and it encodes approximately 1.5 kb mRNA. Translation will encode a protein with 323–325 amino acids, including a signal peptide of 19 residues and two potential sites for N-glycosylation. Very recently multiple tissue-specific promoters were identified [40], solving the problem with contradictory results with several transcription start points (tsp). Walsh and colleagues were able to identify three distinct mRNAs with differential 5′UTRs, but similar 3′ ends of the mRNA from the first base of exon 2. The novel 5′-UTRs represent alternative first exons located upstream of the known 5′UTR [41]. This genomic structure is conserved at least between mouse and human. Expression of the most distal 5′-UTR (exon 1A) in mouse is restricted to adult bone and spleen tissue, while exon 1B is expressed primarily in tissues containing TRACP-positive nonhematopoietic cells. The known 5′-UTR (exon 1C) is expressed in cells

Table I.

E. Isoforms 5a and 5b Soon after the discovery of TRACP, Lam and co-workers demonstrated that TRACP exists in two distinct isoforms in human serum [36]. Two tartrate-resistant bands with slightly different molecular weights were observed and named according to their electrophoretic mobility as TRACP 5a and TRACP 5b. The difference in size was

Properties of TRACP 5a and 5b TRACP 5a

TRACP 5b

Sialic acid pH optimum for AcP Specific activity of AcP Polypeptide chain

Present 4.9–5.1 Low Intact ∼35 kDa

Origin

Macrophages and dendritic cells

Not present 5.7–5.9 High Cleaved 16 kDa + 20–25 kDa Osteoclasts

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originating from a myeloid lineage in osteoclasts and macrophages [40]. Although the genomic structure of the 5′-flanking region of the human and mouse TRACP genes is conserved, the sequence identity is limited. Studies on the 5′-flanking region of the TRACP gene have indicated a complex transcriptional regulation. The most conserved region of the TRACP 5′-flanking region lies within the 2-kb segment from the tsp A+1TG, and this area has been under extensive study. The possibility that TRACP might be involved in iron transport or storage has led several groups to examine the regulatory influence of intracellular iron on TRACP expression. Simmen and co-workers found a putative iron-response element in the 5′-flanking region of the uteroferrin gene in 1989 [42], and a few years later Alcantara and colleagues were able to show that expression of TRACP was regulated by iron and the iron responsive element was located between base pairs –1846 and –1240 in the mouse promoter [43]. This iron regulation is very complex and occurs at the transcriptional level. Two iron-delivering agents, hemin (ferric protoporphyrin IX) and transferrin have opposite effects on TRACP expression, hemin decreases and transferrin increases [43, 44] TRACP expression. Two heminresponsive elements (HRE) were found within the human 5′-flanking region, representing binding sites for an HRE-binding protein (HRE-BP), and both are needed for the hemin-induced regulation of the human TRACP gene expression [45]. Monocyte-associated transcription factor PU.1 acts synergistically with osteoclast commitment factor MiTF (microphthalmia transcription factor) regulating the TRACP expression during terminal differentiation of osteoclasts [46]. TRACP expression is stimulated by RANKL, and at least upstream stimulatory factors (USF) 1 and 2 are involved in RANKL-mediated activation [47]. In addition to these, the promoter area of TRACP contains numerous candidate transcription factor binding sequences, including those for Sp1, AP1, GT-1, and H-APF-1 [41].

G. Functions of TRACP The biological function(s) as well as natural substrate(s) of TRACP are still unknown despite intensive studies within recent decades. However, the highly conserved structure of TRACP enzymes throughout the animal kingdom provides strong evidence for an important biological role for the enzyme. Location in distinct tissues and cells, as well as functions shown in vitro, suggest that TRACP may have several functions in different cells and cell compartments.

1. G.1. AcP

TRACP is primarily an AcP catalyzing the hydrolysis of a wide range of phosphate monoesters such as β-umbelliferyl phosphate, p-nitrophenyl phosphate, α-naphthyl phosphate and phosphotyrosine. Also, phosphoanhydrides such as pyrophosphate and nucleoside tri- and diphosphates are hydrolyzed by TRACP [12], but aliphatic phosphoesters such as monophosphates, β-glycerophosphate, mannose-6-phosphate, phosphoserine, and phosphothreonine are not. However, TRACP can catalyze the release of phosphate from certain phosphoproteins that carry phosphoserine residue such as osteopontin and osteonectin [49]. Nevertheless, the AcP activity of TRACP prefers phosphotyrosine-containing proteins over other phosphoproteins, suggesting a specific protein tyrosine phosphatase (PTPase) activity for TRACP [9]. Phosphorylation of specific tyrosine residues by kinases plays an important role in various cell events such as signal transduction, activation, proliferation, and differentiation. As a PTPase, TRACP could regulate protein-tyrosine kinases and thereby influence various cellular events [50]. The AcP activity of TRACP is competitively inhibited by inorganic phosphate and its analogs, e.g. vanadate, arsenate, and molybdate. Fluoride, tungstate, copper, and zinc cause noncompetitive inhibition of AcP activity [51]. Reducing agents such as β-mercaptoethanol, dithiothreitol, ascorbic acid, cysteine, and glutathione, activate the AcP activity of TRACP by reducing the ferric iron into the ferrous form and simultaneously changing the purple color to pink [51]. The reverse reaction from pink to purple (Fe2+ → Fe3+) causes inactivation and is achieved by oxidizing agents such as H2O2 and ferricyanide. Dithionite inactivates TRACP irreversibly by removing the metal ions from the active site, thereby bleaching its color [48, 51]. In contrast, common inhibitors for AcPs such as cyanide, azide, tartrate, and p-nitrophenol show no inhibition of the AcP activity of TRACP [51]. Combined structural and mechanistic studies provide a model for the catalytic mechanism of TRACP, differing from the mechanism of PTPases. In the first step, the phosphate group of the substrate is coordinated to the divalent metal ion. Formation of the enzyme–substrate complex is followed by a nucleophilic attack on the phosphorus, leading to the release of the product alcohol. Three possible candidates have been proposed for the attacking group: (1) a terminal Fe3+-bound hydroxide [52]; (2) a bridging hydroxide [53]; or (3) a hydroxide residing in the second coordination sphere of Fe3+ [54]. In the final step of catalysis, the metal-bound phosphate group is released. TRACP has been hypothesized to participate in bone resorption by dephosphorylating bone matrix phosphoproteins osteopontin, osteonectin, and bone sialoprotein [49].

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Osteopontin has been shown to bind to osteoclast cellsurface integrins via the RGD motif, thereby mediating substrate adhesion, at least in vitro [55]. After dephosphorylation osteopontin is no longer able to support osteoclast binding to the substrate, suggesting that TRACP could regulate osteoclast attachment [49]. Dephosphorylation of bone matrix phosphoproteins suggests a catabolic function for TRACP in bone resorption.

H. TRACP-Deficient and Transgenic Mice

2. G.2. Generator of Reactive Oxygen Species (ROS)

Three individual research groups have been able to demonstrate by different methods that TRACP is capable of generating ROS via Fenton’s reaction [19, 56, 57]. In an enzymatically active reduced state, TRACP has a mixedvalent Fe3+–Fe2+ active site, in which the ferrous iron (Fe2+) is able to react with hydrogen peroxide and produce ferric iron (Fe3+) and hydroxyl radical (•OH) (Equation 1). The newly formed ferric iron is still able to react with hydrogen peroxide to form superoxide anion (•O2−) and a ferrous iron (Equation 2). Hydroxyl radical formation: TRACP-Fe2+ + H2O2 → TRACP-Fe3+ + •OH + OH−

two activities are functionally independent [61]. Beck and co-workers were able to demonstrate that the AcP activity of TRACP is inhibited by the substrate of ROS-generating activity, H2O2, and vice versa, inorganic phosphate blocks H2O2 inhibition, confirming the use of the same active site [62].

(1)

TRACP regeneration and superoxide formation: TRACP-Fe3+ + H2O2 → TRACP-Fe2+ + •O2− + 2H+ (2) The generation of these destructive oxygen radicals can continue as long as H2O2 is available. When hydroxyl radicals alone attack proteins, they cause protein aggregation, but in combination with superoxide anions they cause protein fragmentation without formation of aggregates [58]. Halleen and co-workers were able to demonstrate in vitro that TRACP is able to destroy the major component of the bone matrix, type I collagen, without disruption of the enzyme itself [19]. They also showed that TRACP is localized in transcytotic vesicles of osteoclasts together with bone degradation products, hypothesizing that TRACP would participate in the final degradation of bone matrix molecules intracellularly in the transcytotic vesicles [19]. In general, ROS have been shown to stimulate osteoclastic bone resorption both in vivo and in vitro, and they also enhance the recruitment of osteoclasts [59]. Based on studies with TRACP-deficient or TRACP overexpressing macrophages, TRACP may also have a role in antigen processing by generating destructive ROS [21, 60]. Both the ROS generation and the AcP activity of TRACP use the redox active iron for the catalysis. However, the acid phosphatase activity has an acidic pH-optimum, whereas ROS are generated in a neutral pH [61]. Also, single amino acid TRACP mutants that are completely inactive as phosphatase are still able to produce ROS, suggesting that the

The role of TRACP in bone metabolism and also in other tissues has been elucidated by the TRACP knockout [63] and TRACP over-expressing mouse models [64]. Both models exhibit a bone phenotype; mice lacking TRACP have a mild osteopetrosis resulting from defective bone resorption [63], whereas mice over-expressing TRACP have increased bone resorption that is accompanied by increased bone formation, yielding a net increase in bone turnover [64]. The mechanism by which TRACP increases bone resorption in transgenic mice remains unclear, but nevertheless mice exhibit mild osteoporosis [64]. TRACP-deficient mice suffer from progressive foreshortening and deformity of the long bones and axial skeleton [63]. Tooth eruption and skull plate development is normal in these mice, indicating a role for TRACP in endochondral ossification [65]. Furthermore, the formation of osteoclasts is normal in TRACP-deficient mice, suggesting that TRACP is not essential during osteoclastogenesis [65]. However, osteoclasts isolated from TRACP knockout mice demonstrate defective bone resorption also in vitro determined by a conventional resorption pit assay [63]. Ultrastructural examination of young TRACP null mice revealed that osteoclasts exhibit an increased relative area of ruffled border and accumulation of cytoplasmic vesicles [65] containing filamentous material [66]. The accumulation of intracellular vesicles is probably not due to the impaired secretion, since the location of cathepsin K is normal. Some explanations have been offered for the accumulated vesicles; TRACP-deficient osteoclasts are incapable of totally digesting the internalized bone degradation products, and vacuoles containing filamentous material accumulate into the cytoplasm instead of normal transcytosis and secretion through the FSD [19]; or TRACP may have a function in modulating intracellular vesicular transport by dephosphorylating phosphoproteins [65]. A possible role for TRACP in the immune system has been studied both with TRACP-deficient and TRACP overexpressing macrophages [60]. TRACP-null macrophages have disordered inflammatory responses, such as enhanced superoxide and nitrite production, and increased secretion of proinflammatory cytokines TNFα, IL-1β, and IL-12 [60]. Additionally, TRACP knockout mice exhibit delayed

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clearance of Staphylococcus aureus in vivo [60]. Vice versa, phorbol 12-myristate 13-acetate (PMA) activated macrophages over-expressing TRACP showed increased overall production of ROS and a tendency to higher capacity for bacterial killing in vitro [67]. The biological roles and functional relationship between two acid phosphatases, TRACP and LAP, have remained unknown. Single TRACP [63] or LAP [68] knockouts have distinct, but rather mild phenotypes, suggesting that the phosphatases may in part substitute each other. The double knockout mouse model of both TRACP and LAP demonstrates a more severe phenotype than the sum of the single knockouts, thus strengthening the hypothesis [66]. Mice deficient for both phosphatases showed massive lysosomal storage, for example in Kupffer cells, in bone marrow macrophages and in osteoclasts. Also, the bone phenotype, which was more evident in TRACP- than in LAP-deficient mice, was even more severe in double-deficient mice. The dephosphorylation of osteopontin was severely affected in double-knockout mice, and the authors suggest that TRACP represents the major enzyme implicated in this process [66]. Nevertheless, no data are available from dephosphorylation of osteopontin in single-knockout mice to verify this hypothesis. Furthermore, no attention has been addressed to the ROS generation activity of TRACP, which is not possible to overcome by LAP.

I. TRACP as a Clinical Tool TRACP was considered as a specific histochemical marker for osteoclasts more than 20 years ago [23]. Since then TRACP has been widely accepted as a histological marker of osteoclasts both in tissue sections and in in vitro cell cultures. Without knowing the exact biological role of TRACP in bone metabolism it has become one of the best-known markers for bone-resorbing osteoclasts and their precursors. At present there are up to 1000 references in the database of National Library of Medicine with keywords “TRACP and osteoclast”. The first assays for analytical purposes were kinetic assays to measure the total TRACP activity in serum or plasma. The main problem with these assays was the lack of specificity for TRACP, since they also detected tartrateresistant AcPs from erythrocytes and platelets, which are not related to type-5 TRACP. Kinetic assays were improved by inhibiting non-TRACP AcPs by using fluoride or incubating the samples at 37°C for an hour before measurement [9]. However, these assays still measured total TRACP activity, and lacked the specificity for osteoclastderived TRACP 5b. An important finding was the substrate specificity of distinct AcPs and also TRACP 5a

and 5b. The most common substrate used to detect AcP activity is a 4-nitrophenylphosphate (4-NPP), which is hydrolyzed well by nontype 5 AcPs, as well as TRACP 5a and 5b. Instead, α-naphthyl phosphate and naphtholASBI phosphate have been shown to be more selective substrates and are hydrolyzed effectively by TRACP 5b, but only poorly by TRACP 5a and not at all by other nontype 5 AcPs [69]. Another approach for TRACP 5b-specific kinetic assay was developed using fluoride to inhibit nontype 5 AcPs and heparin to inhibit TRACP 5a activity during measurement [70]. The production of TRACP antibodies made it possible to develop immunoassays to specifically detect TRACP activity or TRACP protein concentration from serum or plasma. Most probably these assays are not selective for TRACP 5a or TRACP 5b, since the TRACP isoforms show very high immunological identity. Until this millennium, the lack of absolute specificity for bone-specific TRACP 5b had held back progress on the use of biochemical assay of TRACP 5b activity or protein concentration as a marker of bone resorption. Both kinetic assays and immunoassays for TRACP showed that individuals with normally or pathologically high rates of bone turnover have increased levels of serum total TRACP activity or protein concentration. Children with physiologically active bone growth and postmenopausal women were shown to have significantly elevated serum TRACP levels compared to healthy premenopausal adults [36, 71]. Marker levels were also elevated in cases where bone resorption is known to be increased, such as osteoporosis, Paget’s disease, humoral hypercalcemia of malignancy, multiple myeloma, osteomalacia, immobilization, and chronic renal failure. An early finding of elevated serum TRACP levels in patients with metastatic bone diseases was later confirmed by several research groups. Also, patients with primary hyperparathyroidism had markedly elevated TRACP levels, which after removal of parathyroid adenoma decreased to normal levels within 2 weeks [72]. In contrast, individuals with hypoparathyroidism, who were expected to have decreased rates of bone resorption, had appropriately less TRACP in serum than healthy controls. The association between serum TRACP and bone resorption has also been confirmed by investigating bone density simultaneously with serum TRACP. A significant inverse correlation has been observed between serum TRACP and bone mineral content or bone mineral density. Similarly, bone density assessed by quantitative ultrasound (QUS) from calcaneus showed a significant negative association with serum TRACP [73]. Furthermore, a significant correlation was observed between histomorphometric parameters of bone resorption and serum TRACP 5b in uremic patients [74]. Additionally, therapies known to

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decrease bone resorption, such as hormone replacement therapy, also decreased serum TRACP, confirming the close relation of the enzyme to bone degradation [11]. Similar to other bone metabolic markers, levels of serum TRACP are also strongly influenced by age, therefore age-matched reference material is essential for proper experimentation [73]. In healthy individuals, the activity of total serum TRACP measured at the pH optimum 5.5 comprises, in approximately equal amounts, the isoforms TRACP 5a and 5b [39]. However, total serum TRACP protein measured by two-site immunoassays contains almost 90% of nonosteoclastic, low-activity TRACP 5a isoform, the remaining 10% being osteoclastic, highly active TRACP 5b [39]. Thus, neither total TRACP activity nor total TRACP protein provides a sensitive and specific tool for assessing the rate of bone resorption.

J. Serum TRACP 5b as a Bone Resorption Marker TRACP 5b expression is strictly limited to boneresorbing osteoclasts, activated macrophages and dendritic cells, which in principle could all be the sources of circulating active TRACP 5b. However, it has been shown in in vitro studies that TRACP 5b is retained almost entirely intracellularly in macrophages and dendritic cells, which secrete exclusively TRACP 5a [37]. Thus, serum TRACP 5b is exclusively secreted by bone-resorbing osteoclasts. Indeed, the existing data demonstrate that osteoclastic TRACP has identical biochemical and physical properties as serum TRACP 5b. Therefore, serum TRACP 5b provides an excellent tool for assessing the rate of bone resorption. Elevated levels of serum TRACP 5b have been observed in conditions with increased bone resorption, such as healthy postmenopausal women and individuals with osteopenia, osteoporosis, or Paget’s disease [26]. Also, children have significantly elevated levels of serum TRACP 5b, which are even further elevated in individuals with vitamin D deficiency [75]. In contrast, antiresorptive treatment such as HRT and alendronate treatment decrease serum TRACP 5b levels in postmenopausal women, as expected [26]. Breast cancer patients with bone metastases have significantly elevated serum TRACP 5b levels [38], and the enzyme levels are closely associated with the metastatic burden. Bisphosphonates are normally used in pain relief and improvement of symptoms of breast cancer patients with bone lesions. Studies revealed that serum TRACP 5b levels specifically reflect the efficacy of bisphosphonate treatment. Similar to breast cancer, TRACP 5b was proven to be associated with the severity of bone disease in multiple myeloma, and specifically monitors the efficacy of antimyeloma treatment [76]. The progression of

the disease was also reflected by serum TRACP 5b, suggesting that TRACP 5b could be used as a predictive tool for bone disease in multiple myeloma. The value of markers for bone metabolism is not only to detect the rate of high bone turnover, but also to predict future bone loss or fractures. A large prospective study with over 1000 postmenopausal women showed that serum TRACP 5b can predict fracture and, in particular, fractures that engage the trabecular bone, such as vertebral fractures [77]. For further discussion of the applications of TRACP as a bone resorption marker, see Chapter 35.

K. Clinical Significance of Serum TRACP 5a Based on the recent data, it seems that serum TRACP 5a has its own specific biological significance in some specific diseases such as rheumatoid arthritis (RA) [78]. Serum total TRACP concentration, but not activity, was shown to be significantly elevated in patients with RA [78]. The elevated levels of TRACP concentration did not correlate with bone metabolic markers bone ALP or NTX, suggesting that the TRACP elevation was not osteoclastic in origin and may not be related to bone turnover. Instead, the levels of TRACP concentration in RA were significantly correlated with C-reactive protein, an acute-phase protein marker of inflammation and indicator of disease activity in RA [37]. Since as much as 90% of the total serum TRACP protein in RA is TRACP 5a, the total TRACP concentration gives a good estimate of TRACP 5a concentration [39]. Based on these results, the authors conclude that increased TRACP protein is a low-activity TRACP 5a isoform from a source other than osteoclasts, probably inflammatory macrophages and dendritic cells abundant in the synovial tissues of affected joints [37]. Further investigations are needed to confirm the clinical significance of serum TRACP 5a concentrations in RA and other chronic inflammatory conditions.

III. PROSTATIC ACID PHOSPHATASE (PAP) Human prostatic acid phosphatase (hPAP) is a glycoprotein synthesized in the epithelial cells of the prostate gland [79–82], and present in spermatic ejeculate [83–85]. This enzyme hydrolyzes a wide range of alkyl and aryl orthophosphate monoesters, including phosphotyrosine [86, 87] and nucleotides [88] and has also been found to dephosphorylate macromolecules, such as phosphopeptides and phosphoproteins [89, 90]. hPAP is categorized as an acid phosphatase, since in has an optimum pH of 4–6.

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A. Gene Structure of hPAP The gene encoding hPAP is located at chromosome 3q21-qter [91]. cDNA encodes a 354-residue protein with a calculated molecular mass of 41 ± 126 Da. The hPAP gene contains ten exons of 170, 96, 87, 153, 99, 93, 133, 83, 104, and 2098 bp, respectively. A 32 amino acid signal sequence and the first eight amino acids of the protein are encoded by exon 1, and the rest of the coding region and the 3′-untranslated region are covered by the exons 2–10 and 10, respectively [92]. When prostatic carcinoma or benign hyperplasia tissue is used as a source of RNA, a major transcription start site can be found 50 nt upstream of the ATG codon of the gene [92, 93]. The length of the 3′ noncoding region in hPAP cDNA varies between 646 and 1913 bp, although, in Northern blot analysis, a 3.3-kb mRNA species is usually observed [94]. The heterogeneity in the length of cDNA is explained by the multiple polyadenylation signals (AAATAA) following two copies of alu-type repetitive sequences in the 3′ noncoding region.

B. Protein Structure of hPAP Figure 2

The hPAP protein is a homodimer with subunits related by a twofold axis [95]. Each subunit comprises two domains. The larger, α/β type domain is composed of a central seven-stranded mixed β-sheet with helices on both sides, while the smaller α domain contains six α-helices and is formed mostly of a long-chain excursion from the first domain. The hPAP active site, which contains an essential histidine (His12) residue [96], is located in a large open cleft between the two domains. This allows the enzyme to accept a large variety of substrates [95]. The rate-limiting step is the breakdown of the covalent phosphoenzyme intermediate by the attack of water on the phosphoroamidate, resulting in the formation of a noncovalent enzyme–inorganic phosphate complex (Fig. 2). The active site was shown to contain two arginines probably involved in the binding of the negatively charged phosphate group of the substrate [97]. The presence of carboxylic acid residues, Asp or Glu, at the active site was indicated by Saini and van Etten [98] and van Etten [99]. Site-directed mutagenesis of hPAP shows His12 acts as an acceptor of the phospho group, Asp258 is a proton donor for the substrate-leaving group, and His257 may participate in substrate-binding or may facilitate the breakdown of the phosphoenzyme complex [100]. These results are consistent with the crystal structure of rPAP [95]. Most of the active site residues of rPAP come from the loops after strands 1 and 4 of the large domain.

Minimal reaction mechanism of rPAP. Modified from

[101a].

Residues Arg11, His12 and Arg15 are part of a sequence motif RHGXRXP characteristic for acid phosphatase [101]. These residues are part of a cluster of conserved amino acid residues in the center of the active site, consisting of residues of Arg11, His12, Arg15, Arg79, His257, and Asp258. The active sites of hPAP and rPAP are very similar in three-dimensional structure, except for the conformation of the Arg15 side chain [102]. A recent study [103] shows that Trp174 was at the active site of the human enzyme, because it was protected by the competitive inhibitor tartrate in the DNPS-Cl modification studies. This is also consistent with the location of a homologous residue in the structure of the rat enzyme. Denaturation–renaturation and subunit reassociation studies show that hPAP activity depends on dimer formation [104]. Oligomerization of rPAP using site-directed mutagenesis found that mutants W106E and H112D, as well as the double mutant W106E/H112D, are monomers without catalytic activity or the ability to bind tartrate [105]. The His112 side-chain forms a hydrogen bond with the Asp76 side-chain between two subunits, and the sidechains of Trp106 are stacked on top of each other across the twofold axis relating the two subunits [95]. Since the PAP active site is located far from the subunits’ interface,

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these results indicate that the formation of dimers induces structural changes which propagate to the active site. Human PAP from native sources possesses molecular heterogeneity. Isoelectric focusing gives up to 30 variants of hPAP from normal, malignant, or hyperplastic prostatic tissue and sera from patients with prostate cancer and seminal fluid [83, 84, 106]. Variation in the amount of sialic acid and other sugar residues and post-translational deamination of glutamine and asparagine account for part of the heterogeneity of hPAP [101, 107]. Furthermore, the carboxyterminal amino acid of hPAP can be threonine, glutamic acid, or aspartic acid [101]. The heterogeneity of hPAP could also be due to the existence of true isoenzymes consisting of different polypeptide chains.

progression [127]. The binding of phosphotyrosine to PAP has been characterized by theoretical modeling and experimental work with peptides derived from the residue sequences of EGFR- and ErbB-2-containing autophosphorysable tyrosine [128, 129] and altogether results confirm that PAP is tyrosine phosphatase as suggested earlier [90]. PAP might function as a dual-specificity protein tyrosine phosphatase as it has been shown to possess affinity also for phosphoserine in proteins in vitro [130]. Binding of PAP to the main high-density lipoprotein apolipoprotein A-I (apoA-I) has also been demonstrated [131]. ApoA-I is lipidated during its progress through intracellular vesicle traffic from the cell surface into early endosomes and further into late endosomes and finally back to the cell surface as a nascent HDL particle [132].

C. Function of hPAP D. Expression of hPAP in Prostate Human PAP is one of the prostate epithelium-specific differential antigens [108–110]. There are two forms of hPAP: one is intracellular and the other secreted [111]. The physiological substrate of hPAP is unknown. However, hPAP is present in large amounts in seminal ejeculate [85], suggesting a physiological role in fertility [112]. The serum activity of the enzyme is frequently elevated in patients with prostate carcinoma and correlates with tumor progression [113–116]. In vitro, hPAP hydrolyzes phosphorycholine, which is found in semen [98], and phosphocreatine, an intracellular high-energy compound present in seminal plasma [117]. Suggested substrates include seminal phosphocholine [105, 118], semenogelins [119], EGFR/ErbB-2 [120], and lysophosphatidic acid [121]. It also has amidolytic activity on semenogelins [122]. hPAP possesses intrinsic protein tyrosine phosphatase (PTP) activity [90, 105, 123], and several lines of evidence indicate that cellular hPAP functions as a neutral PTP, although it shares little sequence homology with other typical PTPs [101, 124, 125]. hPAP also has amidolytic activity on a seminal vesicle protein semenogelin I. The enzyme is able to cleave both peptide substrates derived from the semenogelin sequence and native semenogelin I. The main cleavage sites are at Tyr 292 and Ser 170 [122]. In addition, hPAP interacts with ErbB-2, a member of the erbB receptor tyrosine kinase family, in the androgenpromoted growth of human prostate cancer cells. Androgen-stimulated cell growth concurs with downregulation of cellular hPAP, an elevated p-Tyr level of ErbB-2 and the activation of mitogen-activated protein kinases [126]. Cellular hPAP can down-regulate prostate cancer cell growth, at least partially, by dephosphorylating c-ErbB-2. Therefore, decreased cellular hPAP expression in cancer cells may be involved in prostate cancer

Immunohistochemistry using specific antibodies against PAP has demonstrated that the enzyme is located in the columnar, secretory epithelial cells of the prostate [82, 133–137]. In situ hybridization analysis has shown that hPAP mRNA is confined to the glandular and ductal epithelial cells of the prostate, and that stromal cells are devoid of this mRNA [82]. No hPAP mRNA has been detected in human liver, lung, pancreatic cancer tissue, placenta, breast cancer cells, mononuclear blood cells, or acute promyelocytic leukemia cells, nor in spleen, thymus, testis, ovary, small intestine, colon, or peripheral blood leukocytes [138]. Little is known about the mechanism of tissue-specific regulation of the hPAP gene at the molecular level. Zeliavinski et al. [139] have shown that, in addition to the basic promoter, the region between –1258/−779 is able to enhance the PAP promoter activity in PC-3 and DU-145 human prostate cancer cells, but not in nonprostate cancer cells, indicating that this region is involved in governing the cell type-specific expression of the hPAP gene. Shan et al. [140] showed that a construct containing the sequence of hPAP between the nucleotides –734 and +467 in front of the CAT reporter gene was significantly expressed in the prostate of transgenic mice, while the proximal promoter –734/+50 alone achieved low levels of CAT mRNA in all tissues analyzed. This suggests the involvement of regulatory elements in the intron area and/or the possible interactions of the transcription factors binding along the whole DNA area. It has been shown previously that a 12-bp sequence, GAAAATATGATA is involved in prostatespecific and androgen receptor-dependent gene regulation of the rat probasin gene [141]. In hPAP, five homologs for this previously identified prostate-specific DNA-binding

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site were found between the nucleotides –734 and +467 (Fig. 3). Deletions of the different binding sites of the prostatic transcription factor resulted in bidirectional effects depending on the location of the sites and the hormone status in LNCaP cells. Site C at the proximal promoter of hPAP acts as a positive regulatory element in prostatic cells in an androgen-dependent manner, while site E or sites D and E act as negative regulatory element(s) in the androgendepleted environment [140].

E. Androgen Regulation of hPAP In cell culture models, both up- and down-regulation by androgens have been reported for hPAP. The amount of hPAP released into LNCaP cell culture medium was decreased to 26% of the control level in 7 days when a synthetic androgen, R1881, was present in charcoalstripped serum [142]. Accordingly, 5α-dihydrotestosterone (DHT) treatment was found to decrease the activity of hPAP in these cells. A stimulatory effect of androgen on hPAP secretion has been confirmed by Horozewicz et al. [143] and Lin et al. [144]. hPAP mRNA levels are also increased when tissue slides derived from benign prostatic hyperplasia are treated with DHT and fibroblastic growth factor (bFGF) [145]. A biphasic pattern of the effect of androgen on LNCaP cells has been reported: stimulation of growth and inhibition of hPAP secretion was detected at less than 1 nM concentrations of androgen, while an opposite effect was observed at higher concentrations [146, 147]. Shan et al. [148] showed that reporter constructs of hPAP promoter covering the region –734/+467 were functional in both prostatic and nonprostatic cell lines in transient transfections. This region contained two putative AREs, which have been shown to have androgen receptor-binding ability in vitro, but the promoter could not be induced with androgen, glucocorticoid, or progesterone, indicating that steroids cannot directly regulate hPAP gene expression via receptor binding to these AREs.

Figure 3

Location of the prostate-specific DNA-binding site GAAAATATGATA-related sequences and potential androgen response elements in the schematic representation of the regulatory region of the hPAP gene. Binding capacities of the elements for the prostatic protein in vitro are also marked (−, +, +++). In vitro androgen receptor-binding capacities are indicated by + or ++ above AREs.

Androgen may regulate hPAP expression differently in diverse physiological or pathological conditions. Both up- and down-regulation of hPAP by 12-otetradecanoyl phorbol-13-acetate (TPA), a protein kinase C (PKC) activator, has been reported [142, 149]. Lin et al. [149] reported that TPA is able to increase the secretion of hPAP in a dose- and time-dependent fashion in the androgen-responsive LNCaP cell line. This TPA stimulation of hPAP secretion was more potent than the conventional stimulating agent DHT at the same concentration. Furthermore, the action of TPA and DHT on hPAP secretion was blocked by five different PKC inhibitors. DHT, as well as TPA, could rapidly modulate PKC activity. Therefore, PKC can regulate hPAP secretion and may also be involved in the DHT action on hPAP secretion. So far, very little is known about physiological substrates of acid phosphatases, but increasing interest in kinases/ phosphatases is obviously opening a new insight into this research area.

References 1. Vincent, J. B., Crowder, M. W., and Averill, B. A. (1992). Hydrolysis of phosphate monoesters: a biological problem with multiple chemical solutions. Trends. Biochem. Sci. 17, 105–110. 2. Lohse, D. L., Denu, J. M., and Dixon, J. E. (1995). Insights derived from the structures of the Ser/Thr phosphatases calcineurin and protein phosphatase 1. Structure 3, 987–990. 3. Barford, D., Das, A. K., and Egloff, M. P. (1998). The structure and mechanism of protein phosphatases: insights into catalysis and regulation. Annu. Rev. Biophys. Biomol. Struct. 27, 133–164. 4. Li, C. Y., Yam, L. T., and Lam, K. W. (1970). Studies of acid phosphatase isoenzymes in human leukocytes demonstration of isoenzyme cell specificity. J. Histochem. Cytochem. 18, 901–910. 5. Saltiel, A. R., and Kahn, C. R. (2001). Insulin signalling and the regulation of glucose and lipid metabolism. Nature 414, 799–806. 6. Hashimoto, N. Feener, E. P., Zhang, W. R., and Goldstein, B. J. (1992). J. Biol. Chem. 267, 13811–138142. 7. Jackson, S. P. (1992). Insulin receptor protein-tyrosine phosphatases. Leukocyte common antigen-related phosphatase rapidly deactivates the insulin receptor kinase by preferential dephosphorylation of the receptor regulatory domain. Trends. Cell. Biol. 2, 104–108. 8. Li, C. Y., Yam, L. T., and Lam, K. W. (1970). Acid phosphatase isoenzyme in human leukocytes in normal and pathologic conditions. J. Histochem. Cytochem. 18 (7), 473–481. 9. Lau, K. H., Freeman, T. K., and Baylink, D. J. (1987a). Purification and characterization of an acid phosphatase that displays phosphotyrosylprotein phosphatase activity from bovine cortical bone matrix. J. Biol. Chem. 262 (3), 1389–1397. 10. Lam, K. W., and Yam, L. T. (1977). Biochemical characterization of the tartrate-resistant acid phosphatase of human spleen with leukemic reticuloendotheliosis as a pyrophosphatase. Clin. Chem. 23 (1), 89–94. 11. Halleen, J., Hentunen, T. A., Hellman, J., and Väänänen, H. K. (1996). Tartrate-resistant acid phosphatase from human bone: purification and development of an immunoassay. J. Bone. Miner. Res. 11 (10), 1444–1452. 12. Anderson, T. R., and Toverud, S. U. (1986). Purification and characterization of purple acid phosphatase from developing rat bone. Arch. Biochem. Biophys. 247 (1), 131–139.

Chapter 10 Acid Phosphatases 13. Ek-Rylander, B., Bergman, T., and Andersson, G. (1991). Characterization of a tartrate-resistant acid phosphatase (ATPase) from rat bone: hydrodynamic properties and N-terminal amino acid sequence. J. Bone. Miner. Res. 6 (4), 365–373. 14. Ljusberg, J., Ek-Rylander, B., and Andersson, G. (1999). Tartrate-resistant purple acid phosphatase is synthesized as a latent proenzyme and activated by cysteine proteinases. Biochem. J. 343 Pt 1, 63–69. 15. Funhoff, E. G., Ljusberg, J., Wang, Y., Andersson, G., and Averill, B. A. (2001). Mutational analysis of the interaction between active site residues and the loop region in mammalian purple acid phosphatases. Biochemistry 40 (38), 11614–11622. 16. Yam, L. T., Li, C. Y., and Lam, K. W. (1971). Tartrate-resistant acid phosphatase isoenzyme in the reticulum cells of leukemic reticuloendotheliosis. N. Engl. J. Med. 284 (7), 357–360. 17. Väänänen, H. K., Karhukorpi, E. K., Sundquist, K., Wallmark, B., Roininen, I., Hentunen, T., Tuukkanen, J., and Lakkakorpi, P. (1990). Evidence for the presence of a proton pump of the vacuolar H(+)-ATPase type in the ruffled borders of osteoclasts. J. Cell Biol. 111 (3), 1305–1311. 18. Clark, S. A., Ambrose, W. W., Anderson, T. R., Terrell, R. S., and Toverud, S. U. (1989). Ultrastructural localization of tartrate-resistant, purple acid phosphatase in rat osteoclasts by histochemistry and immunocytochemistry. J. Bone Miner. Res. 4 (3), 399–405. 19. Halleen, J. M., Räisänen, S., Salo, J. J., Reddy, S. V., Roodman, G. D., Hentunen, T. A., Lehenkari, P. P., Kaija, H., Vihko, P., and Väänänen, H. K. (1999b). Intracellular fragmentation of bone resorption products by reactive oxygen species generated by osteoclastic tartrateresistant acid phosphatase. J. Biol. Chem. 274 (33), 22907–22910. 20. Reinholt, F. P., Widholm, S. M., Ek-Rylander, B., and Andersson, G. (1990). Ultrastructural localization of a tartrate-resistant acid ATPase in bone. J. Bone Miner. Res. 5 (10), 1055–1061. 21. Räisänen, S. R., Halleen, J., Parikka, V., and Väänänen, H. K. (2001). Tartrate-resistant acid phosphatase facilitates hydroxyl radical formation and colocalizes with phagocytosed Staphylococcus aureus in alveolar macrophages. Biochem. Biophys. Res. Commun. 288 (1), 142–150. 22. Sheu, T. J., Schwarz, E. M., Martinez, D. A., O’Keefe, R. J., Rosier, R. N., Zuscik, M. J., and Puzas, J. E. (2003). A phage display technique identifies a novel regulator of cell differentiation. J. Biol. Chem. 278 (1), 438–443. 23. Minkin, C. (1982). Bone acid phosphatase: tartrate-resistant acid phosphatase as a marker of osteoclast function. Calcif. Tissue Int. 34 (3), 285–290. 24. Brehme, C. S., Roman, S., Shaffer, J., and Wolfert, R. (1999). Tartrateresistant acid phosphatase forms complexes with alpha2-macroglobulin in serum. J. Bone Miner. Res. 14 (2), 311–318. 25. Lau, K. H., Onishi, T., Wergedal, J. E., Singer, F. R., and Baylink, D. J. (1987b). Characterization and assay of tartrate-resistant acid phosphatase activity in serum: potential use to assess bone resorption. Clin. Chem. 33 (4), 458–462. 26. Halleen, J. M., Alatalo, S. L., Suominen, H., Cheng, S., Janckila, A. J., Väänänen, H. K. (2000). Tartrate-resistant acid phosphatase 5b: a novel serum marker of bone resorption. J. Bone Miner. Res. 15 (7), 1337–1345. 27. Sträter, N., Klabunde, T., Tucker, P., Witzel, H., and Krebs, B. (1995). Crystal structure of a purple acid phosphatase containing a dinuclear Fe(III)-Zn(II) active site. Science 268 (5216), 1489–1492. 28. Guddat, L. W., McAlpine, A. S., Hume, D., Hamilton, S., de Jersey, J., and Martin, J. L. (1999). Crystal structure of mammalian purple acid phosphatase. Arch. Biochem. Biophys. 352 (1), 97–102. 29. Lindqvist, Y., Johansson, E., Kaija, H., Vihko, P., and Schneider, G. (1999). Three-dimensional structure of a mammalian purple acid phosphatase at 2.2 A resolution with a mu-(hydr)oxo bridged di-iron center. J. Mol. Biol. 291 (1), 135–147. 30. Uppenberg, J., Lindqvist, F., Svensson, C., Ek-Rylander, B., and Andersson, G. (1999). Crystal structure of a mammalian purple acid phosphatase. J. Mol. Biol. 290 (1), 201–211.

177 31. Davis, J. C., and Averill, B. A. (1982). Evidence for a spin-coupled binuclear iron unit at the active site of the purple acid phosphatase from beef spleen. Proc. Natl Acad. Sci, USA 79 (15), 4623–4627. 32. Vincent, J. B., and Averill, B. A. (1990b). Sequence homology between purple acid phosphatases and phosphoprotein phosphatases. Are phosphoprotein phosphatases metalloproteins containing oxide-bridged dinuclear metal centers? FEBS Lett 263 (2), 265–268. 33. Kaija, H., Jia, J., Lindqvist, Y., Andersson, G. and Vihko, P. (1999). Tartrate-resistant bone acid phosphatase: large-scale production and purification of the recombinant enzyme, characterization, and crystallization. J. Bone Miner. Res. 14 (3), 424–430. 34. Baumbach, G. A., Saunders, P. T., Bazer, F. W., and Roberts, R. M. (1984). Uteroferrin has N-asparagine-linked high-mannose-type oligosaccharides that contain mannose 6-phosphate. Proc. Natl Acad. Sci. USA 81 (10), 2985–2989. 35. Zhang, Z. Y., Wang, Y., Wu, L., Fauman, E. B., Stuckey, J. A., Schubert, H. L., Saper, M. A., and Dixon, J. E. (1994). The Cys(X)5Arg catalytic motif in phosphoester hydrolysis. Biochemistry 33 (51), 15266–15270. 36. Lam, W. K., Eastlund, D. T., Li, C. Y., and Yam, L. T. (1978a). Biochemical properties of tartrate-resistant acid phosphatase in serum of adults and children. Clin. Chem. 24 (7), 1105–1108. 37. Janckila, A. J., Neustadt, D. H., Nakasato, Y. R., Halleen, J. M., Hentunen, T. and Yam, L. T. (2002a). Serum tartrate-resistant acid phosphatase isoforms in rheumatoid arthritis. Clin Chim Acta 320 (1–2), 49–58. 38. Halleen, J. M., Alatalo, S. L., Janckila, A. J., Woitge, H. W., Seibel, M. J., and Väänänen, H. K. (2001). Serum tartrate-resistant acid phosphatase 5b is a specific and sensitive marker of bone resorption. Clin. Chem. 47 (3), 597–600. 39. Janckila, A. J., Takahashi, K., Sun, S. Z., and Yam, L. T. (2001b). Tartrate-resistant acid phosphatase isoform 5b as serum marker for osteoclastic activity. Clin. Chem. 47 (1), 74–80. 40. Walsh, N. C., Cahill, M., Carninci, P., Kawai, J., Okazaki, Y., Hayashizaki, Y., Hume, D. A., and Cassady, A. I. (2003). Multiple tissuespecific promoters control expression of the murine tartrate-resistant acid phosphatase gene. Gene 307, 111–123. 41. Reddy, S. V., Scarcez, T., Windle, J. J., Leach, R. J., Hundley, J. E., Chirgwin, J. M., Chou, J. Y., and Roodman, G. D. (1993). Cloning and characterization of the 5′-flanking region of the mouse tartrateresistant acid phosphatase gene. J. Bone. Miner. Res. 8 (10), 1263–1270. 42. Simmen, R. C., Srinivas, V., and Roberts, R. M. (1989). CDNA sequence, gene organization, and progesterone induction of mRNA for uteroferrin, a porcine uterine iron transport protein. DNA 8 (8), 543–554. 43. Alcantara, O., Reddy, S. V., Roodman, G. D., and Boldt, D. H. (1996). Inhibition of tartrate-resistant acid phosphatase gene expression by hemin and protoporhyrin IX. Identification of a hemin-responsive inhibitor of transcription. Blood 88 (6), 2288–2297. 44. Reddy, S. V., Alcantara, O., Roodman, G. D., and Boldt, D. H. (1996). Inhibition of tartrate-resistant acid phosphatase gene expression by hemin and protoporphyrin IX. Identification of a hemin-responsive inhibitor of transcription. Blood 88 (6), 2288–2297. 45. Fleckenstein, E. C., Dirks, W. G., and Drexler, H. G. (2000). The human tartrate-resistant acid phosphatase (TRAP): involvement of the hemin responsive elements (HRE) in transcriptional regulation. Leuk Lymphoma 36 (5–6): 603–612. 46. Luchin, A., Suchting, S., Merson, T., Rosol, T. J., Hume, D. A., Cassady, A. I., and Ostrowski, M. C. (2001). Genetic and physical interactions between Microphthalmia transcription factor and PU.1 are necessary for osteoclast gene expression and differentiation. J. Biol. Chem. 276 (39), 36703–36710. 47. Liu, Y., Shi, Z., Silveira, A., Liu, J., Sawadogo, M., Yang, H., and Feng, X. (2003). Involvement of upstream stimulatory factors 1 and 2 in RANKL-induced transcription of tartrate-resistant acid phosphatase

178

48.

49.

50.

51.

52.

53.

54.

55.

56.

57.

58.

59.

60.

61.

62.

63.

64.

gene during osteoclast differentiation. J. Biol. Chem. 278 (23), 20603–20611. Halleen, J. M., Kaija, H., Stepan, J. J., Vihko, P., and Väänänen, H. K. (1998b). Studies on the protein tyrosine phosphatase activity of tartrateresistant acid phosphatase. Arch. Biochem. Biophys. 352 (1), 97–102 Ek-Rylander, B., Flores, M., Wendel, M., Heinegard, D., and Andersson, G. (1994). Dephosphorylation of osteopontin and bone sialoprotein by osteoclastic tartrate-resistant acid phosphatase. Modulation of osteoclast adhesion in vitro. J. Biol. Chem. 269 (21), 14853–14856. Janckila, A. J., Woodford, T. A., Lam, K. W., Li, C. Y., and Yam, L. (1992). Protein-tyrosine phosphatase activity of hairy cell tartrateresistant acid phosphatase. Leukemia 6 (3), 199–203. Schlosnagle, D. C., Sander, E. G., Bazer, F. W., and Roberts, R. M. (1976). Requirement of an essential thiol group and ferric iron for the activity of the progesterone-induced porcine uterine purple phosphatase. J. Biol. Chem. 251 (15), 4680–4685. Klabünde, T., Sträter, N., Frohlich, R., Witzel, H., and Krebs, B. (1996). Mechanism of Fe(III)-Zn(II) purple acid phosphatase based on crystal structures. J. Mol. Biol. 259 (4), 737–748. Wang, X., Randall, C. R., True, A. E., and Que, L., Jr. (1996). X-ray absorption spectroscopic studies of the FeZn derivative of uteroferrin. Biochemistry 35 (44), 13946–13954. Merkx, M., Pinkse, M. W., and Averill, B. A. (1999). Evidence for nonbridged coordination of p-nitrophenyl phosphate to the dinuclear Fe(III)-M(II) center in bovine spleen purple acid phosphatase during enzymatic turnover. Biochemistry 38 (31), 9914–9925. Flores, M. E., Norgard, M., Heinegard, D., Reinholt, F. P., and Andersson, G. (1992). RGD-directed attachment of isolated rat osteoclasts to osteopontin, bone sialoprotein, and fibronectin. Exp. Cell. Res. 201 (2), 526–530. Sibille, J. C., Doi, K., and Aisen, P. (1987). Hydroxyl radical formation and iron-binding proteins. Stimulation by the purple acid phosphatases. J. Biol. Chem. 262 (1), 59–62. Hayman, A. R., and Cox, T. M. (1994). Purple acid phosphatase of the human macrophage and osteoclast. Characterization, molecular properties, and crystallization of the recombinant di-iron-oxo protein secreted by baculovirus-infected insect cells. J. Biol. Chem. 269 (2), 1294–1300. Davies, K. J., and Delsignore, M. E. (1987). Protein damage and degradation by oxygen radicals. III. Modification of secondary and tertiary structure. J. Biol. Chem. 262 (20), 9908–9913. Garrett, I. R., Boyce, B. F., Oreffo, R. O., Bonewald, L., Poser, J., and Mundy, G. R. (1990). Oxygen-derived free radicals stimulate osteoclastic bone resorption in rodent bone in vitro and in vivo. J. Clin. Invest. 85 (3), 632–639. Bune, A. J., Hayman, A. R., Evans, M. J., and Cox, T. M. (2001). Mice lacking tartrate-resistant acid phosphatase (Acp 5) have disordered macrophage inflammatory responses and reduced clearance of the pathogen, Staphylococcus aureus. Immunology 102 (1), 103–113. Kaija, H. Alatalo, S. L., Halleen, J. M., Lindqvist, Y., Schneider, G., Väänänen, H. K., and Vihko, P. (2002). Phosphatase and oxygen radicalgenerating activities of mammalian purple acid phosphatase are functionally independent. Biochem. Biophys. Res. Commun. 292 (1), 128–132. Beck, J. L., Durack, M. C., Hamilton, S. E., and de Jersey, J. (1999). Irreversible inactivation of purple acid phosphatase by hydrogen peroxide and ascorbate. J. Inorg. Biochem. 73 (4), 245–252. Hayman, A. R., Jones, S. J., Boyde, A., Foster, D., Colledge, W. H., Carlton, M. B., Evans, M. J., and Cox, T. M. (1996). Mice lacking tartrate-resistant acid phosphatase (Acp 5) have disrupted endochondral ossification and mild osteopetrosis. Development 122 (10), 3151–3162. Angel, N. Z., Walsh, N., Forwood, M. R., Ostrowski, M. C., Cassady, A. I., and Hume, D. A. (2000). Transgenic mice overexpressing tartrate-resistant acid phosphatase exhibit an increased rate of bone turnover. J. Bone Miner. Res. 15 (1), 103–110.

HELENA KAIJA ET AL. 65. Hollberg, K., Hultenby, K., Hayman, A., Cox, T., and Andersson, G. (2002). Osteoclasts from mice deficient in tartrate-resistant acid phosphatase have altered ruffled borders and disturbed intracellular vesicular transport. Exp. Cell. Res. 279 (2), 227–238. 66. Suter, A., Everts, V., Boyde, A., Jones, S. J., Lullmann-Rauch, R., Hartmann, D., Hayman, A. R., Cox, T. M., Evans, M. J., Meister, T., von Figura, K., and Saftig, P. (2001). Overlapping functions of lysosomal acid phosphatase (LAP) and tartrate-resistant acid phosphatase (Acp5) revealed by doubly deficient mice. Development 128 (23), 4899–4910. 67. Räisänen, S. (2004). Differential effects of carbonic anhydrase III and tartrate-resistant acid phosphatase on intracellular radical balance. Institute of Biomedicine, Department of Anatomy, Turku, University of Turku: 1–93 [Thesis]. 68. Saftig, P., Hartmann, D., Lullmann-Rauch, R., Wolff, J., Evers, M., Koster, A., Hetman, M., von Figura, K., and Peters, C. (1997). Mice deficient in lysosomal acid phosphatase develop lysosomal storage in the kidney and central nervous system. J. Biol. Chem. 272 (30), 18628–18635. 69. Janckila, A. J., Takahashi, K., Sun, S. Z., and Yam, L. T. (2001a). Naphthol-ASBI phosphate as a preferred substrate for tartrate-resistant acid phosphatase isoform 5b. J. Bone. Miner. Res. 16 (4), 788–793. 70. Nakanishi, M., Yoh, K., Miura, T., Ohasi, T., Rai, S. K., and Uchida, K. (2000). Development of a kinetic assay for band 5b tartrate-resistant acid phosphatase activity in serum. Clin. Chem. 46 (4), 469–473. 71. Halleen, J. M., Karp, M., Viloma, S., Laaksonen, P., Hellman, J., Käkönen, S. M., Stepan, J. J., Holmes, S., Väänänen, H., and Pettersson, K. (1999a). Two-site immunoassays for osteoclastic tartrate-resistant acid phosphatase based on characterization of six monoclonal antibodies. J. Bone. Miner. Res. 14 (3), 464–469. 72. Stepan, J. J., Silinkova-Malkova, E., Havranek, T., Formankova, J., Zichova, M., Lachmanova, J., Strakova, M., Broulik, P., and Pacovsky, V. (1983). Relationship of plasma tartrate resistant acid phosphatase to the bone isoenzyme of serum alkaline phosphatase in hyperparathyroidism. Clin. Chim. Acta. 133 (2), 189–200. 73. Nakanishi, M., Yoh, K., Uchida, K., Maruo, S., Rai, S. K., and Matsuoka, A. (1999) Clinical usefulness of serum tartrate-resistant fluoride-sensitive acid phosphatase activity in evaluating bone turnover. J. Bone Miner. Metab. 17 (2), 125–130. 74. Chu, P., Chao, T. Y., Lin, Y. F., Janckila, A. J., and Yam, L. T. (2003). Correlation between histomorphometric parameters of bone resorption and serum type 5b tartrate-resistant acid phosphatase in uremic patients on maintenance hemodialysis. Am. J. Kidney Dis. 41 (5), 1052–1059. 75. Cheng, S., Tylavsky, F., Kröger, H., Kärkkäinen, M., Lyytikäinen, A., Koistinen, A., Mahonen, A., Alen, M., Halleen, J., Väänänen, K., and Lamberg-Allardt, C. (2003). Association of low 25-hydroxyvitamin D concentrations with elevated parathyroid hormone concentrations and low cortical bone density in early pubertal and prepubertal Finnish girls. Am. J. Clin. Nutr. 78 (3), 485–492. 76. Terpos, E., de la Fuente, J., Szydlo, R., Hatjiharissi, E., Viniou, N., Meletis, J., Yataganas, X., Goldman, J. M., and Rahemtulla, A. (2003). Tartrate-resistant acid phosphatase isoform 5b: a novel serum marker for monitoring bone disease in multiple myeloma. Int. J. Cancer. 106 (3), 455–457. 77. Gerdhem, P., Ivaska, K. K., Alatalo, S. L., Halleen, J. M., Hellman, J., Isaksson, A., Pettersson, K., Väänänen, H. K., Åkesson, K., and Obrant, K. J. (2004). Biochemical markers of bone metabolism and prediction of fracture in elderly women. J. Bone Miner. Res. 19 (3), 386–393. 78. Nakasato, Y. R., Janckila, A. J., Halleen, J. M., Väänänen, H. K., Walton, S. P., and Yam, L. T. (1999). Clinical significance of immunoassays for type-5 tartrate-resistant acid phosphatase. Clin. Chem. 45 (12), 2150–2157. 79. Derechin, M., Ostrowski, W., Galka, M., and Barnard, E. A. (1971). Acid phosphomonoesterase of human prostate: molecular weight dissociation and chemical composition. Biochim. Biophys. Acta. 250, 143–154.

Chapter 10 Acid Phosphatases 80. Ostrowski, W., Bhargava, A. K., Dziembor, E., Gizler, M., Gryszkiewicz, J.T. and Barnard, E. A. (1976). Acid phosphomonoesterase of human prostate. Carbohydrate content and optical properties. Biochim. Biophys. Acat. 453, 262–269. 81. Risley, M. J., and van Etten, R. L. (1987). Structures of the carbohydrate moieties of human prostatic acid phosphatase elucidated by 1H nuclear magnetic resonance spectroscopy. Arch. Biochem. Biophys. 258, 404–412. 82. Hakalahti, L., Vihko, P., Henttu, P., Autio-Harmainen, H., Soini, Y., and Vihko, R. (1993). Evaluation of PAP and PSA gene expression in prostatic hyperplasia and prostatic carcinoma using Northern blot analysis, in situ hybridization and immunochemical stainings with monoclonal and bispecific antibodies. Int. J. Cancer 55, 590–597. 83. Vihko, P. (1978). Characterization of the principal human prostatic acid phosphatase isoenzyme, purified by affinity chromatography and isolectric focusing. Part II. Clin. Chem. 24, 1783–1787. 84. Vihko, P., Kontturi, M., and Korhonen, L. K. (1978). Purification of human prostatic acid phosphatase by affinity chromatography and isoelectric focusing. Part I. Clin. Chem. 24, 466–470. 85. Rönnberg, L., Vihko, P., Sajanti, E., and Vihko, R. (1981). Clomiphene citrate administration to normogonadotropic subfertile men: Blood hormone changes and activation of acid phosphatase in seminal fluid. Int. J. Androl. 4, 372–378. 86. Apostol, I., Kuciel, R., Wasylewska, E., and Ostrowski, W. S. (1985). Phosphotyrosine as a substrate of acid and alkaline phosphatases. Acta. Biochem. Pol. 32, 187–197. 87. Vihko, P., Kurkela, R., Porvari, K., Herrala, A., Lindfors, A., Lindqvist, Y., and Schneider, G. (1993). Rat acid phosphatase: Overexression of active secreted enzyme by recombinant baculovirus-infected insect cells, molecular properties and crystallization. Proc. Natl Acad. Sci. USA. 90, 799–803. 88. Dziembor-Gryszkiewicz, E., Fikus, M., Kazimierczuk, Z., and Ostrowski, W. (1978). Activity of human prostatic acid phosphatase toward purine 5′-phosphonucleosides. Bull. Acad. Pol. Sci. Biol. 26, 815–821. 89. Wasylewska, E., Czubak, J., and Ostrowski, W. S. (1983). Phosphoprotein phosphatase activity of human prostate acid phosphatase. Acta. Biochim. Pol. 30, 175–184. 90. Li, H. X., Chernoff, J., Chen, L. B., and Kirschonbaum, A. (1984). A phosphotyrosyl-protein activity associated with acid phosphatase from human prostate gland. Eur. J. Biochem. 138, 45–51. 91. Winqvist, R., Virkkunen, P., Grzeschik, K. H., and Vihko, P. (1989). Chromosomal localization to 3q21→qter and two TaqI RFLPs of the human prostae-specific acid phosphatase gene (ACCP). Cytogenet. Cell. Genet. 52, 68–71. 92. Virkkunen, P., Hedberg, P., Palvimo, J., Birr, E., Porvari, K., Ruokonen, M., Taavitsainen, P., Jänne, O., and Vihko, P. (1994). Structure comparison of human and rat prostate-specific acid phosphatase genes and their promoters: identification of putative androgen response elements. Biochem. Biophys. Res. Comm. 202, 49–57. 93. Sharief, F. S., and Li, S. S. L. (1992). Structure of human prostatic acid phosphatase gene. Biochem. Biophys. Res. Comm. 184, 1468–1476. 94. Vihko, P., Virkkunen, P., Henttu, P., Roiko, K., Solin, T., and Huhtala, M. L. (1988). Molecular cloning and sequence analysis of cDNA encoding human prostatic acid phosphatase. FEBS Lett 236, 275–281. 95. Schneider, G., Lindqvist, Y., and Vihko, P. (1993). Three-dimensional structure of rat acid phosphatase. EMBO. J. 12, 2609–2615. 96. Davidson, R. E. (1990). Structural aspects of human prostatic acid phosphatase and fast atom bombardment mass spectrometric analysis of E. coli L-threonine deaminase. Ph.D. thesis, Purdue University, West Lafayette, IN. 97. Mctigue, J. J., and van Etten, R. L. (1978). An essential arginine residue in human prostatic acid phosphatase. Biochim. Biophys. Acta. 523, 422–429.

179 98. Saini, M. S., and van Etten, R. L. (1979). An essential carboxylic acid group in human prostate acid phosphatase. Biochim. Biophys. Acta. 568, 370–376. 99. van Etten, R. L. (1982). Human prostatic acid phosphatase: a histidine phosphatase. Ann. N. Y. Acad. Sci. 390, 27–51. 100. Ostanin, K., Saeed, A., and van Etten, R. L. (1994). Heterologous expression of human prostatic acid phosphatase and site-directed mutagenesis of the enzyme active site. J. Biol. Chem. 269, 9680–9684. 101. Van Etten, R. L., Davidson, R., Stevis, P. E., MacArthur, H., and Moore, D. L. (1991). Covalent structure, disulfide bonding, and identification reactive surface and active site residues of human prostatic acid phosphatase. J. Biol. Chem. 266, 2313–2319. 101a.Lindqvist, Y., Schneider, G., and Vihko, P. (1994). Crystal structure of rat acid phosphatase complexed with the transition-state analogs vanadate and molybdate. Implications for the reaction mechanism. Eur. J. Biochem. 221, 139–142. 102. Lindqvist, Y., Schneider, G., and Vihko, P. (1993). Three-dimensional structure of rat acid phosphatase in complex with L(+)-tartrate. J. Biol. Chem. 268, 20744–20746. 103. Zhang, Z., Ostanin, K., and van Etten, R. L. (1997). Covalent modification and site-directed mutagenesis of an active site tryptophan on human prostatic acid phosphatase. Acta. Biochim. Pol. 44, 659–672. 104. Kuciel, R., Bakalova, A., Mazurkiewicz, A., Bilska, A., and Ostrowski, W. (1990). Is the subunit of prostatic acid phosphatase active? Reversible denaturation of prostatic acid phosphatase. Biochem. Int. 22, 329–334. 105. Porvari, K. S., Herrala, A. M., Kurkela, R. M., Taavitsainen, P. A., Lindqvist, Y., Schneider, G., and Vihko, P. T. (1994). Site-directed mutagenesis of prostatic acid phosphatase: catalytically important aspartic acid 258, substrate specificity, and oligomerization. J. Biol. Chem. 269, 22642–22646. 106. Chu, T. M., Wang, M. C., Merrin, C., Valenzuela, L., and Murphy, G. P. (1978). Isoenzymes of human prostatic acid phosphatase. Ann, N. Y. Acad. Sci. 390, 16–26. 107. Smith, J. K., and Whitby, L. B. (1968). The heterogeneity of prostatic acid phosphatase. Biochim. Biophys. Acta. 151, 607–618. 108. Yam, L. T. (1974). Clinical significance of the human prostatic acid phosphatase. Am. J. Med. 56, 604–616. 109. Lam, K. W., Li, C. Y., Yam, L. T., Smith, R. S., and Hacker, B. (1982). Comparison of prostatic and nonprostatic acid phosphatase. Ann. N. Y. Acad. Sci. 390, 1–15. 110. Kamoshida, S., and Tsutsumi, Y. (1990). Extraprostatic localization of prostatic acid phosphatase and prostate-specific antigen: distribution in cloacogenic glandular epithelium and sex-dependent expression in human anal gland. Hum. Pathol. 21, 1108–1111. 111. Vihko, P. (1979). Human prostatic acid phosphatase. Purification of a minor enzyme and comparisons of the enzymes. Invest. Urol. 16, 349–352. 112. Coffey, D. S. and Pienta, K. J. (1987). New concepts of study in the control of normal and cancer growth of the prostate. In, D. S. Coffey, N. Bruchovsky, W. A, Gardner, M. I. Resnick, and J. P. Karr (eds), “Current Concepts and Approaches to the Study of Prostate Cancer.” Alan R Liss, Inc., New York, p. 1–73. 113. Gutman, A. B., and Gutman, E. N. (1938). An acid phosphatase occurring in the serum of patients with metastasizing carcinoma of prostate gland. J. Clin. Invest. 17, 473–478. 114. Choe, B. K., Pontes, E. J., Dong, M. K. and Rose, N. R. (1980). Double-antibody immunoassay for human prostatic acid phosphatase. Clin. Chem. 26, 1854–1959. 115. Griffiths, J. C. (1989). Prostate-specific acid phosphatase: re-evaluation of radioimmunoassay in diagnosing prostatic disease. Clin. Chem. 26, 433–436.

180 116. Vihko, P., Kostama, A., Jänne, O., Sajanti, E. and Vihko, R. (1980). Rapid radioimmunoassay for prostate-specific acid phosphatase in human serum. Clin. Chem. 26, 1544–1547. 117. Lee, H. J., Fillers, W. S., and Iyyengar, M. R. (1988). Phosphocreatine, an intracellular high-energy compound, is found in the extracellular fluid of the seminal vesicles in mice and rats. Proc. Natl Acad. Sci. USA 85, 7265–7269. 118. Saini, M. S., and Van Etten, L. R. (1981). A clinical assay for prostatic acid phosphatase using choline phosphate as a substrate: comparison with thymolphthalein phosphate. Prostate 2, 359–368. 119. Ek, P., Malm, J., Lilja, H., Carlsson, L., and Ronquist, G. (2002). Exogenous protein kinases, A and C, but not endogenous prostasomeassociated protein kinase, phosphorylate semenogelins I and II from human semen. J. Androl. 23, 806–814. 120. Lin, M.-F., and Clinton, G. M. (1988). The epidermal growth factor receptor from prostate cells is dephosphorylated by a prostate-specific phosphotyrosyl phosphatase. Mol. Cell Biol. 8, 5477–5485. 121. Tanaka, M., Kishi, Y., Takanezawa, Y., Kakehi, Y., Aoki, J., and Arai, H. (2004). Prostatic acid phosphatase degrades lysophosphatidic acid in seminal plasma. FEBS Lett. 571, 197–204. 122. Brillard-Bourdet, M., Rehault, S., Juliano, L., Ferrer, M., Moreau, T. and Gauthier, F. (2002). Amidolytic activity of prostatic acid phosphatase on human semenogelins and semenogelin-derived synthetic substrates. Eur. J. Biochem. 269, 390–395. 123. Boissonneault, M., Chapdelaine, A., and Chevalier, S. (1995). The enhancement by pervanadate of tyrosine phosphorylation of prostatic proteins occurs through the inhibition of membrane-associated tyrosine phosphatase. Mol. Cell Biochem. 153, 139–144. 124. Guan, K., Haun, R. S., Watson, S. J., Geahlen, R. L., and Dixon, J. E. (1990). Cloning and expression of a protein-tyrosine-phosphatase. Proc. Natl Acad. Sci. USA 87, 1501–1505. 125. Chernoff, J., Schievella, A. R., Jost, C. A., Erikson, R. L., Neel, B. G. (1990). Cloning of a cDNA for a major human protein-tyrosinephosphatase. Proc. Natl Acad. Sci. USA 87, 2735–2739. 126. Meng, T. C., Lee, M. S., and Lin, M. F. (2000). Interaction between protein tyrosine phosphatase and protein tyrosine kinase is involved in androgen-promoted growth of human prostate cancer cells. Oncogene 19, 2664–2677. 127. Lin, M. F., Lee, M. S., Zhou, X. W., Andressen, J. C., Meng, T. C., Johansson, S. L., West, W. W., Taylor, R. J., Anderson, J. R., and Lin, F. F. (2001). Decreased expression of cellular prostatic acid phosphatase increases tumorigenicity of human prostate cancer cells. J. Urol. 166, 1943–1950. 128. Vihko, P. (1993). 129. Sharma, S., Pirilä, P., Kaija, H., Porvari, K., Vihko, P., and Juffer, A. H. Theoretical investigations of prostatic acid phosphatase. Proteins – structure, function and genetics (in press). 130. Lin, M. F., and Clinton, G. M. (1986). Human prostatic acid phosphatase has phosphotyrosyl protein phosphatase activity. Biochem. J. 235, 351–357. 131. Vihko, P., Wahlberg, L., Ehnholm, C., Lukka, M., and Vihko, R. (1986). Acid phosphatases bind to the main high densitylipoprotein apolipoprotein A-I. FEBS Lett. 202, 309–313. 132. Neufeld, E. B., Stonik, J. A., Demosky, S. J., Jr., Knapper, C. L., Combs, C. A., Cooney, A., Comly, M., Dwyer, N., BlanchetteMackie, J., Remaley, A. T., et al. (2004). The ABCA1 transporter modulates late endocytic trafficking: insights from the correction of the genetic defect in Tangier disease. J. Biol. Chem. 279, 15571–15578. 133. Aumüller, G., and Seitz, J. (1985). Cytochemistry and biochemistry of acid phosphatases. VI: Immunoelectron microscopic studies on

HELENA KAIJA ET AL.

134.

135. 136.

137.

138.

139.

140.

141.

142.

143.

144.

145.

146.

147.

148.

149.

human prostatic and leucocytic acid phosphatases. Prostate 17, 31–40. Mori, K., and Wakasugi, C. (1985). Immunocytochemical demonstration of prostatic acid phosphatase: different secretion kinetics between normal, hyperplastic and neoplastic prostate. J. Urol. 133, 877–883. Lilja, H., and Abrahamsson, P. A. (1988). Three predominant proteins secreted by the human prostate gland. Prostate 12, 29–38. Sinha, A. A., Gleason, D. F., Wilson, M. J., Wick, M. R., Reddy, P. K., and Blackard, C. E. (1988). Relationship of prostatic acid phosphatase localization in human prostate by a monoclonal antibody with the Gleason grading system. Prostate 13, 1–15. Lam, K. W., Li Cy., Yam, L. T., Sun, T., Lee, G., and Ziesmer, S. (1989). Improved immunohistochemical detection of prostatic acid phosphatase by a monoclonal antibody. Prostate 15, 13–21. Zeliavinski, S., Comeau, D., and Lin, M. F. (1998). Cloning and analysis of the promoter activity of the human prostatic acid phosphatase gene. Biochem. Biophys. Res. Commun. 245, 108–112. Zeliavinski, S., Igawa, T., Lim, S., Taylor, R., and L M.-F. (2002). Identification and characterization of regulatory elements of the human prostatic acid phosphatase promoter. Oncogene 21, 3696–3705. Shan, J. D., Porvari, K., Kivinen, A., Patrikainen, L., Halmetkytö, M., Jänne, J., and Vihko, P. (2003). Tissue-specific expression of the prostatic acid phosphatase promoter constructs. Biochem. Biophys. Res. Comm. 311, 864–869. Patrikainen, L., Shan, J., Porvari, K., and Vihko, P. (1999). Identification of the DNA-binding site of a regulatory protein involved in prostate-specific and androgen receptor-dependent gene expression. Endocrinology 140, 2063–2070. Henttu, P., and Vihko, P. (1996). Effect of tumor-promoting phorbol ester on androgen-regulated gene expression in prostatic LNCaP cells. “In vitro Biology of Sex Steroid Hormone Action.” Churchill Livingstone, Tokyo, pp. 384–395. Horozewicz, J., Leong, S., Kawinski, E., Karr, J., Rosenthal, H., Chu, T., Mirand, E., and Murphy, G. (1983). LNCaP model of human prostatic carcinoma. Cancer Res. 43, 1809–1818. Lin, M. F., Garcia-Arenas, R., Chao, Y. C., Lai, M., Patel, P. C., and Xia, X. Z. (1993). Regulation of prostatic acid phosphatase expression and secretion by androgen in LNCaP human prostate carcinoma cells. Arch. Biochem. Biophys. 300, 384–390. Dulinska, J., Laidler, P., and Labrfz, M. (2002). Comparative analysis of prostatic acid phosphatase and prostate-specific antigen mRNA levels in hyperplastic prostate stimulated with steroid hormones and growth factors. Acta. Biochim. Pol. 49, 357–368. Henttu, P., Liao, S., and Vihko, P. (1992). Androgens up-regulate the human prostate-specific antigen mRNA but down-regulate the prostatic acid phosphatase mRNA in the LNCaP cell line. Endocrinology 130, 766–772. Langler, E. G., van Uffelen, C. J., Blankenstein, M. A., van Steenbrugge, G. J., and Mulder, E. (1993). Effect of culture conditions on androgen sensitivity of the human prostatic cancer cell line LNCaP. Prostate 23, 213–223. Shan, J. D., Porvari, K., Ruokonen, M., Karhu, A., Launonen, V., Hedberg, P., Oikarinen, J., and Vihko, P. (1997). Steroid-involved transcriptional regulation of human genes encoding prostatic acid phosphatase, prostate-specific antigen, and prostate-specific glandular kallikrein. Endocrinology 138, 3764–3770. Lin, M. F., Zhang, X. Q., Dean, J., and Lin, F-F. (2001). Protein kinase C pathway is involved in regulating the secretion of prostatic acid phosphatase in human prostate cancer cells. Cell Biol. Int. 25(11), 1139–1148.

Chapter 11

Matrix Proteinases Ian M. Clark Gillian Murphy

School of Biological Sciences, University of East Anglia, Norwich NR4 7TJ, UK Dept of Oncology, Cambridge University, Cambridge CB2 2XY, UK

I. Introduction II. Aspartic Proteinases III. Cysteine Proteinases

IV. Serine Proteinases V. Metalloproteinases References

I. INTRODUCTION

II. ASPARTIC PROTEINASES

Proteolysis of the extracellular matrix (ECM) of tissues is a feature of the normal remodeling associated with physiological processes such as morphogenesis, growth, and wound healing. Degradation of the ECM is also associated with many pathologies including arthritic diseases, osteoporosis, and periodontal disease. More recently, the concept of the role of proteinases in the modulation of cell–cell and cell–ECM interactions as a means of signaling to the cell and the determination of cell phenotype has emerged. This more subtle activity is clearly of importance not only in developmental processes, but also in pathology. Endopeptidases are thought to be the key enzymes in ECM degradation and in vitro evidence is available that members of all four major classes (metallo-, serine, cysteine, and aspartate at the active site) can cleave individual components of the ECM (Table IA, B). Although the emphasis on different proteinase activities varies according to the situation, the metalloproteinases appear to have the most ubiquitous role in matrix turnover. In this review, we will summarize the role of other proteinase types and describe the metalloproteinases in rather more detail.

Cathepsin D, a lysosomal enzyme [1], is the major aspartic proteinase involved in matrix degradation. It has optimal proteolytic activity at acid pH between 3 and 5, showing little activity at neutral pH. It can degrade proteoglycan core protein, gelatin, and collagen telopeptides, but not native triple helical collagen. A recent study showed that cathepsin D cleaves aggrecan within both interglobular and chondroitin sulfate domain over a pH range of 5.2–6.5 [2]. Although early reports indicated that cathepsin D might be involved in the extracellular degradation of matrix molecules, later studies have demonstrated that its role is confined to the intracellular breakdown of phagocytosed matrix molecules. For example, pepstatin, an inhibitor of cathepsin D, and anticathepsin D antiserum do not inhibit vitamin A-induced cartilage breakdown in vitro [3]; in rat osteoclasts, cathepsin D was immunolocalized in granules or vacuoles, but negligible staining was seen along resorption lacunae and in eroded bone matrix, in contrast to the staining pattern seen for other cathepsins [4].

Dynamics of Bone and Cartilage Metabolism

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Ian M. Clark and Gillian Murphy Table IA.

Proteolysis of the Extracellular Matrix

Proteinase class range Serine Plasmin Tissue plasminogen activator Urokinase-type plasminogen activator Neutrophil elastase Proteinase 3 Cathepsin G Plasma kallikrein Tissue kallikrein Tryptase Chymase Granzymes Cysteine Cathepsin B Cathepsin L Cathepsin S Cathepsin K Calpains Aspartate Cathepsin D

Matrix substrates

Inhibitors

pH

Fibrin, fibronectin, aggrecan pro MMPs Plasminogen Plasminogen Most ECM components pro MMPs Similar to elastase Aggrecan, elastin collagen II, pro MMPs pro UPA, pro MMP1, MMP3 pro MMP8 Collagen VI, fibronectin pro MMP3 Many ECM components, pro MMP1, MMP3 Aggrecan

α2 anti-plasmin PAI-1, PAI-2 PAI-1, PAI-2, PN-1 α1-PI α1-PI, elafin α1-PI Aprotinin Kallistatin Trypstatin α1-PI

Neutral Neutral Neutral Neutral Neutral Neutral Neutral Neutral Neutral Neutral

Collagen telopeptides, collagen IX, XI, aggrecan As cathepsin B, elastin As cathepsin B, elastin Collagen telopeptides, collagen, elastin Proteoglycan, fibronectin, vitronectin

Cystatins Cystatin Cystatin Cystatin Calpastatin

5.0–6.5 4.0–6.5 6.0–6.5 Neutral Neutral

Many phagocytosed ECM components

Pepstatin

3.0–6.0

III. CYSTEINE PROTEINASES The cysteine proteinases which degrade extracellular matrix components include cathepsins B, L, S, K, and calpains I and II. Cathepsins B, L, and S are lysosomal enzymes with acid pH optima; ultrastructural studies using the electron microscope have demonstrated their involvement in intracellular degradation of phagocytosed matrix molecules. However, they may also function as extracellular proteinases at pH closer to neutral. The lysosomal active form of cathepsin B is unstable at neutral pH, but both pro-cathepsin B and a higher Mr active form (which may be a noncovalent complex between cleaved propeptide and active enzyme) are secreted extracellularly from stimulated connective tissue cells and activated macrophages; these are more stable and possibly active at near neutral pH. A study of cathepsin B activity in synovial fluid demonstrated a higher level of activity in RA fluid than OA fluid, with 42 kDa pro-enzyme and 29 kDa active enzyme being present [5]. These data were recently replicated with the demonstration that the majority of cathepsin activity in synovial fluid was from cathepsin B [6]. Human osteoblasts are reported to secrete a form of cathepsin B which is stable at neutral pH and can be induced with IL-1 and PTH [7]. At acid pH in vitro, cathepsins B, L, and S cleave the telopeptides of types I and II collagen, causing depolymerization; they also cleave proteoglycan core protein, gelatin, and nonhelical regions of types IX and XI collagen [8, 9]. Cathepsins L and S are both potently elastolytic

and, at neutral pH, cathepsin S has elastolytic activity comparable with that of neutrophil elastase [9]. Cathepsin S is stable over a broader pH range than cathepsin B or L. If cathepsins B and L are added to cartilaginous matrix at near neutral pH, then proteoglycan and collagen are released [10]. Cathepsin B has been shown to be the major cysteine proteinase activity in OA chondrocytes and cartilage [11]. Indeed, cathepsin B has been shown to be increased in lysosomes of OA chondrocytes compared to normal, and also extracellularly at sites of cartilage degradation in OA [12]. A lipophilic cathepsin B inhibitor can block proteoglycan release induced by interleukin 1 in an explant model and this can also be blocked by MMP inhibitors, perhaps revealing a role for the cathepsin in the activation of proMMP [13]. In endochondral bone formation, it has been proposed that the complete degradation of type X collagen requires both the action of MMP-13 from the chondrocyte and then cathepsin B from the osteoclast [14]. In bone resorption by osteoclasts, studies with specific inhibitors in a murine calvarial assay suggest that cathepsins L and S might act both intracellularly and extracellularly, but that cathepsin B only acts in the intracellular compartment, possibly via activation of other proteinases [15]. In rat osteoclasts, cathepsin B and L immunostained along lacunae with strong extracellular staining of both on collagen fibrils and the bone matrix under ruffled border, with a stronger signal for cathepsin L than cathepsin B [4]. A further ultrastructural study using specific

183

Chapter 11 Matrix Proteinases Table IB. Proteolysis of the Extracellular Matrix MMP No

Mr enzyme

Mr latent

MMP-1

Interstitial collagenase-1

55 000

45 000

MMP-8

Neutrophil collagenase

75 000

58 000

MMP-13

Collagenase-3

60 000

48 000

MMP-18 MMP-3

Xenopus collagenase Stromelysin-1

55 000 57 000

? 45 000

MMP-10

Stromelysin-2

57 000

44 000

MMP-11 MMP-19

Stromelysin-3 RASI

51 000 ?

44 000 ?

MMP-20 MMP-2

Enamelysin gelatinase A

72 000

66 000

MMP-9

gelatinase B

92000

86 000

MMP-7

Matrilysin (PUMP-1)

28 000

19 000

MMP-26

Matrilysin 2

MMP-12

Macrophage

54 00

45 000/22 000

MMP-14

MT-1 MMP-66000

MMP-15

MT-2 MMP

72 000

MMP-16

MT-3MMP

64 000

MMP-17 MMP-24 MMP-25 MMP23A MMP23B MMP27 MMP28

MT-4MMPMT5 MMP MT6 MMP

Epilysin

Active

56 000

52 000

Known substrates Collagens I, II, III, VII, VIII, X; gelatin: aggrecan, versican, proteoglycan link protein; laminin; perlecan; nidogen, α1-proteinase inhibitor, α2- macroglobulin (α2M); fibrin; fibrinogen; pregnancy zone protein; ovostatin; myelin basic protein (MBP); IGFBP; proTNF; CXCL12; proIL1β; stromal cell-derived factor, SDF; L-Selectin; proMMP-2; proMMP-9 Collagens I, II, III, V, VII, VIII, X; gelatin; aggrecan; brevican; α2-M; α1-proteinase inhibitor; α2-antiplasmin; fibronectin; CXCL5; IGFBP; ADAM-TS-1; C1q Collagens I, II, III, IV, VI, IX, X, XIV; gelatin; aggrecan; perlecan; brevican; fibrillin; fibronectin tenascin; α2-M; plasminogen activator inhibitor-2; CXCL12; SDF; proMMP-9 Collagen; gelatin Collagens III, IV, V, VII, IX, X, XI; gelatin; aggrecan; versican, perlecan; proteoglycan link protein; fibronectin; laminin; elastin; nidogen; decorin; osteonectin, osteopontin; tenascin; vitronectin; fibrin; fibrinogen; MBP; L-selectin; E-cadherin; antithrombin-III; proteoglycan link protein; α2-M ovostatin; α1- proteinase inhibitor; proHB-EGF; proTNF; pro MMP-1; proMMP-7; proMMP-8; proMMP-9; proMMP-13; uPA Collagens III, IV, V; gelatin; casein; aggrecan; brevican; elastin; proteoglycan link protein; fibronectin; fibrinogen; α2-M; proMMP-1; proMMP-8; proMMP-9 α1-proteinase inhibitor; IGFBP; α2-M Collagens I, IV; gelatin, aggrecan, COMP, nidogen; fibronectin; laminin; tenascin Amelogenin; aggrecan; COMP; Collagens I, IV, V, VII, X, XI, XIV; gelatin; elastin; fibronectin; aggrecan; versican; proteoglycan link protein; laminin; nidogen; fibulins; vitronectin; osteonectin; fibrillin; MBP; α2-M; α1-proteinase inhibitor; C1q; CCL7; CXCL12; proTNF; latent TGFβ; FGFR1; proIL1β; proMMP-9; proMMP-13; ADAM TS1; plasminogen Collagens IV, V, VII, X, XIV; gelatin; elastin; aggrecan; versican; decorin; nidogen; proteoglycan link protein; fibronectin; vitronectin; osteonectin; MBP; α2-M; α1-proteinase inhibitor; C1q; CXCL1; CXCL4; CXCL7; CXCL 12; endothelin, galectin-3; IL8; proTNF; IL1β; IL-2Rα; latent TGFβ; plasminogen Collagens IV, X; gelatin; aggrecan; brevican; decorin; elastin; fibulins; osteonectin; osteopontin; tenascin; vitronectin; proteoglycan link protein; fibronectin; laminin; nidogen; MBP; fibrinogen; E-cadherin; FASligand; α1-proteinase inhibitor; proHB-EGF; proTNF; β4integrin; pro MMP-1; pro MMP-2; proMMP-9 Collagen IV; gelatin; fibronectin; fibrinogen; vitronectin; α1-proteinase inhibitor; proMMP9 Collagen IV; gelatin; aggrecan; elastin; fibronectin; fibrillin; nidogen; metalloelastase vitronectin; laminin; MBP; α1-proteinase inhibitor; proTNF; plasminogen Collagens I, II, III gelatin; aggrecan; nidogen; fibrillin; perlecan; fibrinogen; elastin; fibronectin; laminin B chain; vitronectin; tenascin; α2-M; α1-proteinase inhibitor; CD44; CXCL12; αVintegrin; tissue transglutaminase; proMMP-2; pro MMP-13; proTNFα Aggrecan, nidogen; fibronectin; laminin; perlecan; tenascin; transglutaminase; proMMP-2; ADAM TS Collagen III; gelatin; fibronectin; laminin; perlecan; tenascin; tissue transglutaminase; proMMP-2; ADAM TS1 Gelatin; fibrin; fibrinogen; tissue transglutaminase; proMMP2; proTNFα Gelatin; fibronectin; proMMP2 Collagen IV; gelatin; fibronectin; fibrinogen; fibrin; proMMP2

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inhibitors also suggested that cathepsin L played a key role in bone resorption by cultured osteoclasts [16]. More recently a new cysteine proteinase, cathepsin K (previously called cathepsin O or O2), has been discovered in osteoclasts. Cathepsin K has 56% sequence homology with cathepsin S; it has a pH optimum of 6–6.5 similar to cathepsin S, but has activity over a broader range, remaining largely active at neutral pH. It degrades elastin, collagen telopeptides, and gelatin better than the other cysteine proteinase cathepsins [9, 17]. Aside from its collagen telopeptidase activity, cathepsin K also cleaves both types I and II collagen within the triple helix [18, 19]. Cathepsin K is stabilized by interactions with chondroitin sulfates [20] and requires complex formation with chondroitin sulfate for collagenase activity [21, 22]. Inhibitors of cathepsin K have been shown to reduce bone resorption in vivo or in vitro [23] and an antisense approach blocks osteoclastic bone resorption in vitro [24]. It has been suggested that only cathepsin K, and not cathepsins B, L, or S, is abundantly expressed in human osteoclasts, and that this enzyme is responsible for osteoclastic bone resorption [25]. Interestingly, pycnodyostosis, an osteochondrodysplasia is caused by mutation in the cathepsin K gene [26]. Cathepsin K is also expressed in synovial fibroblasts and chondrocytes and is raised in both OA and RA [27–29]. A family of cysteine proteinase inhibitors, the cystatins, has also been described [30]. Their role in controlling the turnover of cartilage and bone is largely unknown; however, cystatin C is reported to inhibit bone resorption in different in vitro systems due to inhibition of osteoclastic proteolytic enzymes and to be produced by bone cells [31]. Calpains are cytosolic, Ca2+-dependent, cysteine proteinases with neutral pH optima. Calpains 1 and 2 require micromolar and millimolar Ca2+ respectively for activation. These enzymes can degrade proteoglycan, fibronectin, and vitronectin [32]. Calpains have been shown to be raised in RA and OA synovial fluid and tissues [33, 34] and in some animal models of arthritis [35]. Recently, Calpain 2 has been shown to cleave aggrecan at a distinct C-terminal site consistent with the presence of such cleaved species in cartilage [36]. Calpastatin, a specific cellular inhibitor of calpains has been identified as an autoantigen in some RA patients [37].

IV. SERINE PROTEINASES A. Leukocyte Enzymes Neutrophil elastase, cathepsin G, and proteinase 3 are found in the azurophil granules of polymorphonuclear leukocytes. They are synthesized as precursors in

promyelocytes of bone marrow during development [38]. Granule contents may be discharged into the extracellular space during inflammation where they have the opportunity to degrade matrix. These enzymes have pH optima between 7 and 9 and can degrade many components of the matrix. Neutrophil elastase and proteinase 3 are both potently elastolytic; neutrophil elastase and cathepsin G degrade the telopeptides of fibrillar collagens and type IV collagen; all three enzymes can degrade other matrix glycoproteins, gelatin, and proteoglycan core protein [39–41]. Human neutrophil elastase has also been shown to cleave within the collagen triple helix [42]. Proteinase 3 is an autoantigen in Wegener’s granulomatosis [43]. The role of these enzymes in cartilage degradation may only be in specialized situations where they can escape inhibition by synovial fluid inhibitors (see below), for example during joint sepsis. Beige mice lack both PMN elastase and cathepsin G, but still undergo severe cartilage loss during antigen-induced arthritis [44]; rabbits depleted of neutrophils by nitrogen mustard show the same rate of cartilage degradation in arthritis models as the nontreated animals [45]. A number of inhibitors of these enzymes exist, including the general proteinase inhibitor α2-macroglobulin. α1 proteinase inhibitor is the major specific serum inhibitor of PMN elastase and also inhibits proteinase 3 [46]; α1 antichymotrypsin and secreted leukocyte proteinase inhibitor (SLPI) inhibit both PMN elastase and cathepsin G; SLPI has been isolated from cartilage [47]. Thrombospondin 1, a matrix protein, is also a potent inhibitor of PMN elastase and cathepsin G; this molecule is found to be induced in RA synovium proportionally to the numbers of surrounding leukocytes [48]. Cross-talk may exist between proteinase families via action on proteinase inhibitors, e.g. serpins may be inactivated by MMPs and neutrophil elastase may inactivate TIMPs [49, 50].

B. Plasminogen Activators and Plasmin The conversion of plasminogen (Pg, 90 kDa) to plasmin by plasminogen activators (PA) is a key step in connective tissue remodeling since plasmin can degrade a number of matrix macromolecules (aside from fibrin, plasmin can degrade the core protein of proteoglycan, fibronectin, laminin, and type IV collagen) [51] and also take part in the activation of proMMPs (e.g. proMMP-1 and -3). There are two types of PA, tissue-type (tPA) and urokinase-type (uPA). tPA (70 kDa) is mainly associated with endothelial cells, but is also produced by fibroblasts and chondrocytes; it also binds to extracellular matrix components

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such as laminin, fibronectin, and thrombospondin [52]. tPA is the major activator of Pg in fibrinolysis. uPA (54 kDa, two subunits of 30 kDa and 24 kDa which are disulfide bonded) is produced by many connective tissue cells. It binds a specific cell surface receptor (uPAR) via a growth factor domain in uPA, and this enhances activation of cellassociated Pg and protects the uPA from inhibition; uPA may also have mitogenic effects via this receptor [53]. Inhibitors of plasmin and PA come from the serpin superfamily [46]. α2-antiplasmin is the major plasmin inhibitor in plasma. α1-proteinase inhibitor and α2-antichymotrypsin are also active against plasmin [54]. PA inhibitors (PAIs) control the rate of plasmin production in the local extracellular environment. PAI-1, originally isolated from endothelial cells, is produced by a number of cell types and is present in platelets; it inhibits both tPA and uPA, but forms more stable complexes with the former [55]. PAI-1 binds to the pericellular matrix which enhances its extracellular stability and therefore may inhibit proteolysis in this environment. PAI-2 is a better inhibitor of uPA than tPA; it is present in plasma and is expressed by a number of cell types including macrophages [56]. PAI-3 is also an inhibitor of activated protein C, and is less well characterized [57]. Finally, protease nexin 1 (PN-1) inhibits uPA and tPA with the PN–1 proteinase complex binding a specific cell surface receptor via the inhibitor, followed by internalization and degradation [58]. In synovial fluid, uPA, PAI-1, and uPAR are all raised in RA compared to OA, and in OA compared to normal, though there was no direct association with clinical parameters [59]. There is also evidence for local production of uPA and PAI-1, with synovial fluid levels raised compared to plasma. At the invasive pannus front in RA synovium, uPA, uPAR, PAI-1, and PAI-2 are all induced compared to normal tissue [60, 61], whilst tPA is repressed [61]. In human OA and animal models of OA where enhanced bone remodeling may trigger cartilage damage, the uPA/plasmin system is up-regulated [62]. Plasminogen added to chondrocyte cultures causes an increase in interleukin-1-induced matrix degradation, implicating PA in this process; indeed, IL-1 induces uPA and PAI-1 in these cells. Similarly, plasminogen added to IL-1-stimulated cartilage explants potentiates increased collagen and proteoglycan loss, and this can be blocked with PA inhibitors or MMP inhibitors. Hence, there may be a cascade of enzyme activation with PA activating plasminogen, and plasmin activating MMPs [63–65]. Furthermore, intra-articular injection of protease nexin-1 blocks IL-1 or FGF-induced proteoglycan loss from rabbit knee and tranexamic acid (TEA), another antiplasmin agent, used orally, has the same effect [66]. However, a

recent pilot-scale double-blind placebo-controlled trial of TEA in RA patients showed no therapeutic efficacy [67]. In a murine model of mild arthritis (antigen-induced arthritis), uPA- or tPA-null mice have increased disease severity [68] whilst PAI-1 null mice have decreased disease severity [69]; disease severity also correlates with fibrin deposition. However, in a systemic and more aggressive model (collagen-induced arthritis), uPA-null mice develop less severe disease whilst tPA-null mice develop more severe disease. Disease severity still correlates with fibrin deposition, with the uPA-null mice also having defects in T-cell responses [70]. Osteoblasts also produce uPA, tPA, PAI-1, and uPAR, and their synthesis is regulated by osteotropic hormones [71, 72]; it has been postulated that they function as activators of proMMPs or latent growth factors in bone remodeling, or that uPA may be acting as a mitogen via its growth factor domain. Similarly, the direct resorptive activity of osteoclasts does not require tPA or uPA, but the latter may be involved in migration of the pre-osteoclasts to the mineral surface [73]. It has been proposed that this enzyme system is key to the initiation of bone resorption [74]. In mice lacking plasminogen activators increased bone formation is observed [75].

C. Mast Cell Proteinases Mast cell granules contain tryptase, chymase, and cathepsin G. Tryptase has a trypsin-like specificity and degrades collagen type VI and fibronectin. Chymase has a chymotrypsin-like specificity and degrades a number of matrix substrates. Both enzymes have been implicated in the activation of proMMPs [76]. Activated, degranulated mast cells have been localized to regions of cartilage erosion in RA [77].

D. Granzymes Granzymes A and B are serine proteinases which are stored in the granules of activated cytotoxic T-cells and NK cells. Granzyme B has been reported to degrade aggrecan in the extracellular matrix synthesized by chondrocytes [78, 79] and recently this enzyme has been reported to be expressed by chondrocytes themselves [80]. Granzymes A- and B-expressing lymphocytes have been localized in the synovium of patients with RA and OA, and may be increased in RA tissue [81]. Granzymes A and B are elevated in the plasma and synovial fluid of patients with RA compared to OA or reactive arthritis [82].

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V. METALLOPROTEINASES At least three classes of zinc-dependent endopeptidase activity have been implicated in the degradation of cellsurface molecules and the ECM [83]: the matrix metalloproteinase (MMP) family, the reprolysin-related proteinases (ADAMs or MDCs) that contain both a metalloproteinase and a disintegrin domain, and the astacin-related proteinases. The astacins discovered to date include the enzyme that cleaves the carboxy-terminal propeptides of the fibrillar collagens during collagen assembly (bone morphogenetic protein-1; [84]) and others involved in cell differentiation and pattern formation during development. Recent studies have provided direct morphological and functional links between its proteolytic activity and the ECM [85]. The ADAM family and a related ADAMTS family are discussed in more detail below.

A. ADAM Family The ADAMs (a disintegrin and metalloproteinase) are a family of 21 human proteinases involved in numerous physiological and developmental processes. They each contain an amino-terminal signal sequence, a pro-domain, a metalloproteinase domain, a disintegrin domain with homology to the snake venom disintegrins, a cysteinerich domain, an EGF-like region, and transmembrane and cytoplasmic domains. Not all of the ADAMs contain a complete metalloproteinase active site consensus, suggesting that only 13 are catalytically active. The ADAMs were initially described as having roles in fertilization, but have now been implicated in cell migration, differentiation, activation of signaling pathways, and shedding of cell-surface molecules [86]. Levels of soluble TNFα are significantly elevated in RA, largely due to the action of ADAM-17, TNFα-converting enzyme (TACE). Expression of TACE is predominantly localized to the synovium in RA [87]. Both TNFα and TACE can also be expressed in cartilage chondrocytes [87–89]. There have been several reports of mRNA for ADAM 9, ADAM 10, ADAM 12, and ADAM 15 being expressed in chondrocytes, with ADAM 10 and ADAM 15 being up-regulated in OA cartilage compared to normal [90–94]. Inoue et al. [95] have shown that ADAMs 11, 12, 15, and 19 are expressed by osteoblasts and ADAMs 11 and 15 by osteoclasts. Other studies suggest a function for ADAM 10 and ADAM 12S in the formation of osteoclasts from human peripheral blood mononuclear cells and subsequent resorption activity [96]. These proteins have proposed functions in the proteolytic processing of cell-surface

ectodomains, in membrane fusion events, and in cell–cell and cell–matrix interactions [97].

B. ADAMTS Family The ADAMTS family are a recently described group of secreted proteinases which, like ADAMs, contain a signal sequence, pro-domain, metalloproteinase domain and disintegrin-like region, but this is followed by a thrombospondin-1 repeat, then a cysteine-rich domain, one or more additional thrombospondin-1 repeats and in some cases an additional C-terminal domain often containing a PLAC motif [98]. There are currently 19 human family members including six enzymes reported to degrade aggrecan (ADAMTS 1, ADAMTS 4, ADAMTS 5, ADAMTS 8, ADAMTS 9, ADAMTS 15) and these are termed aggrecanases [98, 99]. Cleavage of aggrecan in the G1–G2 interglobular domain is believed to be a key event in aggrecan degradation and this region contains cleavage sites for both MMPs (Asn341-Phe342) and aggrecanases (Glu373–Ala374). The major aggrecan fragments released from resorbing cartilage are not products of MMP action [100]. Using neoepitope antibodies to study cleavage products of both enzyme families in cartilage, it appears that aggrecanases may be active early in the arthritic disease process with subsequent increases in MMP activity [101–103]. ADAMTS 4 and ADAMTS 5 also cleave the chondroitin sulfate proteoglycans brevican and versican [104, 105]. The most studied aggrecanase enzymes are ADAMTS 4 and ADAMTS 5. Western blot analysis has demonstrated expression of ADAMTS 4 and ADAMTS 5 in normal bovine cartilage explants, human osteoarthritic cartilage, and rheumatoid, osteoarthritic and normal synovium. In cartilage explant cultures aggrecanase activity can be induced by the addition of catabolic cytokines. Levels of ADAMTS 4 are induced in cartilage in response to IL-1 and TNFα and in fibroblast-like synoviocytes in response to TGFβ [106–109]. ADAMTS5 expression tends to be less responsive to stimulation, though this is dependent on cell type [107, 109]. Immunodepletion of ADAMTS4 from bovine articular cartilage cultures following IL-1 stimulation resulted in a 75% reduction of aggrecanase activity in the culture medium, demonstrating that ADAMTS4 is the major aggrecanase in this model, presuming the antibody is monospecific [106]. In addition to cleavage at the Glu373–Ala374 bond, at least ADAMTS4 and ADAMTS5 cleave aggrecan at four other sites within the chondroitin sulfate-rich region of the core protein between G2 and G3. Cleavage of these sites in vitro appears to be more efficient than cleavage

Chapter 11 Matrix Proteinases

within the G1–G2 IGD [106, 110, 111]. Using recombinant ADAMTS4, the full-length active form of the enzyme was an effective aggrecanase, but exhibited little activity against the Glu373–Ala374 bond, preferentially cleaving at Glu1480–Gly1481. However, deletion of the spacer region led to a reduction in aggrecanase activity, but a markedly increased cleavage of Glu373–Ala374; this C-terminally truncated form of the enzyme also diminishes binding to extracellular matrix and may be the primary form found in, for example, interleukin-1-stimulated cartilage [112]. In the cell, C-terminal processing of ADAMTS4 appears to be mediated by MT4-MMP [113]. The presence of the TSP1 domain in ADAMTS4 and the GAG substitutions in aggrecan are also important for substrate recognition [111, 114]. Despite a growing understanding of the biochemistry and expression of the known aggrecanases, the relative contribution of each enzyme in cartilage destruction remains to be determined.

C. Matrix Metalloproteinases The MMPs are key ECM-degrading proteinases and have been divided into subgroups according to their structure and function. To date 23 individual human enzymes have been described (Table IB). The MMPs have a common domain structure with a signal peptide, a propeptide, a catalytic domain, a hinge region and a hemopexinlike C-terminal domain. Individual MMPs have variations on this general structure: the MT-MMPs have a transmembrane domain and cytoplasmic tail (MMP14–16 and 24) at the C-terminal end or are GPI-anchored (MMP17 and 25) and, in common with MMP11, 21, 23, and 28, have a sequence containing a potential furin-cleavage site at the N-terminal end of the catalytic domain. Matrilysins (MMP7 and 26) lack the C-terminal domain and hinge region, the gelatinases A and B (MMP2 and 9) have an insert of three fibronectin type II repeats in the catalytic domain; and MMP9 has a collagen-like sequence at the C-terminal end of the catalytic domain. MMP23 occurs in two forms, A and B, encoded by separate genes. These are type II transmembrane proteins containing a unique cysteine array and an immunoglobulin-like domain at the C-terminus [115]. The collagenases-1, -2, and -3 (MMP1, MMP8, and MMP13, respectively) have the specific ability to degrade fibrillar collagens within the native triple helix. They also have limited ability to cleave other proteins, MMP13 being the most active, with significant activity against type IV collagen and denatured collagens (gelatins). The catalytic domains of collagenases alone retain proteolytic activity

187 but they do not digest native triple helical collagen. MMP2 and MMP9 actively degrade denatured collagens and MMP2 has low but significant activity on soluble native fibrillar collagens. They also degrade soluble type IV collagen but activity on more native preparations is extremely weak [116]. Both enzymes degrade type V collagen, elastin, aggrecan, and other matrix proteins to a limited extent. Some of the MT MMPs have collagenolytic activity (see below). The stromelysin subgroup, MMP3 and MMP10 cleave a number of matrix components, notably aggrecan, fibronectin, laminin, and type IV collagen. MMP3 has limited activity against collagens III, IX, and X and the telopeptides of collagens I, II, and XI [117]. Stromelysin 3, MMP11, has some of the substrate specificity of the other stromelysins but its activity is so weak that its role as a proteinase may be against a specific, as yet unidentified, substrate. Matrilysin, MMP7, and metalloelastase, MMP12, are also somewhat similar to the stromelysins, but are rather more active. The more recently discovered MT MMPs, MT 1–3 (MMP14–16), are also known to cleave a number of ECM proteins, including fibrillar collagens. The other newer MMPs also degrade some matrix proteins (Table IB). The catalytic domains of the MMPs are spherical or ellipsoidal and have an active site cleft containing a catalytic zinc ion at the bottom. The structure is stabilized by a second zinc ion and a calcium ion. Crystal structures have been obtained for the catalytic domains of human MMP1 [118–120], human MMP3 [121], human MMP8 [122–124], and human MMP9 [125] complexed with synthetic inhibitors. The three-dimensional structures of the catalytic domain of several human MMPs have also been determined by NMR and used in the design of lowmolecular-weight inhibitors [126]. Some of the differences in substrate specificity can be accounted for by variations in the six specificity subsites in the active site cleft and in sequences around the entrance to the cleft. The S1′ pocket (the first specificity subsite on the carboxy-terminal side of the substrate scissile bond) is deeper in MMP3 [121, 127] and larger in MMP8 [122, 124] than in MMP1. The type II fibronectin-like repeats that form the gelatin-binding domain of MMP2 occur as a three pronged “fish hook” to the side of the active site cleft, conferring specificity on the catalytic domain [128]. The hinge region varies in length between the different MMPs. The crystal structure of porcine MMP1 showed the hinge to be highly exposed and with no secondary structure [129]. In the collagenases this region is prolinerich, and has been suggested to mimic the triple helix of collagen and to contribute to the binding of MMP8 to type I collagen [130]. It has been shown that collagenases

188 bind to and locally unwind triple helical collagens. MMP1 preferentially interacts with the α2(I) chain of type I collagen, by both catalytic domain and hemopexin domain interactions and cleaves the three α chains in succession and the flexible hinge may be critical to this mechanism, coordinating the binding of the domains to the collagen [131]. The crystal structures of full-length porcine MMP1 [129], the C-terminal domain of human MMP13 [132], and human proMMP2 [128, 133] showed their hemopexinlike domains to comprise four similar units that form a four-bladed β-propeller structure. This propeller is stabilized by a disulfide bridge between the first and fourth units and has cations, postulated to be calcium, and chloride ions at the core. The whole structure has a disk-like shape with a funnel-like tunnel running through the short axis [132, 133]. On MMP2 there is a surface patch of positive residues extending from the exit side of the tunnel across propeller blade III that has been proposed as an anchoring point for TIMP-2 [133] and this was confirmed by the proMMP2-TIMP-2 crystal structure [134]. The hemopexin-like domain contributes towards the specificity of the collagenases. Studies on the isolated N-terminal domain of MMP1 [135], on truncated mutants of human collagenases and on collagenase/stromelysin-1 chimeric enzymes, have shown that the ability of collagenase-1 [136] and MMP8 [137] to cleave native type I collagen is influenced by the C-terminal domain. This domain is also involved in the binding of active MMPs to TIMPs [135, 136, 138, 139] and in the binding of latent and active MMP3 [136] and MMP8 [140] to extracellular matrix molecules. The C-terminal domains of MMPs 2 and 9 allow them to form complexes with certain of the TIMPs when latent. This is the basis for the cellular activation of proMMP2 by MT-1 MMP, involving the formation of a multiprotein cell-surface cluster where TIMP-2 tethers proMMP2 by its hemopexin domain to MT1 MMP, facilitating proteolysis of the propeptide by free MT1 MMP and active MMP2 [141]. The genes encoding for example MMP1, MMP3, MMP7, MMP9, and MMP10, have a TATA box and a TPA-responsive element (TRE), a sequence that binds to Fos and Jun (AP1), which is thought to mediate the induction of these MMPs by inflammatory cytokines such as IL-1 and TNF-α. MMP2 appears to be constitutively produced by cells in culture and is not generally significantly modulated by cytokines. The 5′-flanking region has no TATA box or identifiable TRE sequence, but does contain a p53-sensitive sequence and an adenovirus ElAresponsive element resembling the AP2-binding site and two “silencer” regions [142]. Glucocorticoids, retinoids, and TGFβ repress the expression of at least MMP1 and MMP3 in connective tissue fibroblasts [143, 144].

Ian M. Clark and Gillian Murphy

Agents may induce the expression of one MMP, whilst repressing expression of another. Furthermore, the effect of any one agent may differ with cell type; this makes it difficult to predict the effects of blocking cytokine/growth factor action. Table II shows some of the factors which modulate expression of the MMP-1 gene. The extracellular regulation of MMP activity is a key feature of their function. The regulation of zymogen activation is possibly the most critical level of control of enzyme activity. The MMPs have latency conferred on them by their propeptide domain, which effectively blocks the active site in the catalytic domain. In the latent enzyme, a cysteine residue in the N-terminal propeptide PRCGVPDV sequence co-ordinates with the catalytic zinc ion, preventing water from occupying this site. In the active enzyme, water co-ordinated with this zinc ion is important in the mechanism of peptide bond hydrolysis. Upon enzymic cleavage in the prodomain the Zn2+–cysteine complex becomes destabilized, and a conformational

Table II. Gene MMP-1

TIMP-1

TIMP-2 TIMP-3

Factors that Modulate Expression of MMP and TIMP Genes Inducing agent

Interleukin-1 Tumor necrosis factor α Epidermal growth factor Basic fibroblast growth factor Platelet-derived growth factor Lipopolysaccharide Phorbol ester Colchicine Cytochalasin B Concanavalin A Calcium pyrophosphate Calcium ionophore A23187 UV irradiation Transforming growth factor β Interleukin-6 Interleukin-11 Oncostatin M Leukemia inhibitory factor Basic fibroblast growth factor Epidermal growth factor Interleukin-1 Progesterone Oestrogen Retinoic acid Phorbol ester Lipopolysaccharide Progesterone Transforming growth factor β Epidermal growth factor Platelet-derived growth factor Phorbol ester Dexamethasone

Repressing agent Transforming growth factor β Retinoic acid Estrogen Progesterone Interferon γ Dexamethasone

Dexamethasone Concanavalin A

Transforming growth factor β Lipopolysaccharide Oncostatin M

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change results in autoproteolysis of the entire propeptide [145]. Most of the MMPs are secreted in their latent forms and activation takes place outside the cell. The MMPs with a furin cleavage site in the propeptide (MMP11, 21, 23, 28) are exceptions to this, and are activated intracellularly by furin, or other proprotein convertases, prior to secretion [146]. The MT-MMPs have a similar furincleavage site (RXKR) and are also likely to be activated before leaving the cell. In vitro the MMPs can be activated by chemical agents that modify cysteine residues, such as the organomercurial 4-aminophenymercuric acetate (APMA). Activation may also occur when the conformation of the proenzyme is perturbed, as is evident from the SDS-activation of proMMP2 and proMMP9 during zymography. In vivo, however, activation probably takes place upon the enzymic cleavage of the propeptide. This cleavage may be an autolytic step, as occurs with proMMP2 at high concentrations in vitro [147], but more usually requires a second proteolytic activity. As can be seen from Table III, many of the MMPs share common activators and many are activated by other MMPs. This has given rise to speculation that the MMPs are involved in activation cascades [148]. An important cascade is initiated by the action of plasminogen activator (tPA or uPA) on plasminogen, which results in the active enzyme plasmin. This enzyme can activate several MMPs, as depicted in Figure 1. MMP2 is not involved in the plasmin cascade, and appears to be regulated differently to the plasmin-activated MMPs. It is not cleaved by trypsin, chymotrypsin, plasmin, thrombin, Table III. Human MMP

MMP No.

Collagenase-1 Collagenase-2 Collagenase-3 Stromelysins-1, -2 Stromelysin-3 Matrilysin Metalloelastase Gelatinase A

1 8 13 3,10 11 7 12 2

Gelatinase B MT1-MMP MT2-MMP MT3-MMP MT4-MMP MT5-MMP MT6-MMP MMP-19 MMP-20 MMP-23 MMP-26 MMP-28

9 14 15 16 17 24 25 19 20

elastase, cathepsin G, plasma kallikrein, or MMP3 [149, 150]. The mechanism involving MT MMPs described above appears to be the most likely pathway for MMP2 activation and may lead to the activation of MMP13 and MMP9 [151].

D. Tissue Inhibitors of Metalloproteinases There is a group of specific inhibitors of the MMPs called the TIMPs (tissue inhibitors of metalloproteinases; [152]), Four TIMPs have been described to date; TIMP-1 [153], a 28-kDa glycosylated protein; TIMP-2 [154], a 22-kDa nonglycosylated protein; TIMP-3 [155–161], a 21–27-kDa protein that binds to the extracellular matrix [144,148]; and TIMP-4 [162, 163], the human form of which was cloned from a cDNA library and expressed as a 22-kDa recombinant protein [162]. TIMP-4 is more closely related to TIMPs-2 and -3 than to TIMP-1 [163]. The TIMPs have 12 conserved cysteines, giving a protein structure of six loops which can be divided into two subdomains [164]. TIMP-1 has the shape of an elongated wedge in which the long edge, consisting of five different chains, occupies the active site cleft of MMP3 in the complex and are responsible for around 75% of the interactions between TIMP and MMP3. The central disulfide-linked segments Cysl–Val4 and Ser68–Val69 bind either side of the MMP catalytic zinc, with ligation of the zinc by the carbonyl oxygen and α amino groups of Cys1 [115]. Although they are able to inhibit most

Activation of MMPs

Exogenous activators Plasmin, kallikrein, chymase, MMP-3 Plasmin, MMP-3, MMP-10 Plasmin, MMP-2, MMP-3, MMP-14 Plasmin, kallikrein, chymase, tryptase, elastase, cathepsin G Furin Plasmin, MMP-3 Plasmin MMP-1, MMP-2, MMP-7, MMP-9, MMP-13, MMP-14, MMP-15, MMP-16, MMP-17, MMP-24, MMP-25, elastase, cathepsin G Plasmin, neutrophil elastase, MMP-3, MMP-2, MMP-13, MMP-26 Furin Probably a proprotein convertase Probably a proprotein convertase Probably a proprotein convertase Probably a proprotein convertase Probably a proprotein convertase Not known MMP-14 Probably a proprotein convertase MMP-26 Probably a proprotein convertase

Activating MMP-2 Not known MMP-2, MMP-9 MMP-1, MMP-8, MMP-9, MMP-13 Not known MMP-1, MMP-2, MMP-3, MMP-9 Not known MMP-1, MMP-2, MMP-9, MMP-13 Not known MMP-2, MMP-13 MMP-2 MMP-2 ADAMTS-4 (C-terminal processing) MMP-2 MMP-2 Not known Not known Not known MMP-9 Not known

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Figure 1

A scheme for the cellular matrix metalloproteinase activation cascades that occur pericellularly. The activation of proMMPs is largely limited to the pericellular environment where cell-associated proteinases can function in a privileged environment away from an excess of proteinase inhibitors. Key initiators of the MMP activation cascades are thought to be MT-MMPs and plasmin-mediated proteolysis. The generation of partially active or active MMPs allows a cascade of cleavages to generate fully active enzymes. The efficiency of these interactions is dependent upon mechanisms for the concentration of MMPs at the cell surface or on the extracellular matrix. (Courtesy of Dr V. Knäuper, Dept. of Biology, University of York, UK.)

active MMPs, TIMP-1 binds very poorly to many MT MMPs and MMP 19 [152]. In addition to their ability to bind to active MMPs the TIMPs can bind to certain latent MMPs. TIMP-1 and -3 bind to pro-MMP9 [165, 166] and TIMPs-2, -3, and -4 bind to pro-MMP2 [138, 167–172]. The C-terminal loops of TIMP-1 are involved in the binding of this inhibitor to proMMP9 [170]. Similarly, the C-terminal domain of TIMP-2 is essential for binding to pro-MMP2 which occurs through interactions between a negatively charged “tail” on TIMP-2 [172] and a positively charged area on the hemopexin domain of MMP2 [134]. Since pro-MMP2 complexed to TIMP-2 is less prone to autoactivation, it was suggested that TIMP-2 acts to stabilize the proenzyme [173–175]; however, more recent work has implicated TIMP-2 involvement in the activation of pro-MMP2 by cellular MT1 MMP [141, 176]. Pro-MMP9 complexed to TIMP-1 resists activation by MMP3 [166] but can be activated by organomercurial compounds, albeit more slowly than noncomplexed pro-MMP9 [165, 177, 178]. TIMP-3, which appears to be sequestered in the ECM, is a good inhibitor of ADAM17 and may also regulate other ADAMs [152]. It is also the most effective inhibitor of ADAMTS enzymes [179], and is likely to be an endogenous inhibitor of aggrecanases.

TIMP-1 is produced by many cell types and can also be up-regulated by a variety of factors such as serum, bFGF, EGF, TGFβ, IL-6 family members, retinoids, progesterone, and phorbol ester; expression is repressed by dexamethasone. In murine embryonic development, TIMP-1 is expressed at sites of osteogenesis in the limbs, ribs, digits, skull, and vertebrae [180]. TIMP-2 is often constitutively produced by cells and does not generally show a response to actions of cytokines, however, where expression is regulated, it is often in the opposite direction to TIMP-1 and -3, e.g. expression of TIMP-2 is repressed by TGFβ in various cell types. TIMP-3 has been shown to be induced by factors such as EGF, PDGF, TGFβ, phorbol ester, and dexamethasone [180]. TIMP-4 exhibits the most tissuerestricted expression pattern of the TIMPs, being expressed predominantly in heart and brain [152]. Table II shows some of the factors which regulate TIMP gene expression. The TIMP-1, -2, and -3 promoters have potential Sp1binding sites and share other features which are suggestive of housekeeping genes; however, TIMP-1 and -3 are known to be highly stimulus-responsive. The TIMP-1 promoter has an AP-1 and PEA3 motif in close proximity reminiscent of the inducible MMP genes described above; this gene also appears to have important regulatory sequences

Chapter 11 Matrix Proteinases

downstream of the transcription start sites. The murine TIMP-3 gene has six upstream AP-1 sites, which may be involved in basal expression of the gene, although data from the murine and human genes are in conflict here: the murine gene also has a putative p53-binding site, but this appears to be nonfunctional. The TIMP-2 gene has a promoter proximal AP-1 motif, but this appears not to have a major role in basal expression of the gene [180]. The TIMP-4 gene promoter has at least a functional CCAAT and Sp1 site [181].

E. RECK MMP activity can also be inhibited by a recently identified GPI-anchored inhibitor termed RECK (reversioninducing cysteine-rich protein with Kazal motifs) [182]. RECK was shown to inhibit the activity of MMP-2, -9, and -14, though it is unclear how this occurs at the molecular level.

F. α2-Macroglobulin α2-macroglobulin (α2M) is a plasma proteinase inhibitor which is able to inhibit almost all proteinases including MMPs. Human α2M is a 720 kDa tetrameric protein consisting of four 180 kDa subunits. Each subunit contains an exposed “bait” region, a sequence which can be cleaved by virtually all proteinases. Human MMP1 cleaves human α2-macroglobulin at the Gly–Leu bond in the bait region sequence –Gly–Pro–Glu–Gly679–Leu–Arg–Val–Gly–, which strongly resembles the collagenase-1 cleavage site in calf skin collagen α1(I) and α2(I) chains [183]. This cleavage leads to a conformational change in the α2M which traps the proteinase to the exclusion of large substrates; small (peptide) substrates may still be cleaved. ADAMTS4 and ADAMTS5 cleave α2M at Met690–Gly691, which represents a novel bond cleaved for these enzymes [184]. Inhibition by α2-macroglobulin is irreversible. α2M–proteinase complexes are cleared from the circulation by a specific receptor which is a low-density lipoprotein receptor-related protein. Aside from its function as a proteinase inhibitor, α2M has been shown to bind a variety of cytokines such as PDGF, TGF , bFGF, and IL-1β whence it may act as a carrier protein [185, 186]. Interestingly, α2-macroglobulin can inhibit urokinase-type plasminogen activator (uPA) when it is in solution but not when it is bound to uPA receptors at the cell surface, probably due to steric hindrance preventing the engulfing of the uPA by the α2-macroglobulin [187]. The presence of this inhibitor thus localizes uPA activity to the cell surface.

191

G. Metalloproteinases and TIMPs in Cartilage and Bone Extracts of cartilage from osteoarthritic patients show increased metalloproteinase activity compared to normal cartilage [188]. An imbalance of MMP and TIMP activity is implicated in cartilage loss. Recent expression-profiling studies have examined the expression of all MMPs, ADAMTSs, and TIMPs at the mRNA level in cartilage [189, 190]. These studies have unequivocally shown an increase in MMP13 expression in osteoarthritic cartilage. Interestingly, MMP3 expression appears to be strongly reduced in osteoarthritic cartilage compared to normal tissue. Aggrecanases from the ADAMTS family also tend to show lower expression in this end-stage OA cartilage but this may reflect their potential role early in the disease process [189, 190]. MMPs are raised in the synovium and cartilage of RA patients [191]. In the septic joint, high levels of MMP–TIMP complexes have been found in the synovial fluid, with active MMPs present, but no free TIMPs; rapid cartilage destruction is seen in these patients [192]. Control over MMP activity in vivo occurs at different levels and involves factors such as regulation of gene expression, activation of zymogens, and inhibition of active enzymes by specific inhibitors. The importance of activation of proMMPs in cartilage destruction has recently been underlined by the demonstration that cartilage collagen release in an explant model does not take place until activation occurs, even if the expression of procollagenases is induced [65]; furthermore, the overexpression of interleukin-4 prevents MMP-mediated cartilage erosion in a murine model by interfering with proMMP activation [193]. This pivotal role for activation of proMMPs may explain data from both murine immune complex arthritis, and also antigen-induced arthritis where MMP-3-deficient animals show no difference in proteoglycan depletion to wild-type, but fail to degrade collagen (and hence do not develop severe cartilage erosions) with evidence of failure to activate procollagenases [194, 195]. Growth factors such as IL-1, TNFα, bFGF, PDGF, OSM, and ATRA stimulate MMP expression in cartilage and also induce cartilage resorption; factors such as TGFβ, IGF-I, IL-4, IL-10, and IL-13 may oppose these effects. TIMP-1 and -2 have been shown to prevent collagen release in an ex vivo cartilage resorption model, underlining the involvement of collagenase MMPs in this event [196]. TIMP-3 (but not TIMP-1 or TIMP-2) blocks proteoglycan release in a similar model, again highlighting the role of aggrecanase ADAMTSs in this explant system [197]. MMPs have been implicated in bone turnover, either via removal of osteoid to allow osteoclastic bone resorption or more directly as osteoclast enzymes.

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In fetal rat calvarial osteoblasts, a variety of growth factors have been shown to alter expression of collagenase (MMP-13) and also to modulate bone resorption in calvarial explant assays; inducing agents include PTH, bFGF, ATRA, PDGF, and IL-6; suppressing factors include IGF-I and -II, TGFβ, and BMP-2. Some of these factors (bFGF, TGFβ, and BMP-2) also induce expression of TIMP-1 and -3 [198]. Human osteoblasts have been reported to express at least MMP-1, -2, -3, -9, -10, -13, and -14 [199]. Synthetic MMP inhibitors have been demonstrated to suppress bone resorption [200]. In human development, MMP-13 is expressed in mineralizing skeletal tissue, in hypertrophic chondrocytes and osteoblasts involved in ossification; in osteoblasts, MMP14 and MMP-2 were co-localized with MMP-13. In postnatal tissues, MMP-13 is expressed at sites of remodeling such as bone cysts and ectopic bone and cartilage formation, whilst in RA patients strong expression is seen in cartilage. MMP-13 was therefore proposed to function in the degradation of type II collagen in primary ossification, skeletal remodeling and joint disease [201]. Similar patterns of expression are seen in developing rat and mouse bone; rat osteoblasts possess a scavenger receptor which removes MMP-13 from the extracellular space [198]. Expression of MMPs has also been noted in osteoclasts, particularly MMP-9, but also MMP-1, -2, -3, -13, and -14 [202]; indeed, some inhibition of bone resorption by MMP inhibitors has been shown even on osteoid-free bone [200].

References 1. Dunn, B. M. (1990). Structure and function of the aspartic proteinases. Genetics, structures and mechanisms. Adv. Exp. Med. Biol. 306, 1–585. 2. Handley, C. J., Mok, M. T., Ilic, M. Z., Adcocks, C., Buttle, D. J., and Robinson, H. C. (2001). Cathepsin D cleaves aggrecan at unique sites within the interglobular domain and chondroitin sulfate attachment regions that are also cleaved when cartilage is maintained at acid pH. Matrix Biol. 20, 543–553. 3. Hembry, R. M., Knight, C. G., Dingle, J. T., and Barrett, A. J. (1982). Evidence that extracellular cathepsin D is not responsible for the resorption of cartilage matrix in culture. Biochim. Biophys. Acta 714, 307–312. 4. Goto, T., Kiyoshima, T., Moroi, R., Tsukuba, T., Nishimura, Y., Himeno, M., Yamamoto, K., and Tnaka, T. (1994). Localisation of cathepsins B, cathepsins D and cathepsins L in rat osteoclast by immunolight and immunoelectron microscopy. Histochemistry 101, 33–40. 5. Duffy, J. M., Walker, B., Guthrie, D., Grimshaw, J., McNally, G., Grimshaw, J. T., Spedding, P. L., and Mollan, R. A. B. (1994). The detection, quantification and partial characterisation of cathepsin B-like activity in human pathological synovial fluids. Eur. J. Clin. Chem. Clin. Biochem. 32, 429–434. 6. Ikeda, Y., Ikata, T., Mishiro, T., Nakano, S., Ikebe, M., and Yasuoka, S. (2000). Cathepsins B and L in synovial fluids from patients with rheumatoid arthritis and the effect of cathepsin B on the activation of pro-urokinase. J. Med. Invest. 47, 61–75

7. Aisa, M. C., Rahman, S., Senin, U., Maggio, D., and Russell, R. G. G. (1996). Cathepsin B activity in normal human osteoblast-like cells and human osteoblastic osteosarcoma cells (MG-63) – regulation by interluekin-1-beta and parathyroid hormone. Biochim. Biophys. Acta 1290, 29–36. 8. Maciewicz, R. A., Wotton, S. F., Etherington, D. J., and Duance, V. C. (1990). Susceptibility of the cartilage collagens type II, type IX and type XI to degradation by the cysteine proteinases, cathepsin B and cathepsin L. FEBS Lett. 269, 189–193. 9. Brömme, D., Okamoto, K., Wang, B. B., and Biroc, S. (1996). Human cathepsin O2, a matrix protein-degrading cysteine protease expressed in osteoclasts J. Biol. Chem. 271, 2126–2132. 10. Maciewicz, R. A., and Wotton, S. F. (1991). Degradation of cartilage matrix components by the cysteine proteinases, cathepsin B and cathepsin L. Biomed. Biochim. Acta 50, 561–564. 11. Lang, A., Horler, D., and Baici, A. (2000). The relative importance of cysteine peptidases in osteoarthritis. J. Rheumatol. 27, 1970–1979. 12. Baici, A., Lang, A., Horler, D., Kissling, R., and Merlin, C. (1995). Cathepsin B in osteoarthritis – cytochemical and histochemical analysis of human femoral head cartilage. Ann. Rheum. Dis. 54, 289–297. 13. Buttle, D. J., Handley, C. J., Ilic, M. Z., Saklatvala, J., Murata, M., and Barrett, A. J. (1993). Inhibition of cartilage proteoglycan release by a specific inactivator of cathepsin B and an inhibitor of matrix metalloproteinases – evidence for 2 converging pathways of chondrocytemediated proteoglycan degradation Arth. Rheum. 36, 1709–1717. 14. Sires, U. I., Schmid, T. M., Fliszar, C. J., Wang, Z. Q., Gluck, S. L., and Welgus, H. G. (1995). Complete degradation of type X collagen requires the combined action of interstitial collagenase and osteoclastderived cathepsin B. J. Clin. Invest. 95, 2089–2095. 15. Hill, P. A., Buttle, D. J., Jones, S. J., Boyde, A., Murata, M., Reynolds, J. J., and Meikle, M. C. (1994). Inhibition of bone resorption by selective inactivators of cysteine proteinases. J. Cell. Biochem. 56, 118–130. 16. Debari, K., Sasaki, T., Udagawa, N., and Rifkin, B. R. (1995). An ultrastructural evaluation of the effects of cysteine proteinases inhibitors on osteoclastic resorptive functions. Calcif. Tiss. Int. 56, 566–570. 17. Bossard, M. J., Thaddeus, T. A., Thompson, S. K., Amegadzie, B. Y., Hannings, C. R., Jones, C., Kurdyla, J. T., McNulty, D. E., Drake, F. H., Gowen, M., and Levy, M. A. (1996). Proteolytic activity of human osteoclast cathepsin K. J. Biol. Chem. 271, 12517–12524. 18. Garnero, P., Borel, O., Byrjalsen, I., Ferreras, M., Drake, F. H., McQueney, M. S., Foged, N. T., Delmas, P. D., and Delaisse, J. M. (1998). The collagenolytic activity of cathepsin K is unique among mammalian proteinases. J. Biol. Chem. 273, 32347–32352. 19. Kafienah, W., Bromme, D., Buttle, D. J., Croucher, L. J., and Hollander, A. P. (1998). Human cathepsin K cleaves native type I and II collagens at the N-terminal end of the triple helix. Biochem. J. 331, 727–732. 20. Li, Z., Hou, W. S., and Bromme, D. (2000). Collagenolytic activity of cathepsin K is specifically modulated by cartilage-resident chondroitin sulfates. Biochemistry 39, 529–536. 21. Li, Z., Hou, W. S., Escalante-Torres, C. R., Gelb, B. D., and Bromme, D. (2002). Collagenase activity of cathepsin K depends on complex formation with chondroitin sulfate. J. Biol. Chem. 277, 28669–28676. 22. Li, Z., Yasuda, Y., Li, W., Bogyo, M., Katz, N., Gordon, R. E., Fields, G. B., and Bromme, D. (2004). Regulation of collagenase activities of human cathepsins by glycosaminoglycans. J. Biol. Chem. 279, 5470–5479. 23. Votta, B. J., Levy, M. A., Badger, A., Bradbeer, J., Dodds, R. A., James, I. E., Thompson, S., Bossard, M. J., Carr, T., Connor, J. R., Tomaszek, T. A., Szewczuk, L., Drake, F. H., Veber, D. F., and Gowen, M. (1997). Peptide aldehyde inhibitors of cathepsin K inhibit bone resorption both in vitro and in vivo. J. Bone Min. Res. 12, 1396–1406. 24. Inui, T., Ishibashi, O., Inaoka, T., Origane, Y., Kumegawa, M., Kokubo, T., and Yamamura, T. (1997). Cathepsin K antisense

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25.

26.

27.

28.

29.

30. 31.

32. 33.

34.

35.

36.

37.

38.

39.

40.

41.

oligodeoxynucleotide inhibits osteoclastic bone resorption. J. Biol. Chem. 272, 8109–8112. Drake, F. H., Dodds, R. A., James, I. E., Connor, J. R., Debouck, C., Richardson, S., Leerykaczewski, E., Coleman, L., Rieman, D., Barthlow, R., Hastings, G., and Gowen, M. (1996). Cathepsin K, but not cathepsin B, cathepsin L or cathepsin S, is abundantly expressed in human osteoclasts. J. Biol. Chem. 271, 12511–12516. Gelb, B. D., Shi, G. P., Chapman, H. A., and Desnick, R. J. (1996). Pycnodyostosis, a lysosomal disease caused by cathepsin K deficiency. Science 273, 1236–1238. Hou, W. S., Li, W., Keyszer, G., Weber, E., Levy, R., Klein, M. J., Gravallese, E. M., Goldring, S. R., and Bromme, D. (2002). Comparison of cathepsins K and S expression within the rheumatoid and osteoarthritic synovium. Arthritis Rheum. 46, 663–674. Hou, W. S., Li, Z., Buttner, F. H., Bartnik, E., and Bromme, D. (2003). Cleavage site specificity of cathepsin K toward cartilage proteoglycans and protease complex formation. Biol. Chem. 384, 891–897. Konttinen, Y. T., Mandelin, J., Li, T. F., Salo, J., Lassus, J., Liljestrom, M., Hukkanen, M., Takagi, M., Virtanen, I., and Santavirta, S. (2002). Acidic cysteine endoproteinase cathepsin K in the degeneration of the superficial articular hyaline cartilage in osteoarthritis. Arthritis Rheum. 46, 953–960. Barrett, A. J. (1987). The cystatins: a new class of peptidase inhibitors. Trends Biochem. Sci. 12, 193–196. Lerner, U. H., Johansson, L., Ransjo, M., Rosenquist, J. B., Reinholt, F. P., and Grubb, A. (1997). Cystatin C, an inhibitor of bone resorption produced by osteoblasts. Acta Phys. Scand. 161, 81–92. Ménard, H. A., and El-Amine, M. (1996). The calpain-calpastatin system in rheumatoid arthritis. Immunol. Today 17, 545–547. Yamamoto, S., Shimizu, K., Niibayashi, H., Yasuda, T., and Yamamuro, T. (1994). Immunocytochemical demonstration of calpain in synovial cells in human arthritic synovial joints. Biomed. Res. 15, 77–88. Szomor, Z., Shimizu, K., Yamamoto, S., Yasuda, T., Ishikawa, H., and Nakamura, T. (1999). Externalization of calpain (calcium-dependent neutral cysteine proteinase) in human arthritic cartilage. Clin. Exp. Rheumatol. 17, 569–554. Szomor, Z., Shimizu, K., Fujimori, Y., Yamamoto, S., and Yamamuro, T. (1995). Appearance of calpain correlates with arthritis and cartilage destruction in collagen-induced arthritic knee joints of mice. Ann. Rheum. Dis. 54, 477–483. Oshita, H., Sandy, J. D., Suzuki, K., Akaike, A., Bai, Y., Sasaki, T., and Shimizu, K. (2004) Mature bovine articular cartilage contains abundant aggrecan that is C-terminally truncated at Ala719-Ala720, a site which is readily cleaved by m-calpain. Biochem. J. 382, 253–259. Despres, N., Talbot, G., Plouffe, B., Boire, G., and Menard, H. A. (1995). Detection and expression of a cDNA clone that encodes a polypeptide containing 2 inhibitory domains of human calpastatin and its recognition by rheumatoid arthritis sera. J. Clin. Invest. 95, 1891–1896. Salvesen, G., and Enghild, J. J. (1991). Zymogen activation, specificity and genomic structures of human neutrophil elastase and cathepsin G reveal a new branch of the chymotrypsinogen superfamily of serine proteinases. Biomed. Biochim. Acta 50, 665–671. Starkey, P. M., Barrett, A. J., and Burleigh, M. C. (1977). The degradation of articular collagen by neutrophil proteinases. Biochim. Biophys. Acta 483, 386–397. Rao, N. V., Wehner, N. G., Marshall, B. C., Gray, W. R., Gray, B. H., and Hoidal, J. R. (1991). Characterisation of proteinase-3 (PR-3), a neutrophil serine proteinase. J. Biol. Chem. 266, 9540–9548. McDonnell, J., Lobner, J. M., Knight, W. B., Lark, M. W., Green, B., Poe, M., and Moore, V. L. (1993). Comparison of the proteoglycanolytic activities of human leukocyte elastase and human cathepsin G in vitro and in vivo. Connect. Tissue Res. 30, 1–9.

193 42. Kafienah, W., Buttle, D. J., Burnett, D., and Hollander, A. P. (1998). Cleavage of native type I collagen by human neutrophil elastase. Biochem. J. 330, 897–902. 43. Campanelli, D., Melchior, M., Fu, Y. P., Nakata, M., Shuman, H., Nathan, C., and Gabay, J. E. (1990). Cloning of a cDNA for proteinase 3: A serine protease, antibiotic, and autoantigen from human neutrophils. J. Exp. Med. 172, 1709–1715. 44. Schalkwijk, J., Joosten, L. A. B., Van den Berg, W. B., and Van de Putte, L. B. A. (1990). Antigen-induced arthritis in beige (Chediak-Higashi) mice. Ann. Rheum. Dis. 49, 607–610. 45. Pettipher, E. R., Henderson, B., Hardingham, T. E., and Ratcliffe, A. (1989). Cartilage proteoglycan depletion in acute and chronic antigeninduced arthritis. Arth. Rheum. 32, 601–607. 46. Potempa, J., Korzus, E., and Travis, J. (1994). The serpin superfamily of proteinase inhibitors: structure function and regulation. J. Biol. Chem. 269, 15957–15960. 47. Thompson, R. C., and Ohlsson, K. (1986). Isolation, properties and complete amino acid sequence of human secretory leukocyte protease inhibitor, a potent inhibitor of leukocyte elastase Proc. Natl Acad. Sci. USA 83, 6692–6696. 48. Gotis-Graham, I., Hogg, P. J., and McNeil, H. P. (1997). Significant correlation between thrombospondin 1 and serine proteinase expression in rheumatoid synovium. Arth. Rheum. 40, 1780–1787. 49. Mast, A. E., Enghild, J. J., Nagase, H., Suzuki, K., Pizzo, S. V., and Salvesen, G. (1991). Kinetics and physiological relevance of the inactivation of α1-proteinase inhibitor, α1-antichymotrypsin and antithrombin III by matrix metalloproteinases-1 (tissue collagenase), -2 (72-kDa gelatinase / type IV collagenase), and -3 (stromelysin) J. Biol. Chem. 266, 15810–15816. 50. Okada, Y., Watanabe, S., Nakanishi, I., Kishi, J., Hayakawa, T., Watorek, W., Travis, J., and Nagase, H. (1988). Inactivation of tissue inhibitor of metalloproteinases by neutrophil elastase and other serine proteinases. FEBS Lett. 229, 157–160. 51. Dano, K., Andreasen, P. A., Grondahl-Hansen, J., Kristensen, P., Nielsen, L. S., and Skriver, L. (1985). Plasminogen activators, tissue degradation and cancer. Adv. Cancer Res. 44, 139–266. 52. Saksela, O., and Rifkin, D. B. (1988). Cell-associated plasminogen activation: regulation and physiological functions. Ann. Rev. Cell Biol. 4, 93–126. 53. Ellis, V., Behrendt, N., and Dano, K. (1991). Plasminogen activation by receptor-bound urokinase: a kinetic study with both cell-associated and isolated receptor. J. Biol. Chem. 266, 12752–12758. 54. Andreasen, P. A., Georg, B., Lund, L. R., Riccio, A., and Stacey, S. N. (1990). Plasminogen activator inhibitors: hormonally regulated serpins. Mol. Cell Endocrinol. 68, 1–19. 55. Blasi, F., Vassalli, J-D., and Dano, K. (1987). Urokinase-type plasminogen activator: proenzyme, receptor and inhibitors. J. Cell Biol. 104, 801–804. 56. Chapman, H. A., and Stone, O. L. (1985). Characterisation of a macrophage-derived plasminogen-activator inhibitor. Similarities with placental urokinase inhibitor Biochem. J. 230, 109–116. 57. Kruithof, E. K. O (1988). Plasminogen activator inhibitors – a review. Enzyme 40, 113–121 58. Conese, M., Olson, D., and Blasi, F. (1994). Protease nexin-1-urokinase complexes are internalised and degraded through a mechanism that requires both urokinase receptor and α2-macroglobulin receptor. J. Biol. Chem. 269, 17886–17892. 59. Belcher, C., Fawthrop, F., Bunning, R., and Doherty, M. (1996). Plasminogen activators and their inhibitors in synovial fluids from normal, osteoarthritis, and rheumatoid arthritis knees. Ann. Rheum. Dis. 55, 230–236. 60. Ronday, H. K., Smits, H. H., Van Muijen, G. N. P., Pruszczynski, M. S. M., Van Langelaan, E. J., Breedveld, F. C., and Verheijen, J. H. (1996). Difference in expression of the plasminogen activator system in synovial tissue of patients with rheumatoid arthritis and osteoarthritis. Brit. J. Rheumatol. 35, 416–423.

194 61. Busso, N., Peclat, V., So, A., and Sappino, A. P. (1997). Plasminogen activation in synovial tissues: differences between normal, osteoarthritis, and rheumatoid arthritis joints. Ann. Rheum. Dis. 56, 550–557. 62. Lajeunesse, D. (2004). The role of bone in the treatment of osteoarthritis. Osteoarthritis Cartilage 12 Suppl A, S34–38. 63. Oleksyszyn, J., and Augustine, A. J. (1996). Plasminogen modulation of IL-1-stimulated degradation in bovine and human articular cartilage explants – the role of the endogenous inhibitors PAI-1, α2-antiplasmin, α1-PI, α2-macroglobulin and TIMP. Inflammation Res. 45, 464–472. 64. Saito, S., Katoh, M., Matsumoto, M., Matsumoto, S., and Masuho, Y. (1997). Collagen degradation induced by the combination of IL-1α and plasminogen in rabbit articular cartilage explant culture. J. Biochem. 122, 49–54. 65. Milner, J. M., Elliott, S. F., and Cawston, T. E. (2001). Activation of procollagenases is a key control point in cartilage collagen degradation: interaction of serine and metalloproteinase pathways. Arthritis Rheum. 44, 2084–2096. 66. Vignon, E., Mathieu, P., Ejui, J., Descotes, J., Hartmann, D., Patricot, L. M., and Richard, M. (1991). Study of an inhibitor of plasminogen activator (tranexamic acid) in the treatment of experimental osteoarthritis. J. Rheumatol. 18 (Suppl 27) 131–133. 67. van der Laan, W. H., Ronday, H. K., TeKoppele, J. M., Breedveld, F. C., Huizinga, T. W., and Verheijen, J. H. (2003). No therapeutic effect of plasmin antagonist tranexamic acid in rheumatoid arthritis. A doubleblind placebo-controlled pilot study. Clin. Exp. Rheumatol. 21, 359–362. 68. Yang, Y. H., Carmeliet, P., and Hamilton, J. A. (2001) Tissue-type plasminogen activator deficiency exacerbates arthritis. J. Immunol. 167, 1047–1052. 69. Van Ness, K., Chobaz-Peclat, V., Castellucci, M., So, A., and Busso, N. (2002). Plasminogen activator inhibitor type-1 deficiency attenuates murine antigen-induced arthritis. Rheumatology 41, 136–141. 70. Cook, A. D., Braine, E. L., Campbell, I. K., and Hamilton, J. A. (2002). Differing roles for urokinase and tissue-type plasminogen activator in collagen-induced arthritis. Am. J. Pathol. 160, 917–926. 71. Martin, T. J., Allan, E. H., and Fukumoto, S. (1993). The plasminogen activator and inhibitor system in bone remodeling. Growth Regulation 3, 209–214. 72. Hoekman, K., Lowik, C. W. G. M., Van Derruit, M., Bijvoet, O. L. M., Verheijen, J. H., and Papapoulos, S. E. (1991). Regulation of the production of plasminogen activators by bone resorption-enhancing and inhibiting factors in 3 types of osteoblast-like cells. Bone Min. 14, 189–204. 73. Leloup, G., Lemoine, P., Carmeliet, P., and Vaes, G. (1996). Bone resorption and response to calcium regulating hormones in the absence of tissue or urokinase plasminogen activator or of their type 1 inhibitor. J. Bone Min. Res. 11, 1146–1157. 74. Tumber, A., Papaioannou, S., Breckon, J., Meikle, M. C., Reynolds, J. J., and Hill, P. A. (2003). The effects of serine proteinase inhibitors on bone resorption in vitro. J. Endocrinol. 178, 437–447. 75. Daci, E., Everts, V., Torrekens, S., Van Herck, E., Tigchelaar-Gutterr, W., Bouillon, R., and Carmeliet, G. (2003). Increased bone formation in mice lacking plasminogen activators. J. Bone. Miner. Res. 18, 1167–1176. 76. Suzuki, K., Lees, M., Newlands, G. F. J., Nagase, H., and Woolley, D. E. (1995). Activation of precursors for matrix metalloproteinases 1 (interstitial collagenase) and 3 (stromelysin) by rat mast cell proteinases I and II. Biochem. J. 305, 301–306. 77. Tetlow, L. C., and Woolley, D. E. (1995). Distribution, activation and tryptase chymase phenotype of mast cells in the rheumatoid lesion. Ann. Rheum. Dis. 54, 549–555. 78. Froelich, C. J., Zhang, X. L., Turbov, J., Hudig, D., Winkler, U., and Hanna, W. L. (1993). Human granzyme B degrades aggrecan proteoglycan in matrix synthesized by chondrocytes. J. Immunol. 151, 7161–7171. 79. Ronday, H. K., van der Laan, W. H., Tak, P. P., de Roos, J. A., Bank, R. A., TeKoppele, J. M., Froelich, C. J., Hack, C. E., Hogendoorn, P. C.,

Ian M. Clark and Gillian Murphy

80.

81.

82.

83. 84.

85.

86.

87.

88.

89.

90.

91.

92.

93.

94.

95.

Breedveld, F. C., and Verheijen, J. H. (2001). Human granzyme B mediates cartilage proteoglycan degradation and is expressed at the invasive front of the synovium in rheumatoid arthritis. Rheumatology 40, 55–61. Horiuchi, K., Saito, S., Sasaki, R., Tomatsu, T., and Toyama, Y. (2003). Expression of granzyme B in human articular chondrocytes. J. Rheumatol. 30, 1799–1810. Tak, P. P., Kumer, J. A., Hack, C. E., Daha, M. R., Smeets, T. J. M., and Erkelens, G. W. (1994). Granzyme-positive cytotoxic cells are specifically increased in early rheumatoid synovial tissue. Arth. Rheum. 12, 1735–1743. Tak, P. P., Spaeny-Dekking, L., Kraan, M. C., Breedveld, F. C., Froelich, C. J., and Hack, C. E. (1999). The levels of soluble granzyme A and B are elevated in plasma and synovial fluid of patients with rheumatoid arthritis (RA). Clin. Exp. Immunol. 116, 366–370. Werb, Z. (1997). ECM and cell surface proteolysis: regulating cellular ecology. Cell 91, 439–442. Kessler, E., Takahara, K., Biniaminov, L., Brusel, M., and Greenspan, D. S. (1996). Bone morphogenetic protein-1: The type I procollagen C- proteinase. Science 271, 360–362. Sarras, M. P., Jr. (1996). BMP-1 and the astacin family of metalloproteinases: A potential link between the extracellular matrix, growth factors and pattern formation. BioEssays 18, 439–442. Moss, M. L., White, J. M., Lambert, J. H., and Andrews, R. C. (2001). TACE and other ADAM proteases as targets for drug discovery. Drug Discov. Today 6, 417–426. Ohta, S., Harigai, M., Tanaka, M., Kawaguchi, Y., Sugiura, T., Takagi, K., Fukasawa, C., Hara, M., and Kamatani, N. (2001). Tumour necrosis factor-alpha (TNF-alpha) converting enzyme contributes to production of TNF-alpha in synovial tissues from patients with rheumatoid arthritis. J. Rheumatol. 28, 1756–1763. Patel, I. R., Attur, M. G., Patel, R. N., Stuchin, S. A., Abagyan, R. A., Abramson, S. B., and Amin, A. R. (1998). TNF-alpha convertase enzyme from human arthritis-affected cartilage: isolation of cDNA by differential display, expression of the active enzyme, and regulation of TNF-alpha. J. Immunol. 160, 4570–4579. Amin, A. R. (1999). Regulation of tumour necrosis factor-alpha and tumour necrosis factor converting enzyme in human osteoarthritis. Osteoarthritis Cartilage 7, 392–394. Bohm, B. B., Aigner, T., Gehrsitz, A., Blobel, C. P., Kalden, J. R., and Burkhardt, H. (1999). Up regulation of MDC15 (metargidin) messenger RNA in human osteoarthritic cartilage. Arthritis Rheum. 42, 1946–1950. Chubinskaya, S., Cs-Szabo, G., and Kuettner, K. E. (1998). ADAM-10 message is expressed in human articular cartilage. J. Histochem. Cytochem 46, 723–729. Chubinskaya, S., Mikhail, R., Deutsch, A., and Tindal, M. H. (2001). ADAM-10 protein is present in human articular cartilage primarily in the membrane-bound form and is upregulated in response to IL-1alpha in bovine nasal cartilage. J. Histochem. Cytochem. 49, 1165–1176. McKie, N., Edwards, T., Dallas, D. J., Houghton, A., Stringer, B., Graham, R., Russell, G., and Croucher, P. I. (1997). Expression of members of a novel membrane linked metalloproteinase family (ADAM) in human articular chondrocytes. Biochem. Biophys. Res. Commun. 230, 335–339. Flannery, C. R., Little, C. B., Caterson, B., and Hughes, C. E. (1999). Effects of culture conditions and exposure to catabolic stimulators (IL-1 and retinoic acid) on the expression of matrix metalloproteinases (MMPs) and disintegrin metalloproteinases (ADAMs) by articular cartilage chondrocytes. Matrix Biol. 18, 225–237. Inoue, D., Reid, M., Lum, L., Kratzschmar, J., Weskamp, G., Myung, Y. M., Baron, R., and Blobel, C. P. (1998). Cloning and initial characterization of mouse meltrin beta and analysis of the expression of four metalloprotease-disintegrins in bone cells. J. Biol. Chem. 273, 4180–4187.

Chapter 11 Matrix Proteinases 96. Verrier, S., Hogan, A., McKie, N., and Horton, M. (2004). ADAM gene expression and regulation during human osteoclast formation. Bone 35, 34–46. 97. Wolfsberg, T. G., Primakoff, P., Myles, D. G., and White, J. M. (1995). ADAMs, membrane proteins containing a disintegrin and metalloprotease domain – multipotential roles in cell-cell and cellmatrix interaction. J. Cell Biol. 131, 275–278. 98. Apte, S. S. (2004). A disintegrin-like and metalloprotease (reprolysin type) with thrombospondin type 1 motifs: the ADAMTS family. Int. J. Biochem. Cell Biol. 36, 981–985. 99. Collins-Racie, L. A., Flannery, C. R., Zeng, W., Corcoran, C., Annis-Freeman, B., Agostino, M. J., Arai, M., DiBlasio-Smith, E., Dorner, A. J., Georgiadis, K. E., Jin, M., Tan, X. Y., Morris, E. A., and LaVallie, E. R. (2004). ADAMTS-8 exhibits aggrecanase activity and is expressed in human articular cartilage. Matrix Biol. 23, 219–230. 100. Sandy, J. D., Flannery, C. R., Neame, P. J., and Lohmander, L. S. (1992). The structure of aggrecan fragments in human synovial fluid. Evidence for the involvement in osteoarthritis of a novel proteinase which cleaves the Glu373 – Ala374 bond of the interglobular domain. J. Clin. Invest. 89, 1512–1516. 101. Lark, M. W., Bayne, E. K., Flanagan, J., Harper, C. F., Hoerrner, L. A., Hutchinson, N. I., Singer, I. I., Donatelli, S. A., Weidner, J. R., Williams, H. R., Mumford, R. A., and Lohmander, L. S. (1997). Aggrecan degradation in human cartilage. Evidence for both matrix metalloproteinase and aggrecanase activity in normal, osteoarthritic and rheumatoid joints. J. Clin. Invest. 100, 93–106. 102. Singer, I. I., Scott, S., Kawka, D. W., Bayne, E. K., Weidner, J. R., Williams, H. R., Mumford, R. A., Lark, M. W., McDonnell, J., Christen, A. J., Moore, V. L., Mudgett, J. S., and Visco, D. M. (1997). Aggrecanase and metalloproteinase-specific aggrecan neo-epitopes are induced in the articular cartilage of mice with collagen II-induced arthritis. Osteoarthritis Cartilage 5, 407–418. 103. Van Meurs, J. B., van Lent, P. L., Holthuysen, A. E., Singer, I. I., Bayne, E. K., and van den Berg, W. B. (1999). Kinetics of aggrecanase- and metalloproteinase-induced neoepitopes in various stages of cartilage destruction in murine arthritis. Arthritis Rheum. 42, 1128–1139. 104. Matthews, R. T., Gary, S. C., Zerillo, C., Pratta, M., Solomon, K., Arner, E. C., and Hockfield, S. (2000). Brain-enriched hyaluronan binding (BEHAB)/brevican cleavage in a glioma cell line is mediated by a disintegrin and metalloproteinase with thrombospondin motifs (ADAMTS) family member. J. Biol. Chem. 275, 22695–22703. 105. Sandy, J. D., Westling, J., Kenagy, R. D., Iruela-Arispe, M. L., Verscharen, C., Rodriguez-Mazaneque, J. C., Zimmermann, D. R., Lemire, J. M., Fischer, J. W., Wight, T. N., and Clowes, A. W. (2001). Versican V1 proteolysis in human aorta in vivo occurs at the Glu441Ala442 bond, a site that is cleaved by recombinant ADAMTS-1 and ADAMTS-4. J. Biol. Chem. 276, 13372–13378. 106. Tortorella, M. D., Malfait, A. M., Deccico, C., and Arner, E. (2001). The role of ADAM-TS4 (aggrecanase-1) and ADAM-TS5 (aggrecanase-2) in a model of cartilage degradation. Osteoarthritis Cartilage 9, 539–552. 107. Malfait, A. M., Liu, R.Q, Ijiri, K., Komiya, S., and Tortorella, M. D. (2002). Inhibition of ADAM-TS4 and ADAM-TS5 prevents aggrecan degradation in osteoarthritic cartilage. J. Biol. Chem. 277, 22201–22208. 108. Yamanishi, Y., Boyle, D. L., Clark, M., Maki, R. A., Tortorella, M. D., Arner, E. C., and Firestein, G. S. (2002). Expression and regulation of aggrecanase in arthritis: the role of TGF-beta. J. Immunol. 168, 1405–1412. 109. Vankemmelbeke, M. N., Holen, I., Wilson, A. G., Ilic, M. Z., Handley, C. J., Kelner, G. S., Clark, M., Liu, C., Maki, R. A., Burnett, D., and Buttle, D. J. (2001). Expression and activity of ADAM-TS5 in synovium. Eur. J. Biochem. 268, 1259–1268.

195 110. Sugimoto, K., Takahashi, M., Yamamoto, Y., Shimada, K., and Tanzawa, K. (1999). Identification of aggrecanase activity in medium of cartilage culture. J. Biochem. 126, 449–455. 111. Tortorella, M. D., Pratta, M., Liu, R. Q., Austin, J., Ross, O. H., Abbaszade, I., Burn, T., and Arner, E. (2000). Sites of aggrecan cleavage by recombinant human aggrecanase-1 (ADAMTS-4). J. Biol. Chem. 275, 18566–18573. 112. Kashiwagi, M., Enghild, J. J., Gendron, C., Hughes, C., Caterson, B., Itoh, Y., and Nagase, H. (2004). Altered proteolytic activities of ADAMTS-4 expressed by C-terminal processing. J. Biol. Chem. 279, 10109–10119. 113. Gao, G., Plaas, A., Thompson, V. P., Jin, S., Zuo, F., and Sandy, J. D. (2004). ADAMTS4 (aggrecanase-1) activation on the cell surface involves C-terminal cleavage by glycosylphosphatidyl inositolanchored membrane type 4-matrix metalloproteinase and binding of the activated proteinase to chondroitin sulfate and heparan sulfate on syndecan-1. J. Biol. Chem. 279, 10042–10051. 114. Pratta, M. A., Tortorella, M. D., and Arner, E. (2000). Age-related changes in aggrecan glycosylation affect cleavage by aggrecanase. J. Biol. Chem. 275, 39096–39102. 115. Brinckerhoff, C. E., and Matrisian, L. M. (2002). Matrix metalloproteinases: a tail of a frog that became a prince. Nature Rev. Mol. Cell Biol. 3, 207–214. 116. Eble, J. A., Ries, A., Lichy, A., Mann, K., Stanton, H., Gavrilovic, J., Murphy, G., and Kühn, K. (1996). The recognition sites of the integrins α1β1 and α2β1 within collagen IV are protected against gelatinase A attack in the native protein. J. Biol. Chem. 271, 30964–30970. 117. Nagase, H. (1995). Human stromelysins 1 and 2. Methods Enzymol. 248, 449–470. 118. Borkakoti, N., Winkler, F. K., Williams, D. H., D’Arcy, A., Broadhurst, M. J., Brown, P. S., Johnson, W. H., and Murray, E. J. (1994). Structure of the catalytic domain of human fibroblast collagenase complexed with an inhibitor. Nat. Struct. Biol. 1, 106–110. 119. Spurlino, J. C., Smallwood, A. M., Carlton, D. D., Banks, T. M., Vavra, K. J., Johnson, J. S., Cook, E. R., Falvo, J., Wahl, R. C., Pulvino, T. A., Wendoloski, J. J., and Smith, D. L. (1994). 1.56 A structure of mature truncated human fibroblast collagenase. Proteins 19, 98–109. 120. Lovejoy, B., Cleasby, A., Hassell, A. M., Longley, K., Luther, M. A., Weigl, D., McGeehan, G., McElroy, A. B., Drewry, D., Lambert, M. H., and Jordan, S. R. (1994). Structure of the catalytic domain of fibroblast collagenase complexed with an inhibitor. Science 263, 375–377. 121. Dhanaraj, V., Ye, Q. Z., Johnson, L. L., Hupe, D. J., Ortwine, D. F., Dubar, J. B., Jr., Rubin, J. R., Pavlovsky, A., Humblet, C., and Blundell, T. L. (1996). X-ray structure of a hydroxamate inhibitor complex of stromelysin catalytic domain and its comparison with members of the zinc metalloproteinase superfamily. Structure 4, 375–386. 122. Stams, T., Spurlino, J. C., Smith, D. L., Wahl, R. C., Ho, T. F., Qoronfleh, M. W., Banks, T. M., and Rubin, B. (1994). Structure of human neutrophil collagenase reveals large S1′ specificity pocket. Nat. Struct. Biol. 1, 119–123. 123. Bode, W., Reinemer, P., Huber, R., Kleine, T., Schnierer, S., and Tschesche, H. (1994). The X-ray crystal structure of the catalytic domain of human neutrophil collagenase inhibited by a substrate analogue reveals the essentials for catalysis and specificity. EMBO J. 13, 1263–1269. 124. Grams, F., Reinemer, P., Powers, J. C., Kleine, T., Pieper, M., Tschesche, H., Huber, R., and Bode, W. (1995). X-ray structures of human neutrophil collagenase complexed with peptide hydroxamate and peptide thiol inhibitors. Implications for substrate binding and rational drug design. Eur. J. Biochem. 228, 830–841. 125. Rowsell, S., Hawtin, P., Minshull, C. A., Jepson, H., Brockbank, S. M., Barratt, D. G., Slater, A. M., McPheat, W. L., Waterson, D., Henney, A. M., and Pauptit, R. A. (2002). Crystal structure of human

196

126. 127.

128.

129.

130.

131.

132.

133.

134.

135.

136.

137.

138.

139.

140.

141.

Ian M. Clark and Gillian Murphy MMP9 in complex with a reverse hydroxamate inhibitor. J. Mol. Biol. 319, 173–181. Borkakoti, N. (2004). Matrix metalloprotease inhibitors: design from structure. Biochem. Soc. Trans. 32, 17–20. Gooley, P. R., O’Connell, J. F., Marcy, A. I., Cuca, G. C., Salowe, S. P., Bush, B. L., Hermes, J. D., Esser, C. K., Hagmann, W. K., Springer, J. P., and Johnson, B. A. (1994). The NMR structure of the inhibited catalytic domain of human stromelysin-1. Nat. Struct. Biol. 1, 111–118. Morgunova, E., Tuuttila, A., Bergmann, U., Isupov, M., Lindqvist, Y., Schneider, G., and Tryggvason, K. (1999). Structure of human pro-matrix metalloproteinase-2: activation mechanism revealed. Science 284, 1667–1670. Li, J., Brick, P., O’Hare, M. C., Skarzynski, T., Lloyd, L. F., Curry, V. A., Clark, I. M., Bigg, H. F., Hazleman, B. L., Cawston, T. E., and Blow, D. M. (1995). Structure of full-length porcine synovial collagenase reveals a C-terminal domain containing a calcium-linked, four-bladed -propeller. Structure 3, 541–549. De Souza, S. J., and Brentani, R. R. (1994). On the structure/function relationship of polymorphonuclear-leucocyte collagenase. Biochem. J. 300, 605–605. Chung, L., Dinakarpandian, D., Yoshida, N., Lauer-Fields, J. L., Fields, G. B., Visse, R., and Nagase, H. (2004). Collagenase unwinds triple-helical collagen prior to peptide bond hydrolysis. EMBO J. 23, 3020–3030. Gomis-Rüth, F. X., Gohlke, U., Betz, M., Knäuper, V., Murphy, G., López-Otín, C., and Bode, W. (1996). The helping hand of collagenase-3 (MMP-13): 2.7 Å crystal structure of its C-terminal haemopexin-like domain. J. Mol. Biol. 264, 556–566. Gohlke, U., Gomis-Rüth, F. X., Crabbe, T., Murphy, G., Docherty, A. J. P., and Bode, W. (1996). The C-terminal (haemopexinlike) domain structure of human gelatinase A (MMP2): Structural implications for its function. FEBS Lett. 378, 126–130. Morgunova, E., Tuuttila, A., Bergmann, U., and Tryggvason, K. (2002). Structural insight into the complex formation of latent matrix metalloproteinase 2 with tissue inhibitor of metalloproteinase 2. Proc. Natl. Acad. Sci. USA 99, 7414–7419. Clark, I. M., and Cawston, T. E. (1989). Fragments of human fibroblast collagenase: purification and characterisation. Biochem. J. 263, 201–206. Murphy, G., Allan, J. A., Willenbrock, F., Cockett, M. I., O’Connell, J. P., and Docherty, A. J. P (1992). The role of the C-terminal domain in collagenase and stromelysin specificity. J. Biol. Chem. 267, 9612–9618. Hirose, T., Patterson, C., Pourmotabbed, T., Mainardi, C. L., and Hasty, K. A. (1993). Structure-function relationship of human neutrophil collagenase: identification of regions responsible for substrate specificity and general proteinase activity. Proc .Natl. Acad. Sci. USA 90, 2569–2573. Murphy, G., Willenbrock, F., Ward, R. V., Cockett, M. I., Eaton, D., and Docherty, A. J. P (1992). The C-terminal domain of 72 kDa gelatinase A is not required for catalysis, but is essential for membrane activation and modulates interactions with tissue inhibitors of metalloproteinases. Biochem. J. 283, 637–641. Baragi, V. M., Fliszar, C. J., Conroy, M. C., Ye, Q.-Z., Shipley, J. M., and Welgus, H. G. (1994). Contribution of the C-terminal domain of metalloproteinases to binding by tissue inhibitor of metalloproteinases. C-terminal truncated stromelysin and matrilysin exhibit equally compromised binding affinities as compared to full-length stromelysin. J. Biol. Chem. 269, 12692–12697. Knäuper, V., Osthues, A., DeClerck, Y. A., Langley, K. E., Blaser, J., and Tschesche, H. (1993). Fragmentation of human polymorphonuclear leukocyte collagenase. Biochem. J. 291, 847–854. Butler, G. S., Butler, M. J., Atkinson, S. J., Will, H., Tamura, T., van Westrum, S. S., Crabbe, T., Clements, J., d’Ortho, M. P., and

142.

143.

144.

145.

146. 147.

148.

149.

150.

151.

152.

153.

154.

155.

156.

157.

Murphy, G. (1998). The TIMP2 membrane type 1 metalloproteinase “receptor” regulates the concentration and efficient activation of progelatinase A. A kinetic study. J. Biol. Chem. 273, 871–880. Frisch, S. M., and Morisaki, J. H. (1990). Positive and negative transcriptional elements of the type IV collagenase gene. Mol. Cell. Biol. 10, 6524–6532. Jonat, C., Rahmsdorf, H. J., Park, K. K., Cato, A. C. B., Gebel, S., Ponta, H., and Herrlich, P. (1990). Anti-tumor promotion and anti-inflammation – down modulation of AP-1 (Fos, Jun) activity by glucocorticoid hormone. Cell 62, 1189–1204. Nicholson, R. C., Mader, S., Nagpal, S., Leid, M., Rochette-Egly, C., and Chambon, P. (1990). Negative regulation of the rat stromelysin gene promoter by retinoic acid is mediated by an AP1 binding site. EMBO J. 9, 4443–4454. Springman, E. B., Angleton, E. L., Birkedal-Hansen, H., and Van Wart, H. E. (1990). Multiple modes of activation of latent human fibroblast collagenase – evidence for the role of a cys-73 active site zinc complex in latency and a cysteine switch mechanism for activation. Proc. Natl Acad. Sci. USA 87, 364–368. Pei, D., and Weiss, S. J. (1995). Furin-dependent intracellular activation of the human stromelysin-3 zymogen. Nature 375, 244–247. Crabbe, T., Ioannou, C., and Docherty, A. J. P (1993). Human progelatinase A can be activated by autolysis at a rate that is concentrationdependent and enhanced by heparin bound to the C-terminal domain. Eur. J. Biochem. 218, 431–438. Murphy, G., and Knäuper, V. (1997). Relating matrix metalloproteinase structure to function: Why the “hemopexin” domain. Matrix Biol. 15, 511–518. Okada, Y., Morodomi, T., Enghild, J. J., Suzuki, K., Yasui, A., Nakanishi, I., Salvesen, G., and Nagase, H. (1990). Matrix metalloproteinase 2 from human rheumatoid synovial fibroblasts. Purification and activation of the precursor and enzymic properties. Eur. J. Biochem. 194, 721–730. Hipps, D. S., Hembry, R. M., Docherty, A. J. P., Reynolds, J. J., and Murphy, G. (1991). Purification and characterisation of human 72-kDa gelatinase (type IV) collagenase. Use of immunolocalisation to demonstrate the non-coordinated regulation of the 72-kDa and 95-kDa gelatinases by human fibroblasts. Biol. Chem. Hoppe-Seyler 372, 287–296. Cowell, S.,. Knäuper, V., Stewart, M. L., d’Ortho, M. P., Stanton, H., Hembry, R. M., Lopez-Otin, C., Reynolds, J. J., and Murphy, G. (1998). Induction of matrix metalloproteinase activation cascades based on membrane-type 1 matrix metalloproteinase: associated activation of gelatinase A, gelatinase B and collagenase 3. Biochem. J. 331, 453–458. Baker, A. H., Edwards, D. R., and Murphy, G. (2002) Metalloproteinase inhibitors: biological actions and therapeutic opportunities. J. Cell Sci. 115, 3719–3727. Docherty, A. J. P., Lyons, A., Smith, B. J., Wright, E. M., Stephens, P. E., Harris, T. J. R., Murphy, G., and Reynolds, J. J. (1985). Sequence of human tissue inhibitor of metalloproteinases and its identity to erythroid-potentiating activity. Nature 318, 66–69. Boone, T. C., Johnson, M. J., De Clerck, Y. A., and Langley, K. E. (1990). cDNA cloning and expression of a metalloproteinase inhibitor related to tissue inhibitor of metalloproteinases. Proc. Natl Acad. Sci. USA 87, 2800–2804. Pavloff, N., Staskus, P. W., Kishnani, N. S., and Hawkes, S. P. (1992). A new inhibitor of metalloproteinases from chicken – ChIMP-3 - a 3rd member of the TIMP family. J. Biol. Chem. 267, 17321–17326. Apte, S. S., Hayashi, K., Seldin, M. F., Mattei, M.-G., Hayashi, M., and Olsen, B. R. (1994). Gene encoding a novel murine tissue inhibitor of metalloproteinases (TIMP), TIMP-3, is expressed in developing mouse epithelia, cartilage, and muscle, and is located on mouse chromosome 10. Dev. Dyn. 200, 177–197. Apte, S. S., Mattei, M.-G., and Olsen, B. R. (1994). Cloning of the cDNA encoding human tissue inhibitor of metalloproteinases-3

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158.

159.

160.

161.

162.

163.

164.

165.

166.

167.

168.

169.

170.

171.

172.

(TIMP-3) and mapping of the TIMP3 gene to chromosome 22. Genomics 19, 86–90. Apte, S. S., Olsen, B. R., and Murphy, G. (1995). The gene structure of tissue inhibitor of metalloproteinases (TIMP)-3 and its inhibitory activities define the distinct TIMP gene family. J. Biol. Chem. 270, 14313–14318. Leco, K. J., Khokha, R., Pavloff, N., Hawkes, S. P., and Edwards, D. R. (1994). Tissue inhibitor of metalloproteinases-3 (TIMP-3) is an extracellular matrix-associated protein with a distinctive pattern of expression in mouse cells and tissues. J. Biol. Chem. 269, 9352–9360. Silbiger, S. M., Jacobsen, V. L., Cupples, R. L., and Koski, R. A. (1994). Cloning of cDNAs encoding human TIMP-3, a novel member of the tissue inhibitor of metalloproteinase family. Gene 141, 293–297. Uria, J. A., Ferrando, A. A., Velasco, G., Freije, J. M. P., and Lopez-Otin, C. (1994). Structure and expression in breast tumors of human TIMP-3, a new member of the metalloproteinase inhibitor family. Cancer Res. 54, 2091–2094. Greene, J., Wang, M., Liu, Y. E., Raymond, L. A., Rosen, C., and Shi, Y. E. (1996). Molecular cloning and characterization of human tissue inhibitor of metalloproteinase 4. J. Biol. Chem. 271, 30375–30380. Leco, K. J., Apte, S. S., Taniguchi, G. T., Hawkes, S. P., Khokha, R., Schultz, G. A., and Edwards, D. R. (1997). Murine tissue inhibitor of metalloproteinases-4 (Timp-4): cDNA isolation and expression in adult mouse tissues. FEBS Lett. 401, 213–217. Gomis-Ruth, F. X., Maskos, K., Betz, M., Bergner, A., Huber, R., Suzuki, K., Yoshida, N., Nagase, H., Brew, K., Bourenkov, G. P., Bartunik, H., and Bode, W. (1997). Mechanism of inhibition of the human matrix metalloproteinase stromelysin-1 by TIMP-1. Nature, 389, 77–81. Wilhelm, S. M., Collier, I. E., Marmer, B. L., Eisen, A. Z., Grant, G. A., and Goldberg, G. I. (1989). SV40-transformed human lung fibroblasts secrete a 92-kDa type IV collagenase which is identical to that secreted by normal human macrophages. J. Biol. Chem. 264, 17213–17221. Goldberg, G. I., Strongin, A., Collier, I. E., Genrich, L. T., and Marmer, B. L. (1992). Interaction of 92-kDa type IV collagenase with the tissue inhibitor of metalloproteinases prevents dimerization, complex formation with interstitial collagenase, and activation of the proenzyme with stromelysin. J. Biol. Chem. 267, 4583–4591. Goldberg, G. I., Marmer, B. L., Grant, G. A., Eisen, A. Z., Wilhelm, S., and He, C. (1989). Human 72-kilodalton type IV collagenase forms a complex with a tissue inhibitor of metalloproteases designated TIMP-2. Proc. Natl Acad. Sci. USA 86, 8207–8211. Stetler-Stevenson, W. G., Krutzsch, H. C., Wacher, M. P., Margulies, I. M. K., and Liotta, L. A. (1989). The activation of human type IV collagenase proenzyme. Sequence identification of the major conversion product following organomercurial activation. J. Biol. Chem. 264, 1353–1356. Howard, E. W., Bullen, E. C., and Banda, M. J. (1991). Regulation of the autoactivation of human 72-kDa progelatinase by tissue inhibitor of metalloproteinases-2. J. Biol. Chem. 266, 13064–13069. Murphy, G., Houbrechts, A., Cockett, M. I., Williamson, R. A., O’Shea, M., and Docherty, A. J. P (1991). The N-terminal domain of tissue inhibitor of metalloproteinases retains metalloproteinase inhibitory activity. Biochemistry 30, 8097–8101. Fridman, R., Fuerst, R. R., Bird, R. E., Hoyhtya, M., Oelkuct, M., Kraus, S., Komarek, D., Liotta, L. A., Berman, M. L., and Stetler-Stevenson, W. G. (1992). Domain structure of human 72-kDa gelatinase/type IV collagenase. Characterization of proteolytic activity and identification of the tissue inhibitor of metalloproteinases-2 (TIMP-2) binding regions. J. Biol. Chem. 267, 15398–15405. Willenbrock, F., Crabbe, T., Slocombe, P. M., Sutton, C. W., Docherty, A. J. P., Cockett, M. I., O’Shea, M., Brocklehurst, K., Phillips, I. R., and Murphy, G. (1993). The activity of the tissue inhibitors of metalloproteinases is regulated by C-terminal domain

173.

174.

175.

176.

177.

178.

179.

180.

181.

182.

183.

184.

185. 186.

187.

188.

interactions: a kinetic analysis of the inhibition of gelatinase A. Biochemistry 32, 4330–4337. Howard, E. W., Bullen, E. C., and Banda, M. (1991). Preferential inhibition of 72-kDa and 92-kDa gelatinases by tissue inhibitor of metalloproteinases-2. J. Biol. Chem. 266, 13070–13075. Howard, E. W., and Banda, M. J. (1991). Binding of tissue inhibitor of metalloproteinases 2 to two distinct sites on human 72-kDa gelatinase. Identification of a stabilization site. J. Biol. Chem. 266, 17972–17977. Kleiner, D. E., Tuuttila, A., Tryggvason, K., and Stetler-Stevenson, W. G. (1993). Stability analysis of latent and active 72-kDa type IV collagenase: the role of tissue inhibitor of metalloproteinases-2 (TIMP-2). Biochemistry 32, 1583–1592. Strongin, A. Y., Collier, I., Bannikov, G., Marmer, B. L., Grant, G. A., and Goldberg, G. I. (1995). Mechanism of cell surface activation of 72-kDa type IV collagenase. Isolation of the activated form of the membrane metalloprotease. J. Biol. Chem. 270, 5331–5338. Murphy, G., Ward, R., Hembry, R. M., Reynolds, J. J., Kühn, K., and Tryggvason, K. (1989). Characterization of gelatinase from pig polymorphonuclear leucocytes. A metalloproteinase resembling tumour type IV collagenase. Biochem. J. 258, 463–472. Ward, R. V., Hembry, R. M., Reynolds, J. J., and Murphy, G. (1991). The purification of tissue inhibitor of metalloproteinases-2 from its 72 kDa progelatinase complex – demonstration of the biochemical similarities of tissue inhibitor of metalloproteinases-2 and tissue inhibitor of metalloproteinases-1. Biochem. J. 278, 179-187. Kashiwagi, M., Tortorella, M., Nagase, H., and Brew, K. (2001). TIMP-3 is a potent inhibitor of aggrecanase 1 (ADAM-TS4) and aggrecanase 2 (ADAM-TS5). J. Biol. Chem. 276, 12501–12504. Edwards, D. R., Leco, K. J., Leco, P. A., Lim, M. S., Phillips, B. W., Raja, J., and Sharma, R. (2000). Regulation of TIMP gene expression. In “Inhibitors of Metalloproteinases in Development and Disease”, S. P. Hawkes, D. R. Edwards, R. Khokha (eds), Harwood Academic Press, Lausanne, Switzerland, pp. 13–25. Young, D. A., Phillips, B. W., Lundy, C., Nuttall, R. K., Hogan, A., Schultz, G. A., Leco, K. J., Clark, I. M., and Edwards, D. R. (2002). Identification of an initiator-like element essential for the expression of the tissue inhibitor of metalloproteinases-4 (Timp-4) gene. Biochem. J. 364, 89–99. Takahashi, C., Sheng, Z., Horan, T. P., Kitayama, H., Maki, M., Hitomi, K., Kitaura, Y., Takai, S., Sasahara, R. M., Horimoto, A., Ikawa, Y., Ratzkin, B. J., Arakawa, T., and Noda, M. (1998). Regulation of matrix metalloproteinase-9 and inhibition of tumor invasion by the membrane-anchored glycoprotein RECK. Proc. Natl Acad. Sci. USA. 95, 13221–13226. Sottrup-Jensen, L., and Birkedal-Hansen, H. (1989). Human fibroblast collagenase-α2-macroglobulin interactions. Localization of cleavage sites in the bait regions of five mammalian a-macroglobulins. J. Biol. Chem. 264, 393–401. Tortorella, M. D., Arner, E. C., Hills, R., Easton, A., Korte-Sarfaty, J., Fok, K., Wittwer, A. J., Liu, R. Q., and Malfait, A. M. (2004). Alpha2macroglobulin is a novel substrate for ADAMTS-4 and ADAMTS-5 and represents an endogenous inhibitor of these enzymes. J. Biol. Chem. 279, 17554–17561. Borth, W. (1992). α2-macroglobulin, a multifunctional binding protein with targeting characteristics. FASEB J. 6, 3345-3353. Borth, W. (1994). α2-macroglobulin, a multifunctional binding and targeting protein with possible roles in immunity and autoimmunity. Ann. N. Y. Acad. Sci. 737, 267–272. Stephens, R. W., Tapiovaara, H., Reisberg, T., Bizik, J., and Vaheri, A. (1991). Alpha-2-macroglobulin restricts plasminogen activation to the surface of RC2A leukemia cells. Cell Regul. 2, 1057–1065. Dean, D. D., Martel-Pelletier, J., Pelletier, J-P., Howell, D. S., and Woessner, J. F. (1989). Evidence for metalloproteinase and metalloproteinase inhibitor imbalance in human osteoarthritic cartilage J. Clin. Invest. 84, 678–685.

198 189. Kevorkian, L., Young, D. A., Darrah, C., Donell, S. T., Shepstone, L., Porter, S., Brockbank, S. M., Edwards, D. R., Parker, A. E., and Clark, I. M. (2004). Expression profiling of metalloproteinases and their inhibitors in cartilage. Arthritis Rheum. 50, 131–141. 190. Bau, B., Gebhard, P. M., Haag, J., Knorr, T., Bartnik, E., and Aigner, T. (2002). Relative messenger RNA expression profiling of collagenases and aggrecanases in human articular chondrocytes in vivo and in vitro. Arthritis Rheum. 46, 2648–2657. 191. Martel-Pelletier, J., McCollum, R., Fujimoto, N., Obata, K., Cloutier, J-M., and Pelletier, J-P. (1994). Excess of metalloproteases over tissue inhibitor of metalloproteases may contribute to cartilage degradation in osteoarthritis and rheumatoid arthritis. Lab. Invest. 70, 807–815. 192. Cawston, T., McLaughlan, P., Coughlan, R., Kyle, V., and Hazleman, B. (1990). Synovial fluids from infected joints contain metalloproteinasetissue inhibitor of metalloproteinase (TIMP) complexes. Biochim. Biophys. Acta 1033, 96–102. 193. van Lent, P. L., Holthuysen, A. E., Sloetjes, A., Lubberts, E., and van den Berg, W.B. (2002). Local overexpression of adeno-viral IL-4 protects cartilage from metallo proteinase-induced destruction during immune complex-mediated arthritis by preventing activation of pro-MMPs. Osteoarthritis Cartilage 10, 234–243. 194. van Meurs, J., van Lent, P., Holthuysen, A., Lambrou, D., Bayne, E., Singer, I., and van den Berg, W. (1999). Active matrix metalloproteinases are present in cartilage during immune complex-mediated arthritis: a pivotal role for stromelysin-1 in cartilage destruction. J. Immunol. 163, 5633–5639. 195. van Meurs, J., van Lent, P., Stoop, R., Holthuysen, A., Singer, I., Bayne, E., Mudgett, J., Poole, R., Billinghurst, C., van der Kraan, P., Buma, P., and van den Berg, W. (1999). Cleavage of aggrecan at the Asn341-Phe342 site coincides with the initiation of collagen damage

Ian M. Clark and Gillian Murphy

196.

197.

198.

199.

200.

201.

202.

in murine antigen-induced arthritis: a pivotal role for stromelysin 1 in matrix metalloproteinase activity. Arth. Rheum. 42, 2074–2084. Ellis, A. J., Powell, L. K., Curry, V. A., and Cawston, T. E. (1994). TIMP-1 and TIMP-2 prevent the release of collagen fragments in cartilage stimulated with IL-1. Biochem. Biophys. Res. Commun. 201, 94–101. Gendron, C., Kashiwagi, M., Hughes, C., Caterson, B., and Nagase, H. (2003). TIMP-3 inhibits aggrecanase-mediated glycosaminoglycan release from cartilage explants stimulated by catabolic factors. FEBS Lett. 555, 431–436. Partridge, N. C., Walling, H. W., Bloch, S. R., Omura, T. H., Chan, P. T., Pearman, A. T., and Chou, W-Y. (1996). The regulation and regulatory role of collagenase in bone. Crit. Rev. in Euk. Gene Exp. 6, 15–27. Bord, S., Horner, A., Hembry, R. M., Reynolds, J. J., and Compston, J. E. (1997). Distribution of matrix metalloproteinases and their inhibitor, TIMP-1, in developing human osteophytic bone. J. Anatomy 191, 39–48. Hill, P. A., Docherty, A. J. P., Bottomley, K. M., O’Connell, J. P., Morphy, J. R., Reynolds, J. J., and Meikle, M. C. (1995). Inhibition of bone resorption in vitro by selective inhibitors of gelatinase and collagenase. Biochem. J. 308, 167–175. Stahle-Backdahl, M., Sandstedt, B., Bruce, K., Lindahl, A., Jimenez, M. G., Vega, J. A., and Lopez-Otin, C. (1997). Collagenase-3 (MMP-13) is expressed during human fetal ossification and re-expressed in postnatal bone remodeling and in rheumatoid arthritis. Lab. Invest. 76, 717–728. Okada, Y., Naka, K., Kawamura, K., Matsumoto, T., Nakanishi, I., Fujimoto, N., Sato, H., and Seiki, M. (1995). Localization of matrix metalloproteinase-9 (92-kilodalton gelatinase/type IV collagenase, gelatinase B) in osteoclasts – implications for bone resorption. Lab. Invest. 72, 311–322.

Part II

Structure and Metabolism of the Extracellular Matrix of Bone and Cartilage

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Chapter 12

Mineralization, Structure and Function of Bone Adele L. Boskey, Ph.D.

Starr Chair in Mineralized Tissue Research, Hospital for Special Surgery, New York, NY 10021 and Weill Medical College and Graduate School of Medical Sciences of Cornell University, New York, NY 10021

IV. Bone Modeling and Remodeling References

I. Abstract II. The Structure and Function of Bone III. Bone Mineralization

I. ABSTRACT

the locomotion process. In addition to these mechanical functions, bone provides a protective housing for bloodforming marrow, and the bone serves as a reservoir for mineral ions (Ca+2, Mg+2, PO4−3). There are two histologically or radiologically defined bone types [1], dense (also known as compact) bone, and spongy or cancellous (also known as trabecular) bone. When bone is newly formed, as during development or fracture healing, the matrix is loose or woven. With maturation, the bone becomes denser, better organized, and better able to serve its mechanical function. Mature bone is described as compact (dense) or trabecular (meaning composed of little beams). The general features of both compact and trabecular bones are similar. Both are solid mineralized matrices with small canals (canaliculi), and spaces (lacunae), and bone cells. (For discussion of bone cells, see Chapter 14.) In cancellous bone, the matrix, lacunae, and mineral-encased cells (osteocytes) are organized in the form of thin interconnecting spicules. In cortical bone the tissue is organized in Haversian systems or osteons. The Haversian systems consist of a canal containing a blood vessel surrounded by concentric and interstitial lamellae.

The development of bone is a complex process that involves cellular, extracellular, and physicochemical events. Here, the overall process of bone formation is reviewed from a biologic and chemical viewpoint. The factors regulating the initial mineralization of osteoid, and those regulating the entire process of mineralization are the main subjects of this review. Attention is paid to new data on the role of matrix proteins in these processes and to the interaction between collagen and matrix proteins, and matrix proteins and cells in regulating bone mineralization.

II. THE STRUCTURE AND FUNCTION OF BONE Bones are dynamic tissues, replaced as they age or are damaged with newly deposited bone. These dynamic tissues serve numerous essential functions in man and other vertebrates. Bone provides mechanical protection for internal organs, allows the animal to direct motion, and facilitates Dynamics of Bone and Cartilage Metabolism

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In addition to descriptions based on architecture (spicules or lamellae) bone may also be described based on its mechanism of formation during development. Bones that develop by the replacement of a cartilage model (endochondral ossification) are often distinguished from those which form directly (intramembranous ossification). Details of these events will be presented later.

A. The Composition of Bone Bone is a composite material consisting of mineral, collagen, water, noncollagenous proteins, lipids, vascular elements, and cells. The absolute amounts of these constituents vary with animal age, sex, tissue site, health, and dietary status. The arrangement of the bone tissues, as described in the classic treatise by Wolff [2], is such that bones are organized optimally to resist loads imposed by functional activities. Thus with growth and development the tissue must be constantly reshaped and remodeled to maintain this maximization, and maintain a form appropriate to its mechanical function. As used here, modeling refers to formation of bone on sites it has not been before, whereas remodeling refers to its formation on surfaces previously containing bone. Both these processes are required to shape bone. The mineral found in bone is an analog of the naturally occurring geologic mineral hydroxyapatite, Ca10(PO4)6(OH)2 [3]. Bone apatite is a calcium- and hydroxide-deficient apatite containing numerous impurities, the most abundant of which is carbonate [4]. Sodium, potassium, fluoride, and citrate are also common substituents [3–5]. In general, in animals of the same age, the proportion of mineral is greatest in the bones of the ear, and lowest in the ribs. Woven and lamellar bones have a lower mineral content than compact bone. These bones will be described below. Bone mineral apatite crystals are relatively small (~9 nm × 6 nm × 2 nm) in young bone as revealed by atomic force microscopy and electron microscopy [6, 7]. This is in contrast with the slightly larger apatite found in tooth enamel or the much larger geologic apatites. It is the small size of bone mineral crystals that facilitates the incorporation and adsorption of foreign ions, and enables bone mineral to dissolve in the acidic milieu created by the osteoclast during bone remodeling. The small crystal size also enables the crystals to provide the flexible collagen fibrils of the mineral–collagen composite with structural rigidity. Bone mineral crystals, while appearing like needles in some species and plates in others [7] have been shown to be thin curved plates [8], which agglomerate, or fuse, as the bone matures [9].

The variation in the amount of mineral in an osteon was first appreciated by the examination of backscattered electron images [10] and has also been quantified by Fourier transform infrared microspectroscopy (FTIRM) and microscopic imaging (FTIRI) techniques in, which an infrared spectrophotometer, coupled to a light microscope, is used to measure mineral content, composition, and parameters related to mineral crystal size and perfection at 6–20 µm spatial resolution [11]. Figure 1A shows a light micrograph of one such osteon from healthy adult bone, and Figure 1B shows typical FTIR spectral maps from this region. Figure 1C shows the mineral content, calculated from these spectra as the ratio of absorptions of the phosphate (900–1200 cm−1) and amide I (1585–1720 cm−1) peaks respectively in four lines orthogonal to the center of the Haversian canal. Figure 1D shows the variation in carbonate to phosphate ratio along these same directions, and Figure 1E shows the changes in crystal maturity estimated from analyses of the underlying sub-bands in the complex phosphate spectra. These data are presented to illustrate that significant changes in mineral properties occur as the tissue ages, and to emphasize that bone is a dynamic tissue. The carbonate content of bone mineral increases with maturity [12], and there are also variations in the nature of the carbonate substitution. Carbonate can replace hydroxide ions (A-type carbonate) or phosphate ions (B-type carbonate), or can adsorb onto the surface of the apatite (labile carbonate). With increasing bone age the proportion of labile carbonate decreases, and in the most mature bone apatites there is a preponderance of B-substituted carbonate. Less is known about the age and site variation of the other ions that accumulate in bone, and there is some debate as to whether ions such as Mg and Sr are incorporated into the apatite lattice or only reside on the surface of mineral crystals [3]. The second most abundant component of bone is collagen, predominantly type I (for discussion of collagen structure and function see Chapters 1 and 2). Collagen provides bone with elasticity and flexibility and directs the organization of the matrix. The collagen fibers in bone are organized in sheets (periosteal bone), or circumferentially (osteonal bone). They are oriented spirally in the lamellae, and all lie in the same direction in a single lamellae, but in adjacent lamellae they are oriented in a different direction. In physiologically mineralized tissues the long axes of the apatite crystals are always aligned parallel to the axis of the collagen fibrils [9]. This orientation allows the crystals to contribute to the strength of bone. Illustrations of the importance of collagen for the mechanical strength of bone are provided by man and mice with osteogenesis imperfecta. This “brittle” bone disease due

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A

B

C

D

E Figure 1 Mineral property changes in osteonal bone. (A) Light micrograph of an osteon in adult human bone. (B) Fourier transform infrared spectra going from the center of the osteon (0) to 60 µm from the origin at 10 µm spatial resolution. (C) Mineral-to-matrix ratio in the osteon calculated from spectra obtained in four orthogonal directions increases gradually from the center of the Haversian canal. (D) Carbonate-to-phosphate ratio calculated from the spectra increases as mineral content increases in the four orthogonal directions from the center of the Haversian canal. (E) The mineral crystal maturity (size and perfection) as calculated from curve-fit spectra increases with distance from the center of the Haversian canal. Reprinted from Paschalis et al. [11] FTIR microspectroscopic analysis of human osteonal bone.” Calcif. Tissue Int. 59, 480–487, with kind permission of Springer Science and Business Media.

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to genetic abnormalities in collagen structure is associated with frequent fractures, even in the neonate [13, 14]. A second illustration is the observation that when bone is decalcified (by soaking in acid or decalcifying agents) it becomes soft and rubber-like. Water accounts for 5–10% of the weight of bone tissue. Tissue hydration is needed both for nutrition and function of cells, and also for mechanical function of the mineral–collagen composite [15]. Hydrogen bonds between water and collagen contribute to the stabilization of the collagen fibril, and there have been suggestions that dehydration of the bone collagen, i.e., replacement of water by mineral, may take place during the mineralization process. Noncollagenous proteins, reviewed in Chapters 3-5, make up approximately 5% of the dry weight of bone. They are important for a variety of processes described elsewhere [15] as well as in this book. The functions of noncollagenous proteins in mineralization will be reviewed in detail below. Lipids, which account for 2–5 % of the organic matrix of bone, are components of cell membranes, and as such are essential for cell function. They also appear to play an important role in initial mineralization and bone [15].

B. The Cell Biology of Bone Formation The matrix of bone changes with growth and development. To appreciate these changes, it is useful to briefly review the embryological stages of development. More details can be found in a recent article by Reddi [16]. Initial long bone development begins by the vascular invasion of a cartilaginous model. During embryogenesis, a cartilage matrix is formed in the center of the bone surrounded by the periosteum /perichondrium, which throughout development defines the boundary between bone and the soft connective tissues. As the cartilage cells become enlarged, the matrix calcifies, while the cells in the periosteum form a thin layer of bone. The calcified cartilage is removed as a vascular bud and tissue from the periosteum invades. Concurrently, the cartilage cells on either side of this cavity continue to proliferate, causing the bone to grow in length and width. The periosteum continues to form, the cartilage cells closest to the cavity hypertrophy (swell), and a calcified matrix forms. Bone forms on the spicules of calcified cartilage (endochondral bone) and eventually is remodeled, as marrow cells fill in the cavity. As bone starts to form surrounding a marrow cavity, growth continues as the proximal and distal epiphyses move further and further apart. The networks of growth factors, cytokines, hormones, and gene regulatory elements that regulate these processes have been discussed elsewhere

in this book (Chapter 6). For this review, it is only important to note that, during development, calcification of cartilage occurs first, and calcified cartilage is replaced by bone. The calcifying cartilage matrix is enriched in type X collagen [17], extracellular membrane-bound bodies known as matrix vesicles [18], and metalloproteinases [19], all of which are important for mineralization. The matrix vesicles contain Ca-transport proteins known as annexins, complexed acidic phospholipids, and other associated proteins, as part of their “nucleational core” [20]. As endochondral ossification starts, the calcifying cartilage matrix is depleted to some extent of the large aggregating proteoglycans [21]. All of these are important for the calcification process. During intramembranous ossification, as in endochondral ossification, bone is first laid down as a network of spicules. In connective tissue, cells differentiate directly, not osteoblasts, but similar to the accumulation of osteoblasts on the calcified cartilage spicules. These cells aggregate and deposit osteoid, which then becomes mineralized. Like calcifying and uncalcified cartilage, the unmineralized and mineralized osteoid have different composition. During initial osteoid formation, the matrix is rich in cell-binding proteins such as laminin, and fibronectin. Later, bone sialoprotein and osteopontin, also cell-binding proteins, are added [22]. Type I collagen and the small proteoglycans decorin, biglycan, and osteodidherin [23], which regulate collagen fibrollogenesis [23, 24], are laid down upon this initial network. As the tissue matures the collagen fibrils form inter- and intrafibrillar cross-links; the number of such cross-links increases with tissue age [25, 26]. Osteocalcin, the most abundant of the noncollagenous matrix proteins in humans [27], does not accumulate in the osteoid, but only is expressed and appears in the bones as it mineralizes [22, 28]. Several other proteins that are concentrated in the mineralized matrix are listed in Table I, some of these are synthesized by osteoblasts, but some from serum accumulate in bone because of their affinity for hydroxyapatite [15]. As reviewed elsewhere in this volume, the factors regulating the expression and post-translational modification of most of these proteins are known. But how their expression and post-translational modification is linked with the onset of mineralization is less clear.

III. BONE MINERALIZATION Physiologic bone mineralization in mammals involves the ordered deposition of apatite on a type I collagen matrix. The bone apatite crystals are always deposited such that their longest dimension lies parallel to the axis of the collagen fibril (Fig. 2). The cascade of events

Chapter 12 Mineralization, Structure and Function of Bone Table I.

The Extracellular Matrix Proteins of Bone

Collagens Type I Type XI Type V Type III Type XII

Proteoglycans Versican Syndecan CS-decorin CS-biglycan Lumican Osteoadherin Serum proteins Albumin Fetuin (α 2HS-glycoprotein) IgG IgE

Phosphoproteins BAG-75 Bone sialoprotein Dentin matrix protein-1 Dentin sialoprotein MEPE (matrix extracellular phosphoglycoprotein) Osteopontin Osteonectin Gamma carboxy-glutamate containing proteins Osteocalcin Matrix Gla protein

Other Thrombospondin Fibromodulin Proteolipids

includes the formation of that matrix and the oriented deposition of these crystals. Our current knowledge of the process of bone mineralization is derived from a combination of cell and organ culture studies, analysis of bones from normal, mutant, transgenic, and diseased species, analogies drawn from biomineralization in other species and cell free studies in solution [15]. While mineral phases other than apatite have been proposed to be formed as precursors, X-ray diffraction [3], NMR [29], and other studies of newly formed embryonic bone [30–34], have failed to show the presence of other

Figure 2

Electron micrograph of the newly formed mineral in a developing rat bone showing the alignment of the long axis of the electron dense mineral crystals along the collagen fibrils. Courtesy of Dr. S. B. Doty.

205 calcium phosphate phases such as brushite, octacalcin phosphate, or noncrystalline (amorphous) tri-calcium phosphate in the youngest bone. This does not exclude the possibility that these phases may form transiently during the initial steps of bone formation prior to apatite formation. Some nonvertebrates form amorphous mineral phases which may be transient or persistent [35]. Calcium phosphate mineral deposition in bone is, in part, a physical chemical process. Precipitation, in general, takes place from solution when the ion activity product in solution exceeds the solubility product of the precipitating phase. As a first approximation, in the case of hydroxyapatite, that would mean [Ca+2]5 [PO4−3]3 [OH] ≥ 10−57 [36] and most body fluids would be undersaturated. When the concentration of any of these ions is increased, as may be the case during the formation of urinary or salivary stones, crystal deposition may occur. However, even in such cases an energy requiring nucleation process must occur. During nucleation, ions or ion clusters must associate to form a stable configuration containing the building block(s) of the crystal in question. Once this structure (the “critical nucleus”) is large enough to persist in solution, it is easy to continue to add ions or ion clusters to it during the crystal growth process [15]. Secondary nucleation occurs as new crystal starts to form on the initial crystal in a fashion analogous to glycogen branching. The new crystal branches separate and provide additional nuclei. Nucleation, as described above, may occur de novo, or may occur on foreign substances (heterogeneous nucleation). If the foreign substance surface resembles the surface of the nucleus, the process of “epitaxial nucleation” occurs. In biological systems, there are numerous heterogeneous and epitaxial nucleators that facilitate mineralization in a variety of ways. Furthermore, in bone, it is apparent that there are many such nucleation sites, since initial crystals start to form concurrently at numerous separated loci [3]. Examination of newly mineralized collagen shows the first crystals forming in the “e-bands” throughout the collagen matrix [6, 37]. There are two not necessarily mutually exclusive theories concerning how mineral deposition starts at discrete sites in bone. During endochondral ossification, cartilage calcification is widely believed to start within extracellular membrane-bound bodies, known as matrix vesicles (ECMVs) [18, 38]. ECMVs (Fig. 3) released from the chondrocyte, are seen as the site where mineral first appears in the extraterritorial matrix adjacent to the hypertrophic cells of the calcifying cartilage. ECMVs provide protected areas for the accumulation of mineral ions away from the space-filling proteoglycan aggregates that keep the cartilage matrix hydrated, and by virtue of their negative charge and bulk, are effective inhibitors of mineral nucleation

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Figure 3 Electron micrograph showing extracellular matrix vesicles in calcifying cartilage of a young rat. Courtesy of Dr. S. B. Doty.

and growth [39–42]. ECMVs contain a nucleational core [20] consisting of acidic phospholipids, calcium, inorganic phosphate, and a Ca-transport protein, annexin. This nucleational core is believed to be responsible for membraneassociated mineral formation within the ECMVs. The ECMVs also contain enzymes that can modulate inhibitors of mineral formation found in the extracellular matrix [43, 44]. Isolated ECMVs often have matrix proteins such as type X collagen associated with them [38]. This small chain collagen is important to the organization of the matrix and cell produced by hypertrophic chondrocytes [45]. The ECMVs are always adjacent to collagen (Fig. 3), and there is some controversy concerning how the mineral formed within matrix vesicles gets to the collagen matrix upon which the bone mineral is deposited. High-resolution 3D tomographic imaging of the mineralizing turkey tendon, a highly organized model system frequently used to study processes in bone, has demonstrated mineral crystals growing across from mineralizing vesicles to the collagen fibers [46]. These studies also showed that there were mineralizing sites on the tendon collagen not associated with ECMVs, suggesting that ECMV and collagen mineralizing could occur independently of binding [45]. Mineralization of osteoid, as opposed to cartilage and tendon mineralization, is rarely associated with ECMVs;

ADELE L. BOSKEY

however, ECMVs do occur where bone mineralization is impaired, for example in hypocalcemia [47]. It is not known whether the delayed mineralization of the osteoid enables the observation of ECMVs or whether ECMVs are formed specifically to promote mineral formation when other mechanisms are hindered. It is known that purified type I collagen aggregated into fibrils can support apatite formation from metastable calcium phosphate solutions [48]. However, when extracted collagen was implanted in the animal, mineral deposition was relatively slow [49]. Termine showed that, after noncollagenous proteins were extracted from bone collagen with chaotropic solvents and protease inhibitors, the collagen could not be re-mineralized in vitro [50]. Likewise, Endo and Glimcher showed that dephosphorylation of bone collagen retarded its in vitro mineralization [51]. From this and other evidence it is now generally accepted that there are noncollagenous proteins associated with the collagen that can serve both as nucleators of initial mineral formation and regulators of crystal size and shape [3, 15]. The challenge has been in determining which proteins are essential for these processes. The approach to this question has included identifying which proteins are expressed at the boundary between mineralized bone and osteoid (the mineralization front) using molecular [15, 52, 53, 54] and histochemical [22, 55] techniques. These studies have revealed inter alia that a family of phosphoproteins, the so-called SIBLING proteins [56] for small integrin binding ligand N-glycosylated, are all expressed at the start of mineralization. The SIBLING proteins are all expressed on human chromosome 4q. They include osteopontin, bone sialoprotein, matrix extracellular phosphoglycoprotein (MEPE), dentin matrix protein 1 (DMP1), and dentin sialophosphoprotein (DSPP). All these proteins also bind to complement factor H [57] and several of those studied to date activate metalloproteases [58], showing they have multiple functions as signaling molecules and potential regulators of mineralization. Table II lists additional matrix proteins which have been noted to change in distribution or post-translational modification at the mineralization front. Even though a protein may have an altered distribution or composition at the mineralization front, it may not be directly involved in causing mineral crystals to deposit. It may simply be there because it has a high affinity for mineral. To determine which proteins can act as nucleators, investigators have studied the effects of individual proteins, either isolated from bone or prepared using recombinant technology, on in vitro mineralization [15]. Studies of solution-medicated mineralization have included analysis carried out in media containing only calcium and phosphate salts, media made to mimic the composition of

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Chapter 12 Mineralization, Structure and Function of Bone Table II. Modified gene

Mutant Animals with Genetically Altered Matrix Proteins and Bone Phenotypes Phenotype

Mineral properties

Reference

Knockouts Biglycan Matrix glaprotein Ostoecalcin DMP-1

Vanishing bone Osteoporosis-like Vascular calcification Thickened bones Altered bone structure

Osteonectin

Thinning bones

Osteopontin

Thickened bones

Type X collagen MEPE Collagen I

Transgenics Disrupted epiphyses Increased bone mass Brittle bone disease (young animals)

body fluids (synthetic lymph [59]), media in which the calcium, phosphate, and hydroxide ion concentrations are kept fixed (constant composition [60]), and others in which the concentrations of these ions are allowed to vary. Some studies examine growth of calcium phosphates added as “seed crystals” to these solutions, others evaluate nucleators in solutions free of dust and other potential heterogeneous nucleators, while still others look for the effects in agar, silica, or gelatin gels containing the proteins in question [61]. Despite the variation in these conditions, in general, there is consistency as to which proteins are found to act as nucleators and which can bind to nuclei or crystals and inhibit or regulate crystal proliferation and growth. These effects, of course, are all concentration dependent. Only a few bone matrix proteins, and the nucleational core discussed above [20], act as in vitro nucleators. The most effective of the proteins analyzed to date is bone sialoprotein [62]. This bone-specific protein (BSP) causes apatite formation in a variety of in vitro conditions [3, 63, 64]. Both the native and the recombinant form of BSP can inhibit mineral crystal growth as well [15, 65]. BSP is distinguished from the other phosphorylated sialoproteins made by osteoblasts in that it is rich in poly-glutamate repeats, whereas another SIBLING protein, osteopontin, contains poly-aspartate sequence, and BAG-75 has a mixture of poly-glutamate and poly-aspartate repeats. BSP, BAG-75, and alkaline phosphatase form spherical structures in osteoblast cultures that are the foci for initial calcification [64]. It has been suggested based on studies with synthetic poly-peptides and peptidomimetics that this poly-glutamate repeat stabilizes the apatite nuclei [66, 67]. This would explain why BSP at high concentrations could coat apatite crystals and retard their growth [65].

Larger but fewer crystals

Less mineral Larger crystals Increased mineral content Increased crystal size Increased mineral content Increased crystal size

No mineral change ? Decreased mineral content Decreased crystallinity

85 87 88, 89 90 86 86

91 101 102

A variety of other bone and dentin matrix components seem to have the ability to act as nucleators at low concentrations and regulate the extent of proliferation of mineral crystals at higher concentrations. These include the lipids of the “nucleational core” of ECMVs [59], biglycan, the small CS-containing leucine-rich proteoglycan of bone [68], and several phosphoproteins including dentin matrix protein-1, a component of bone as well as dentin [69], and dentin phosphophoryn [70]. Although the precise mechanisms of action of these proteins may differ, it is likely that they can interact with specific faces on the apatite crystal, stabilizing the critical nucleus, and blocking growth in certain directions [15]. There is also evidence that these proteins’ ability to nucleate apatite depends on enzymatic and post-translational modifications [69]. Other extracellular matrix proteins isolated from bone, such as osteonectin, osteopontin, and osteocalcin do not act as nucleators but bind to apatite with high affinities [15] and act as effective inhibitors of apatite proliferation [71, 72]. It is interesting to note that a highly phosphorylated form of osteopontin can act as a nucleator, while a small peptide from MEPE is an inhibitor of cell-medicated mineralization [73]. It is thus likely that many of these proteins are modified at sites of mineralization to either enhance or modulate the calcification process. Although it is known that the proteins shown to be nucleators in solution are associated with collagen [3], to determine their action in situ investigators have used in vitro studies of cell and organ systems [e.g., 15, 64, 74]. These studies provide data on the sequence of expression of the matrix proteins [23, 27, 75] and the effects of altered matrix protein expression on mineralization [76–79]. For example, blocking the synthesis of large proteoglycans, and thereby preventing their sulfation, or accelerating the

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degradation of large proteoglycans, enhanced the rate of mineralization in calcifying cartilage cultures [75]. Blocking phosphoprotein phosphorylation in the same culture system blocked mineralization [76]. These cell culture studies support the postulated role of proteoglycans and phosphoproteins in bone mineralization. The development of embryonic stem cell technology by which proteins synthesized in ovo could be modified, ablated (“knocked out”), or inserted (“knocked in”) provides an additional tool to probe the functions of the bone matrix proteins. Knocking out these proteins does not always produce a bone phenotype, because many of the essential processes, such as mineralization, are regulated in redundant ways. However, proof of function of several proteins has become available through the studies of both transgenic over-expression (expressing an altered gene protein) and/or knockout (gene ablation) animal models. The first example of a transgenic mouse relevant to bone disease was the model of human osteogenesis imperfecta (brittle bone disease), produced by the random insertion of a viral gene sequence into the mouse genome [80]. These transgenic mice showed the typical bone fragility of the human disease. Other models of human bone disease are

seen in the MEPE knockout which forms hypermineralized bones [81], the brittle bone collagen knock-in [82], the dwarf mice created by deletion of the proteoglycan aggregates link protein [83], the rachitic mice created by deletion of vitamin D hydroxylase gene [84], and the osteoporotic-like condition of biglycan knockout mice [85], to name a few. The effect of the general loss of bone matrix proteins or tissue specific loss of these proteins is indicated in Table II. The numerous mutants in which knockout of cytokines and hormones or hormone receptors produce a bone phenotype are not discussed here. The bones of the knockout and other mutant animals are usually described based on histology, radiography, and changes in mechanical testing. We have used FTIR microspectroscopy and imaging to further characterize these bones (Fig. 4) [85–91]. Changes in bone strength and mineral properties have validated most of the predictions made from in vitro data. Thus, for example, biglycan is an in vitro nucleator [68] and when it is absent in the mouse there is less mineral present but whatsoever crystals are present are larger [85]. There must be many nucleators present in bone, but, when one is absent, the crystals formed initially on other nucleating materials are sparse

Figure 4 Infrared imaging of the bone mineral in two representative knockout animals. Top: the DMP1 knockout at 4 weeks has decreased mineral:matrix ratio (mineral content) in its cortical bone, and increased mineral crystal size/perfection (crystallinity) relative to age matched control. Bottom: the osteonectin (ON) knockout mouse has decreased amounts of trabecular bone at 11 weeks of age relative to wild-type controls, and both the mineral content and the crystallinity in the knockout are decreased. Note the scales for mineral:matrix shown in the top and bottom figures are not the same as the trabecular bone has a high mineral:matrix content at this age in the wild-type.

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Chapter 12 Mineralization, Structure and Function of Bone

and tend to grow to greater size. Similarly DMP-1 in vitro is both an inhibitor and a nucleator [69]; when it is absent in the knockout mouse there are fewer crystals (supporting the role as a nucleator), but these crystals become excessively large, implying a role as an inhibitor. In contrast, osteopontin is an in vitro [71], and the osteopontin knockout has increased mineral content and increased crystal size [86], both changes being associated with the absence of an inhibitor of crystal growth and proliferation. Osteopontin functions both as a matrix protein and a signaling molecule, and has an important role in bone remodeling [92, 93].

IV. BONE MODELING AND REMODELING Following initial mineralization of calcified cartilage or osteoid, the mineralized matrix remains in a dynamic state. It must be reshaped as the bones grow, and as the loads applied to the bone vary. The periosteum, where new bone formation starts, is very responsive to mechanical load, enabling the modeling to occur. Mineral crystals grow, agglomerate, dissolve, and thus change in composition during remodeling [3, 8, 9, 11]. During endochondral ossification the calcified cartilage matrix serves as a site for the deposition of mineral in the form of the primary spongiosa. Mineral crystals in the woven bone are smaller and contain more impurities than those in compact bone [30, 94] and larger than those in calcified cartilage [95]. This combination of calcified cartilage and woven bone is removed and replaced by woven bone alone. As the epiphyses move further apart and the bone grows longer, the proportions of the bone also change. This modeling is accomplished by the resorption of bone on the endosteal surface and the formation of periosteal bone. Modeling of the structures formed by intramembranous ossification are similarly governed by the periosteum. Remodeling, as contrasted to modeling, refers to the continuous shaping of the bones in response to repair in which the events of endochondral ossification are recapitulated. Remodeling is the skeletal process that allows mineral ion homeostasis. It is regulated by a number of hormones such as vitamin D, parathyroid hormone, calcitonin, and estrogen [96]. The remodeling process in the healthy bone involves the coupled actions of bone-forming and bone-resorbing cells. Sequentially, the remodeling process involves recruitment of osteoclasts to a site on the bone surface, an osteopontin-mediated process [97], and the removal of bone mineral and matrix, creating a resorption pit. As the osteoclast moves away, osteoblasts move in, fill in the pit with osteoid, which is then mineralized. It is this coupling that is defective in osteoporosis, thus, with age, the bones become more and more porous and less able

to perform their mechanical function. These processes are regulated by growth factors, hormones, and cytokines [98]. Among the recently recognized factors that regulate the cross-talk between the bone-forming ostoeblasts and the bone-resorbing osteoclasts is the TNF receptor superfamily [99, 100]. RANK ligand (made by osteoblasts), also called OPGL and TRANCE, and its receptor RANK and the decoy receptor OPG (osteoprotegrin) control osteoclastogenesis. When RANKL binds to RANK, osteoclast development is triggered, but this can be blocked by another osteoblast product, OPG, which binds RANKL and thus blocks osteoclast formation. While many other signaling processes are involved, this is a clear example of cross-talk between these cells. The ways in which mechanical and other signals are conveyed to the osteoclast and osteoblast are still being probed. It is likely that, as these signals become better understood, more will also be learned about the fundamental aspects of bone formation.

Acknowledgments Dr. Boskey’s data described in this review was supported by NIH grants DE04141, AR 037661, and AR 041325.

References 1. Reith, E. J., and Ross, M. J. (1970). “Atlas of Descriptive Histology,” Second Edition. Harper and Row, Publishers, Hagerstown MD, pp. 23–40. 2. Wolff, J. (1986). “The Law of Bone Remodelling”. Translated from the original 1892 edition, by P. Maquet and R. Furlong. Springer-Verlag, Berlin. 3. Glimcher, M. J. (1998). The nature of the mineral phase in bone: biological and clinical implications. In “Metabolic Bone Disease and Clinically Related Disorders” (L. V. Avioli and S. M. Krane, eds.), Academic Press, San Diego, pp. 23–50. 4. Holden, J. L., Clement, J. G., and Phakey P. P. (1995). Age and temperature related changes to the ultrastructure and composition of human bone mineral. J. Bone Miner. Res. 10, 1400–1409. 5. Loong, C. K., Rey, C., Kuhn, L. T., Combes, C., Wu, Y., Chen, S., and Glimcher, M. J. (2000). Evidence of hydroxyl-ion deficiency in bone apatites: an inelastic neutron-scattering study. Bone 26, 599–602. 6. Tong, W., Glimcher, M. J., Katz, J. L., Kuhn, L., and Eppell, S. J. (2003). Size and shape of mineralites in young bovine bone measured by atomic force microscopy. Calcif. Tissue Int. 72, 592–598. 7. Fratzl, P., Schreiber, S., and Klaushofer, K. (1996). Bone mineralization as studied by small-angle x-ray scattering. Connect. Tissue Res. 34, 247–254. 8. Kim, H-M., Rey, C., and Glimcher, M. J. (1995). Isolation of calciumphosphate crystals of bone by nonaqueous methods of low temperature. J. Bone Miner. Res. 10, 1589–1660. 9. Landis, W. J., Hodgens, K. J., Arena, J., Song, M. J., and McEwen, B. F. (1996). Structural relations between collagen and mineral in bone as determined by high voltage electron microscopic tomography. Microsc. Res. Tech. 33, 192–202.

210 10. Boyde, A., and Jones, S. J. (1996). Scanning electron microscopy of bone: instrument, specimen, and issues. Microsc. Res. Tech. 33, 92–120. 11. Paschalis, E. P., DiCarlo, E., Betts, F., Sherman, P., Mendelsohn, R., and Boskey, A. L. (1996). FTIR microspectroscopic analysis of human osteonal bone. Calcif. Tissue Int. 59, 480–487. 12. Tarnowski, C. P., Ignelzi, M. A. Jr., and Morris, M. D. (2002). Mineralization of developing mouse calvaria as revealed by Raman microspectroscopy. J. Bone Miner. Res. 17, 1118–1126. 13. Roughley, P. J., Rauch, F., and Glorieux, F. (2003). Osteogenesis imperfecta-clinical and molecular diversity. Eur. Cell Mater. 5, 41–47. 14. Camacho, N. P., Raggio, C. L., Doty, S. B., Root, L., Zraick,V., Ilg, W.A., Toledano, T. R., and Boskey, A. L. (2001). A controlled study of the effects of alendronate in a growing mouse model of osteogenesis imperfecta. Calcif. Tissue Int. 69, 94–101. 15. Gokhale, J., Robey, P. G., and Boskey, A. L. (2001). “The Biochemistry of Bone”. In “Osteoporosis”, Second Edition (R. Marcus, D. Feldman and J. Kelsey, eds.), Academic Press, San Diego, Volume 1, pp. 107–189. 16. Reddi, A. H. (2000). Morphogenesis and tissue engineering of bone and cartilage: inductive signals, stem cells, and biomimetic biomaterials. Tissue Eng. 6, 351–349. 17. Schmid, T. M., Bonen, D. K., Luchene, L., and Linsenmayer, T. F. (1991). Late events in chondrocyte differentiation: hypertrophy, type X collagen synthesis and matrix calcification. In Vivo. 5, 533–540. 18. Anderson, H. C. (2003). Matrix vesicles and calcification. Curr. Rheumatol. Rep. 5, 222–226. 19. Ortega, N., Behonick, D., Stickens, D., and Werb, Z. (2003). How proteases regulate bone morphogenesis. Ann. N. Y. Acad. Sci. 995, 109–116. 20. Wu, L. N., Genge, B. R., Dunkelberger, D. G., LeGeros, R. Z., Concannon, B., and Wuthier, R. E. (1997). Physicochemical characterization of the nucleational core of matrix vesicles. J. Biol. Chem. 272, 4404–4411. 21. Buckwalter, J. A., Rosenberg, L. C. (1983). Structural changes during development in bovine fetal epiphyseal cartilage. Collagen Relat. Res. 3, 489–504. 22. Cowles, E. A., DeRome, M. E., Pastizzo, G., Brailey, L. L., and Gronowicz, G. A. (1998). Mineralization and the expression of matrix proteins during in vivo bone development. Calif. Tissues Int. 62, 74–82. 23. Ramstad, V. E., Franzen, A., Heinegard, D., Wendel, M., and Reinholt, F. P. (2003). Ultrastructural distribution of osteoadherin in rat bone shows a pattern similar to that of bone sialoprotein. Calcif. Tissue Int. 72, 57–64. 24. Vogel, K. G., Paulsson, M., Heinegard, D. (1984). Specific inhibition of type I and type II collagen fibrillogenesis by the small proteoglycan of tendon. Biochem. J. 223, 587–597. 25. Kuboki, Y., Kudo, A., Mizuno, M., and Kawamura, M. (1992). Timedependent changes of collagen cross-links and their precursors in culture of osteogeneic cells. Calcif. Tissue Int. 50, 473–480. 26. Eyre, D. R., Dickson, I. R., and Van Ness, K. (1988). Collagen crosslinking in human bone and articular cartilage. Age–related changes in the content of mature hydroxypyridinium residues. Biochem. J. 252, 495–500. 27. Gerstenfeld, L. C., Chipman, S. D., Glowacki, J., and Lian, J. B. (1986). Expression to differentiate function by mineralizing cultures of chicken osteoblasts. Dev. Biol. 122, 49–60. 28. Hauschka, P. V., and Wians, F. H., Jr. (1989). Osteocalcin–hydroxyapatite interaction in the extracellular organic matrix of bone. Anat. Rec. 224, 180–188. 29. Wu, Y., Ackerman, J. L., Strawich, E. S., Rey, C., Kim, H. M., and Glimcher, M. J. (2003). Phosphate ions in bone: identification of a calcium-organic phosphate complex by 31P solid-state NMR spectroscopy at early stages of mineralization. Calcif. Tissue Int. 72, 610–626.

ADELE L. BOSKEY 30. Mbuyi–Muamba, J. M., Dequeker, J., and Gevers, G. (1989). Collagen and noncollagenous proteins in different mineralization stages of human femur. Acta Anat. (Basel). 134, 265–268. 31. Lane, J. M., Betts, F., Posner, A. S., and Yue, D. W. (1984). Mineral parameters in early fracture repair. J. Bone Joint Surg. [Am] 66, 1289–1293. 32. Boskey, A. L., and Marks, S. C. (1985). Mineral and matrix alterations in the bones of incisors–absent (ia/ia). Osteopetrotic rat. Calcif. Tissue Int. 37, 287–292. 33. Kuhn, L. T., Wu, Y., Rey, C., Gerstenfeld, L. C., Grynpas, M. D., Ackerman, J. L., Kim, H. M., and Glimcher, M. J. (2000). Structure, composition, and maturation of newly deposited calcium-phosphate crystals in chicken osteoblast cell cultures. J. Bone Miner. Res. 15, 1301–1309. 34. Mendelsohn, R., Hassenkhani, A., DiCarlo, E., and Boskey, A. L. (1989). FT-IR microscopy of endochondral ossification at 20µ spatial resolution. Calcif. Tissue Int. 44, 20–24. 35. Weiss, I. M., Tuross, N., Addadi, L., and Weiner, S. (2002). Mollusc larval shell formation: amorphous calcium carbonate is a precursor phase for aragonite. J. Exp. Zool. 293, 478–491. 36. Driessens, F. C. M., and Verbeeck, R. M. H. (1990). “Biominerals,” CRC Press, Boca Raton, FL., Ch 3, pp. 37–61. 37. Traub, W., Arad, T., and Weiner, S. (1992). Origin of mineral crystal growth in collagen fibrils. Matrix. 12, 251–255. 38. Garimella, R., Bi, X., Camacho, N., Sipe, J. B., and Anderson, H. C. (2004). Primary culture of rat growth plate chondrocytes: an in vitro model of growth plate histotype, matrix vesicle biogenesis and mineralization. Bone 34, 961–970. 39. Eanes, E. D., Hailer, A. W., Midura, R. J., and Hascall, V. C. (1992). Proteoglycan inhibition of calcium phosphate precipitation in liposomal suspensions. Glycobiology 2, 571–578. 40. Boskey, A. L., Maresca, M., Armstrong, A. L., and Ehrlich, M. G. (1992). Treatment of proteoglycan aggregates with growth plate neutral metalloprotease reduces their ability to inhibit hydroxyapatite proliferation in a gelatin gel. J. Orthop. Res. 10, 313–319. 41. Embery, G., Hall, R., Waddington, R., Septier, D., and Goldberg, M. (2001). Proteoglycans in dentinogenesis. Crit. Rev. Oral Biol. Med. 12, 331–349. 42. Cornelissen, M., and de Ridder, L. (1993). Electron microscopic study of proteoglycan aggregates synthesized by embryonic chick sternal chondrocytes in vitro. Eur. J. Morphol. 31, 17–20. 43. Dean, D. D., Boyan, B. D., Muniz, O. E., Howell, D. S., and Schwartz, Z. (1996). Vitamin D metabolites regulate matrix vesicle metalloproteinase content in a cell maturation–dependent manner. Calcif. Tissue Int. 59, 109–116. 44. Anderson, H. C., Sipe, J. B., Hessle, L., Dhanyamraju, R., Atti, E., Camacho, N. P., and Millan, J. L. (2004). Impaired calcification around matrix vesicles of growth plate and bone in alkaline phosphatasedeficient mice. Am. J. Pathol. 164, 841–847. 45. Luckman, S. P., Rees, E., and Kwan, A. P. (2003). Partial characterization of cell-type X collagen interactions. Biochem. J. 372, 485–493. 46. Landis, W. J., Hodgens, K. J., McKee, M. D., Nanci, A., Song, M. J., Kiyonaga, S., Arena, J., and McEwen, B. (1992). Extracellular vesicles of calcifying turkey leg tendon characterized by immunocytochemistry and high voltage electron microscopic tomography and 3–D graphic image reconstruction. Bone Miner. 17, 237–241. 47. Mocetti, P., Ballanti, P., Zalzal, S., Silvestrini, G., Bonucci, E., and Nanci, A. (2000). A histomorphometric, structural, and immunocytochemical study of the effects of diet-induced hypocalcemia on bone in growing rats. J. Histochem. Cytochem. 48, 1059–1078. 48. Glimcher, M. J., Hodge, A. J., and Schmitt, F. O. (1957). Macromolecular aggregation states in relation to mineralization: the collagen hydroxyapatite system as studied in vitro. Proc. Natl Acad. Sci. U.S.A. 43, 860–967.

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Chapter 12 Mineralization, Structure and Function of Bone 49. Mergenhagen, S. E., Martin, G. R., Rizzo, A., Wright, D. N., and Scott, D. B. (1960). Calcification in vivo of implanted collagens. Biochim. Biophys. Acta. 43, 563–565. 50. Termine, J. D., Kleinman, H. K., Whitson, S. W., Conn, K. M., McGarvey, M. L., and Martin, G. R. (1981). Osteonectin, a bonespecific protein linking mineral to collagen. Cell. 26, 99–105. 51. Endo, A., and Glimcher, M. J. (1989). The effect of complexing phosphoproteins to decalcified collagen on in vitro calcification. Connect. Tissue Res. 21, 179–190. 52. de Bri, E., Reinholt, F. P., Heinegard, D., Mengarelli-Widholm, S., Norgard, M., and Svensson, O. (1996). Bone sialoprotein and osteopontin distribution at the osteocartilaginous interface. Clin. Orthop. Relat. Res. 330, 251–260. 53. Stein, G. S., and Lian, J. B. (1993). Molecular mechanisms mediating proliferation/differentiation interrelationships during progressive development of the osteoblast phenotype. Endocr. Rev. 14, 424–442. 54. Young, M. F., Bi, Y., Ameye, L., and Chen, X. D. (2002). Biglycan knockout mice: new models for musculoskeletal diseases. Glycoconj. J. 19, 257–262. 55. McKee, M. D., and Nanci, A. (1995). Postembedding colloidal-gold immunocytochemistry of noncollagenous extracellular matrix proteins in mineralized tissues. Microsc. Res. Tech. 31, 44–62. 56. Fisher, L. W., and Fedarko, N. S. (2003). Six genes expressed in bones and teeth encode the current members of the SIBLING family of proteins. Connect. Tissue Res. 44, 33–40. 57. Jain, A., Karadag, A., Fohr, B., Fisher, L. W., and Fedarko, N. S. (2002). Three SIBLINGs (small integrin-binding ligand, N-linked glycoproteins) enhance factor H’s cofactor activity enabling MCP-like cellular evasion of complement-mediated attack. J. Biol. Chem. 277, 13700–13708. 58. Fedarko, N. S., Jain, A., Karadag, A., and Fisher, L. W. (2004). Three small integrin binding ligand N-linked glycoproteins (SIBLINGs) bind and activate specific matrix metalloproteinases. FASEB J. 18, 734–746. 59. Wu, L. N., Sauer, G. R., Genge, B. R., Valhmu, W. B., and Wuthier, R. E. (2003). Effects of analogues of inorganic phosphate and sodium ion on mineralization of matrix vesicles isolated from growth plate cartilage of normal rapidly growing chickens. J. Inorg. Biochem. 94, 221–235. 60. Pampena, D. A., Robertson, K. A., Litvinova, O., Lajoie, G., Goldberg, H. A., and Hunter, G. K. (2004). Inhibition of hydroxyapatite formation by osteopontin phosphopeptides. Biochem. J. 378, 1083–1087. 61. Silverman, L., and Boskey, A. L. (2004). Diffusion systems for evaluation of biomineralization. Calcified Tissue Int. 75, 494–501. 62. Ganss, B., Kim, R. H., and Sodek, J. (1999). Bone sialoprotein. Crit. Rev. Oral Biol. Med. 10, 79–98. 63. Hunter, G. K., and Goldberg, H. A. (1993). Nucleation of hydroxyapatite by bone sialoprotein. Proc. Natl Acad. Sci. USA 90, 8562–8565. 64. Midura, R. J., Wang, A., Lovitch, D., Law, D., Powell, K., and Gorski, J. P. (2004). Bone acidic glycoprotein-75 delineates the extracellular sites of future bone sialoprotein accumulation and apatite nucleation in osteoblastic cultures. J. Biol. Chem. 279, 25464–25473. 65. Stubbs, J. T. 3rd, Mintz, K. P., Eanes, E. D., Torchia, D. A., and Fisher, L. W. (1997). Characterization of native and recombinant bone sialoprotein: delineation of the mineral-binding and cell adhesion domains and structural analysis of the RGD domain. J. Bone Miner. Res. 12, 1210–1222. 66. Fujisawa, R., Wada, Y., Nodasaka, Y., and Kuboki, Y. (1996). Acidic amino acid–rich sequences as binding sites of osteonectin to hydroxyapatite crystals. Biochim. Biophys. Acta. 1292, 53–60. 67. Tye, C. E., Rattray, K. R., Warner, K. J., Gordon, J. A., Sodek, J., Hunter, G. K., and Goldberg, H. A. (2003). Delineation of the hydroxyapatite-nucleating domains of bone sialoprotein. J. Biol. Chem. 278, 7949–7955. 68. Boskey, A. L., Spevak, L., Doty, S. B., and Rosenberg, L. (1997). Effects of bone CS-proteoglycans, DS-decorin, and DS-biglycan on

69.

70.

71.

72.

73.

74.

75.

76.

77.

78.

79.

80.

81.

82.

hydroxyapatite formation in a gelatin gel. Calcif. Tissue Int. 61, 298-305. Tartaix, P. H., Doulaverakis, M., George, A., Fisher, L. W., Butler, W. T., Qin, C., Salih, E., Tan, M., Fujimoto, Y., Spevak, L., and Boskey, A. L. (2004). Dentin matrix protein-1: In vitro effects on hydroxyapatite formation provide insights into in vivo functions. J. Biol. Chem. 279, 18115–18120. Boskey, A. L., Maresca, M., Doty, S., Sabsay, B., and Veis, A. (1990). Concentration dependent effects of dentin-phosphophoryn in the regulation of in vitro hydroxyapatite formation and growth. Bone Mineral. 11, 55–65. Boskey, A. L., Maresca, M., Ulrich, W., Doty, S. B., Butler, W. T., and Prince, C. W. (1993). Osteopontin-hydroxyapatite interactions in vitro: Inhibition of hydroxyapatite formation and growth in a gelatin-gel. Bone Miner. 22, 147–159. Hunter, G. K., Kyle, C. L., and Goldberg, H. A. (1994). Modulation of crystal formation by bone phosphoproteins: structural specificity of the osteopontin-mediated inhibition of hydroxyapatite formation. Biochem. J. 300, 723–728. Rowe, P. S., Kumagai, Y., Gutierrez, G., Garrett, I. R., Blacher, R., Rosen, D., Cundy, J., Navvab, S., Chen, D., Drezner, M. K., Quarles, L. D., and Mundy, G. R. (2004). MEPE has the properties of an osteoblastic phosphatonin and inhibin. Bone 34, 303–319. Prince, M., Banerjee, C., Javed, A., Green, J., Lian, J. B., Stein, G. S., Bodine, P. V., and Komm, B. S. (2001). Expression and regulation of Runx2/Cbfa1 and osteoblast phenotypic markers during the growth and differentiation of human osteoblasts. J. Cell. Biochem. 80, 424–440. Boskey, A. L., Stiner, D., Binderman, I., and Doty, S. B. (1997). Effects of proteoglycan modification on mineral formation in a differentiating chick limb-bud mesenchymal cell culture system. J. Cell Biochem. 64, 632–643. Boskey, A. L., Guidon, P., Doty, S. B., Stiner, D., Leboy, P., and Binderman, I. (1996). The mechanism of ß-glycerophosphate action in mineralizing chick limb-bud mesenchymal cells. J. Bone Min. Res. 11, 1694–1702. Luppen, C. A., Leclerc, N., Noh, T., Barski, A., Khokhar, A., Boskey, A. L., Smith, E., and Frenkel, B. (2003). Brief bone morphogenetic protein 2 treatment of lucocorticoid-inhibited MC3T3-E1 osteoblasts rescues commitment-associated cell cycle and mineralization without alteration of Runx2. J. Biol. Chem. 278, 44995– 45003. Thomas, G., Moffatt, P., Salois, P., Gaumond, M. H., Gingras, R., Godin, E., Miao, D., Goltzman, D., and Lanctot, C. (2003). Osteocrin, a novel bone-specific secreted protein that modulates the osteoblast phenotype. J. Biol. Chem. 278, 50563–50571. Schinke, T., Amendt, C., Trindl, A., Poschke, O., Muller-Esterl, W., and Jahnen-Dechent, W. (1996). The serum protein alpha2-HS glycoprotein/fetuin inhibits apatite formation in vitro and in mineralizing calvaria cells. A possible role in mineralization and calcium homeostasis. J. Biol. Chem. 271, 20789–20796. Stacey, A., Bateman, J., Choi, T., Mascara, T., Cole, W., and Jaenisch, R. (1988). Perinatal lethal osteogenesis imperfecta in transgenic mice bearing an engineered mutant pro-alpha 1(i) collagen gene. Nature 332, 131–136. Gowen, L. C., Petersen, D. N., Mansolf, A. L., Qi, H., Stock, J. L., Tkalcevic, G. T., Simmons, H. A., Crawford, D. T., Chidsey-Frink, K. L., Ke, H. Z., McNeish, J. D., and Brown, T. A. (2003). Targeted disruption of the osteoblast/osteocyte factor 45 gene (OF45) results in increased bone formation and bone mass. J. Biol. Chem. 278, 1998–2007. Forlino, A., Porter, F. D., Lee, E. J., Westphal, H., and Marini, J. C. (1999). Use of the Cre/lox recombination system to develop a nonlethal knock-in murine model for osteogenesis imperfecta with an alpha1(I) G349C substitution. Variability in phenotype in BrtlIV mice. J. Biol. Chem. 274, 37923–37931.

212 83. Watanabe, H., and Yamada, Y. (1999). Mice lacking link protein develop dwarfism and craniofacial abnormalities. Nat. Genet. 21, 225–229. 84. Panda, D. K., Miao, D., Tremblay, M. L., Sirois, J., Farookhi, R., Hendy, G. N., and Goltzman, D. (2001) Targeted ablation of the 25-hydroxyvitamin D 1alpha -hydroxylase enzyme:evidence for skeletal, reproductive, and immune dysfunction. Proc. Natl Acad. Sci. USA 98, 7498–7503. 85. Xu, T., Bianco, P., Fisher, L. W., Longenecker, G., Smith, E., Goldstein, S., Bonadio, J., Boskey, A., Heegaard, A. M., Sommer, B., Satomura, K., Dominguez, P., Zhao, C., Kulkarni, A. B., Robey, P. G., and Young, M. F. (1998). Targeted disruption of the biglycan gene leads to an osteoporosis-like phenotype in mice. Nat. Genet. 20, 78–82. 86. Boskey, A. L., Spevak, L., Paschalis, E., Doty, S. B., McKee, M. D. (2002). Osteopontin deficiency increases mineral content and mineral crystallinity in mouse bone. Calcif. Tissue Int. 71, 145–154. 87. Boskey, A. L., Karsenty, G., and McKee, M. D. (2000). Mineral characterization of bones and soft tissues in matrix gla protein deficient mice. In “Chemistry and Biology of Mineralized Tissues” (M. Goldberg, A. Boskey, C. Robinson, eds.), Am. Acad. Orthopaed. Surgeons. Chicago, IL, pp. 63–67. 88. Ducy, P., Desbois, C., Boyce, B., Pinero, G., Story, B., Dunstan, C., Smith, E., Bonadio, J., Goldstein, S., Gundberg, C., Bradley, A., and Karsenty, G. (1996). Increased bone formation in osteocalcin– deficient mice. Nature 382, 448–452. 89. Boskey, A. L., Gadaleta, S., Gundberg, C., Doty, S. B., Ducy, P., and Karsenty, G. (1998). FT-IR microspectroscopic analysis of bones of osteocalcin-deficient mice provided insight into the function of osteocalcin. Bone 23, 187–196. 90. Boskey, A. L., Moore, D. J., Amling, M., Canalis, E., Delany, A. M. (2003). Infrared analysis of the mineral and matrix in bones of osteonectin-null mice and their wildtype controls. J. Bone Miner. Res. 18, 1005–1011. 91. Paschalis, E. P., Jacenko, O., Olsen, B., Mendelsohn, R., and Boskey, A. L. (1996). Fourier transform infrared microspectroscopic analysis identifies alterations in mineral properties in bones from mice transgenic for type X collagen. Bone 19, 151–156

ADELE L. BOSKEY 92. Giachelli, C. M., and Steitz, S. (2000). Osteopontin: a versatile regulator of inflammation and biomineralization. Matrix Biol. 19, 615–622. 93. Denhardt, D. T., Noda, M., O’Regan, A. W., Pavlin, D., Berman, J.S. (2001). Osteopontin as a means to cope with environmental insults: regulation of inflammation, tissue remodeling, and cell survival. J. Clin. Invest. 107(9); 1055–1061. 94. Dequeker, J., and Mundy, G. R. (1998). Bone structure and function. In “Rheumatology,” Second Edition (J. H. Klippel and P. A. Dieppe, eds), Mosby, London, pp. 34:1–12. 95. Bigi, A., Cojazzi, G., Panzavolta, S., Ripamonti, A., Roveri, N., Romanello, M., Noris Suarez, K., and Moro, L. (1997). Chemical and structural characterization of the mineral phase from cortical and trabecular bone. J. Inorg. Biochem. 68, 45–51. 96. Jilka, R. L. (2003). Biology of the basic multicellular unit and the pathophysiology of osteoporosis. Med. Pediatr. Oncol. 41, 182–185. 97. Yoshitake, H., Rittling, S. R., Denhardt, D. T., and Noda, M. (1999). Osteopontin-deficient mice are resistant to ovariectomy-induced bone resorption. Proc. Natl Acad. Sci. USA 96, 8156–8160. 98. Raisz, L. G. (1999). Physiology and pathophysiology of bone remodeling. Clin. Chem. 45, 1353–1358. 99. Cheng, X., Kinosaki, M., Murali, R., and Greene, M. I. (2003). The TNF receptor superfamily: role in immune inflammation and bone formation. Immunol. Res. 27, 287–294. 100. Boyle, W. J., Simonet, W. S., and Lacey, D. L. (2003). Osteoclast differentiation and activation. Nature 423, 337–342. 101. Hashimoto-Gotoh, T., Ohnishi, H., Tsujimura, A., Tsunezuka, H., Imai, K., Masuda, H., and Nakamura, T. (2004). Bone mass increase specific to the female in a line of transgenic mice overexpressing human osteoblast stimulating factor-1. J. Bone Miner. Metab. 22, 278–282. 102. Kozloff, K. M., Carden, A., Bergwitz, C., Forlino, A., Uveges, T. E., Morris, M. D., Marini, J. C., and Goldstein, S. A. (2004). Brittle IV mouse model for osteogenesis imperfecta IV demonstrates postpubertal adaptations to improve whole bone strength. J. Bone Miner. Res. 19, 614–622.

Chapter 13

Bone Structure and Strength Ego Seeman BSc MBBS FRACP MD

V. Strength Maintenance VI. Conclusion References

I. Introduction II. Gravity and the Need for Stiffness, Flexibility, Lightness and Speed III. The Material Composition and Structural Design of Bone IV. Bone Modeling and Remodeling – The Mechanism of Bone’s Construction during Growth and Decay with Advancing Age

I. INTRODUCTION

its structure. A negative bone balance between the volumes of bone resorbed and formed in each remodeling transaction is the morphological basis of irreversible bone loss, a process that compromises structure. Understanding the mechanisms of adaptation and challenges to it requires attention to the individual components of bone’s material composition and structural design so that the regulators and co-regulators of this machinery can be identified. With this approach, rational approaches to intervention can be designed to prevent and to restore structural compromise and ensuring fragility fractures.

Structural failure, another way of referring to skeletal fracture, is a problem in biomechanics. Understanding structural failure, therefore, requires the study of structure. Currently available noninvasive techniques provide a measure of bone mineral mass in a region of interest but they provide little information about bone’s material composition and structural design, two critical determinants of strength [1]. It is self-evident that structure determines tolerable loads but it is less evident as loads also determine structure. Bone has the ability to modify its material composition and structural design to accommodate prevailing loads [2]. This is achieved by the cellular machinery of adaptive modeling and remodeling. Adaptation is most successful when bones are in their growth phase, but with maturation of the skeleton maintaining bone strength is a greater challenge, particularly when aging leads to sex hormone deficiency, secondary hyperparathyroidism, reduced physical activity, loss of muscle mass, and other factors. Age-related abnormalities in the cellular machinery of bone itself can also contribute to the compromised ability of bone to adapt. Alterations in the rate of remodeling can compromise material properties of bone and also indirectly Dynamics of Bone and Cartilage Metabolism

Dept of Endocrinology and Medicine, Austin Hospital, University of Melbourne, Melbourne, Australia.

II. GRAVITY AND THE NEED FOR STIFFNESS, FLEXIBILITY, LIGHTNESS AND SPEED Land-dwelling mammals must be able to move to catch dinner, and move quickly to avoid becoming it. Propulsion against gravity requires levers. Bones are levers and levers must be stiff. They must resist deformation but they must also be light enough to allow rapidity of movement. Impact loading imparts energy to bone. As energy cannot be destroyed, it must be absorbed by bone, a process 213

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EGO SEEMAN

requiring flexibility. Bone must be a spring, able to change shape, deform without cracking, shorten and widen in compression, lengthen and narrow in tension. The elastic properties of bone allow it to absorb energy by deforming reversibly when loaded [2]. If the load imposed exceeds bone’s ability to deform elastically, it can deform further, yielding by changing shape permanently in plastic deformation. The permanent change in shape is associated with microcracks that allow energy release, a compromise of structure that is a final defense against the alternative – complete fracture. Provided these microcracks are small and do not extend in length, bone can remain intact. If both the elastic and plastic zones are exceeded, the only way the energy can be released is fracture (producing two pieces of bone). Bone achieves its strength in each of these contradictory properties – stiffness yet flexibility, lightness yet strength, by its material composition and structural design.

III. THE MATERIAL COMPOSITION AND STRUCTURAL DESIGN OF BONE The right balance between material stiffness and flexibility is achieved by varying the degree of mineral content of

the bone tissue [1]. The greater the mineral content, the greater the material stiffness but the lower its flexibility (Fig. 1). Through millions of years, nature has selected characteristics that are most suited to the particular function bone usually performs. Ossicles of the ear are densely mineralized to be stiff and vibrate like tuning forks without loss of energy in deformation. In contrast, deer antlers are less densely mineralized and so can deform like springs during head butting in the mating season [2] (Fig. 2). The fabric of bone is configured as a structure. Structural stiffness and flexibility (as opposed to material stiffness and flexibility), and lightness, are achieved by geometry and architectural design – not just its mass. Indeed, for tubular bones, larger and smaller bones within a species do not differ by the amount of material used to construct them. Although counterintuitive, there is no correlation between the external volume of long bones and bone mass. The same amount of material is used to build a wider and longer tubular bone by fashioning it with a thinner cortex [3]. This is an example of minimizing mass for optimal function. Nature can fashion a bigger bone using the same material by creating a bigger hole in the middle of the bone. Larger bones have a larger medullary canal, constructed during intrauterine and postnatal life by excavation of this

Figure 1 Increasing tissue mineral content increases material stiffness but decreases the bone material’s ability to absorb energy without cracking (toughness) (data provided by courtesy of JD Currey).

Chapter 13 Bone Structure and Strength

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Figure 2

Bones requiring stiffness like auditory ossicles are selected for high mineral content, they are very brittle. Bones requiring flexibility like antlers are selected for lower mineral content (adapted from JD Currey [2]).

cavity by bone resorption. Larger bones have a lower apparent volumetric BMD than smaller bones. The further displacement of the slightly thinner cortex confers greater resistance to bending (structural stiffness) because bending strength is proportional to the fourth power of the radius [4]. In long bones, structures that are effectively levers, stiffness is favored over flexibility.

Figure 3 During growth before puberty periosteal apposition increases long bone diameter and exceeds endocortical resorption, so the bone develops a thicker and thicker cortex. Cortical thickness is similar by sex because reduced endocortical resorption or increased endocortical apposition during puberty contribute to final cortical thickness in females. After epiphyseal closure, endocortical resorption now exceeds the slow periosteal apposition, producing cortical thinning.

A. Building Levers As long bones grow in length, periosteal apposition increases its diameter while concurrent endocortical resorption excavates the marrow cavity (Fig. 3). As periosteal apposition exceeds net endocortical resorption the lengthening bone develops a wider and wider cortex. The enlarging medullary canal effectively shifts the thickening cortex further and further from the long bone neutral axis, producing the structural stiffness needed for lever function. In females, earlier completion of longitudinal growth and earlier inhibition of periosteal apposition produces a smaller bone. However, cortical thickness is similar across both sexes because a reduction in endocortical resorption and perhaps net endocortical apposition contributes to final cortical thickness [5, 6]. What differs among the sexes, therefore, is the position of the cortex in relationship to the long axis of the long bone. Likewise, between races, the main difference is the position of the cortex in relationship to the long axis of the long bone, not the cortical thickness per se. The absolute and relative growth of periosteal and endocortical surfaces varies by sex, pubertal stage, the type of bone, and varies in degree, in fact, at every position and surface along the bone. At each level, these traits are

determined by the interaction between bone formation on the periosteum, endocortical bone resorption, and formation adjacent to marrow. Long bones are not drinking straws with the same diameter throughout and the same thickness of cortex throughout. The complex and irregular periosteal and endocortical perimeters of long bone shafts create elliptical and triangular structures, adapted locally to the loads they are exposed to and adapted to tolerate. These loads then determine extent and degrees of local modeling and remodeling in adjacent regions. The process of adaptation involves using the same material but fashioning and refashioning the same amount of bone into the shape and contours best suited to tolerated prevailing loads. This notion is important and can be seen in the complex shape of the femoral neck. Near the shaft, where bending moments are greatest the shape is elliptical and the cortex is thicker inferiorly than superiorly. The mass of mineralized bone is mainly cortical. Moving proximally along the femoral neck the amount of bone material remains constant but the shape of the femoral neck is more circular, the proportion of trabecular bone increases and the cortical thickness is similar superiorly and inferiorly, features adapted to the loading pattern present which is more shear

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and compressive than bending [3]. The loading circumstances are changing geometry not bone mass. While this process is successful during growth, it is not very successful after epiphyseal closure has occurred.

B. Building Springs In the axial skeleton (e.g. vertebral body of the spine), bone is constructed differently but also demonstrates the effective use of void space to achieve lightness with a minimal amount of material. The vertebral bodies serve more as springs or shock absorbers than levers. The structure is light yet “strong” in a different sense of the word; these bones are “stronger” in their ability to deform without cracking. While tubular bones can deform by only 1–2% of their original length, vertebral bodies can deform more substantially by absorbing energy but they sacrifice peak stresses tolerated (load per unit area) in favor of greater peak strain (change in length/original length). The ability to deform facilitates flexion, extension, and rotation of the whole vertebral skeleton of the upper body (Fig. 4). For the vertebrae, increasing bone size by periosteal apposition builds a wider vertebral body in males than females and in some races than others. The number of trabeculae, established at the growth plates, does not increase with age [7]. At puberty, trabecular BMD increases by increasing the size and thickness of the trabeculae plates and sheets to a similar degree in boys and girls so that males and females have the same vertebral body trabecular BMD (number and thickness of trabeculae) during prepubertal and pubertal growth, and at peak young

Figure 4

Cortical bone tolerates greater loads (“stronger” in load bearing) at the price of less flexibility while trabecular bone is able to deform more (“stronger” in being more flexible) but sacrifices peak loading ability.

adulthood [8, 9]. Thus, growth does not build a “denser” skeleton in males than females, it builds a bigger skeleton. Strength of the vertebral body is greater in young males than females due to sex differences in size, not BMD.

IV. BONE MODELING AND REMODELING – THE MECHANISM OF BONE’S CONSTRUCTION DURING GROWTH AND DECAY WITH ADVANCING AGE That structure determines the loads that can be tolerated is obvious. The shape of bone is contained within the ancestral genetic code selected for survival in an anticipated environment; fetal lower limb buds removed and grown in vitro, form the shape of the proximal femur [10]. The concept in reverse, namely that loads determine structure, is less obvious but central to understanding the pathogenesis of bone fragility. Bone accommodates load in given circumstances, by adaptive modeling and remodeling. This process occurs throughout the whole of life, but its purpose differs during growth (when the epiphyses are open) and during adulthood (when the epiphyses are closed). During growth, through the processes of bone modeling and remodeling, remarkable adaptations to loading can be achieved. A wealth of literature on skeletal morphology in elite athletes supports this premise [11]. Variability in loading and the forces associated with it help to account for the complex shape of bone described above. For example, in animal studies, collagen abnormalities in the MOV13 mutant mouse result in reduced bone strength but vigorous adaptive modeling by periosteal apposition results in bone strength above that seen in the wild-type [12]. Likewise, in other mouse models of osteogenesis, imperfecta adaptations take place in material composition rather than its structure [13]. The process of adaptive modeling and remodeling can compensate for abnormalities in one trait that reduce whole bone strength to modify another trait to preserve whole bone strength. The compensatory effects are limited. If the severity of the abnormality is too great [14], or the compensatory mechanism is itself abnormal, then adaptation is incomplete, resulting in more fragile bones. The emergence of bone fragility during aging can be viewed in this way. Over many years, accumulating abnormalities due to disease, hormonal deficiency and/or excess, exposure to risk factors such as tobacco and excess alcohol, as well as abnormalities in the machinery of bone modeling and remodeling itself, produce changes in the material composition of bone and its structure that together form the

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basis of bone fragility. Viewed in this way, bone fragility, or osteoporosis, can be regarded as a disorder of failed adaptation.

V. STRENGTH MAINTENANCE After completion of longitudinal growth (closure of the epiphyses), bone modeling and remodeling help to maintain bone strength, rather than to increase it. After bone size has been achieved during the growth period, periosteal apposition ceases to be vigorous and the remodeling rate, reflected in the levels of the biochemical measures of bone remodeling, declines. The remodeling rate is rapid during growth because each remodeling event deposits only a small amount of bone as bone is constructed [15]. Whether there is a relationship between the rate of remodeling and the degree of positive BMU balance is not known. It is possible that smaller positive balances can be compensated for by higher rates of remodeling or vice versa. As growth nears its “programmed” completion rapid remodeling is no longer needed and a positive balance in each remodeling unit is also no longer needed. The remodeling rate slows down and there may be a progressive lessening of the degree of positive balance at the level of the BMU. The first “abnormality” signaling the onset of a change in bone structure may be a reduction in bone formation at the cellular level. Information regarding this is controversial and inconsistent [16–18]. Evidence for a decline in the volume of bone formed is documented at midlife (around 50 years of age), but there is evidence for a decline in peak bone mineral mass and trabecular bone volume well before the menopause [19–21]. After epiphyseal closure, rapid periosteal apposition ceases to the point where bone enlarges no more than a few millimeters over the next 60 years [22–24]. Whereas during growth periosteal apposition is greater than net endocortical resorption, producing an increasingly thicker cortex, during the aging process, endocortical resorption exceeds periosteal apposition. The result of this switch in relative activities of periosteal apposition, declining, and endocortical resorption, maintained or increasing, is a thinner cortex with slight increases in bone diameter. Negative bone balance within each BMU is the metabolic basis of bone loss. On the endocortical surface, this process produces cortical thinning while bone loss on the intracortical surface produces intracortical porosity. The increasing porosity of cortical bone effectively trabecularizes the cortex [25, 26]. The surface or volume ratio increases so that remodeling continues vigorously in cortical bone, producing cortical fragility. The same loads on bone are imposed on a structure diminished

Figure 5

Trabecular bone loss by loss of trabecular numbers and so loss of connectivity (as occurs in women) produces a greater loss of strength than trabecular bone loss by thinning (as occurs in men). Adapted from Ref [28].

in cross-sectional diameter so that the stresses on bone (load per unit area) increase predisposing to buckling, microdamage and ultimately fracture. The amount of trabecular bone lost during aging in women and men is similar, or only slightly less in men than women [4, 5]. However, trabecular bone loss occurs mainly by thinning in men and mainly by loss of connectivity in women [27] (Fig. 5). Strength of the vertebrae decreases more when there is loss of connectivity rather than thinning predisposing to fractures [28]. Loss of connectivity is the result of the accelerated loss of bone that occurs in midlife in women due to estrogen deficiency. Estrogen deficiency is associated with increased remodeling on the endosteal surface. In addition, imbalance in the BMU increases as estrogen deficiency increases the lifespan of osteoclasts and reduces the lifespan of osteoblasts. Increased numbers of BMUs, coupled with increased resorption depth, is responsible for the loss of connectivity seen in women [29]. The contribution of trabecular bone loss to overall bone loss decreases as trabecular plates perforate and disappear because there is less trabecular surface available for remodeling. In men, there is no midlife acceleration of bone remodeling because men do not go through an abrupt period of sex steroid deficiency, but bone loss, nevertheless, occurs due to reduced bone formation and thinning of trabeculae. Relatively greater maintenance of connectivity results in persistence of the trabecular surfaces available for remodeling so that trabecular bone loss probably continues longer in men than in women.

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That is why final trabecular bone volume is similar in men and women. The amount of trabecular surface available for bone remodeling in old age appears to be greater in elderly men than women [27].

A. Periosteal Bone Formation During Aging The periosteal bone formation during aging offsets bone loss from the endosteal surfaces in both men and women. Periosteal apposition may be greater at some sites in men than in women so that the similar loss of bone from the endosteal surface is more greatly offset in men than in women [30, 31]. Thus, the smaller decline in BMD at the spine in men than women is due to the smaller reduction in cortical bone in men than in women. However, the better maintenance of cortical mass in men is the result of greater periosteal bone formation, not less resorptive removal of bone from the endosteal surface [Fig. 6].

B. Why Fewer Men than Women Sustain Fragility Fractures The larger skeleton in men produces a bone that tolerates larger absolute loads. However, the load per unit area (stress) imposed on the vertebral body is no different in young men and women, because the larger bone is subjected to correspondingly larger loads. There is scaling in nature – a bigger bone has a bigger muscle, a relationship that is likely to be largely determined by genes regulating size, not necessarily the amount of exercise undertaken. Fragility fractures are uncommon in young men and women because loads are less than the ability of the bone to withstand them. Structural failure emerges during aging because of the changing relationship between the imposed load and the bone’s ability to tolerate that load. Periosteal apposition increases the cross-sectional diameter of the bone more in men, so that load imposed per unit area decreases more than in women. Greater periosteal

Figure 6 Subperiosteal area increases more in men than in women due to greater periosteal apposition in men. Medullary expansion with age is similar in men and women. The net effect is better maintenance of cortical area and strength in men than in women (adapted from Ruff and Hayes [4]).

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apposition adds more bone to the outer perimeter of the bone in men than women, offsetting the age-related endosteal bone loss more in men than women so that the fall in volumetric BMD is less in men than in women. During aging, the stress on bone decreases more in men, and bone strength decreases less, so that a lower proportion of elderly men than elderly women have bone size and architectural and material properties such as microdamage, tissue mineral density, loss of connectivity, porosity, trabecular and cortical thinning below a critical level (or fracture threshold) where the loads on the bone are greater than the bone’s ability to tolerate them. Structural failure occurs less in men than women because the relationship between load and bone strength is better maintained in men than in women.

4. 5.

6.

7.

8.

9.

10.

VI. CONCLUSION

11. 12.

Bones must be stiff so that they do not bend when loaded. Bones must also be flexible, able to absorb energy by elastic and plastic deformation. Structural failure may occur if bones deform too little or too much. Age-related and menopause-related abnormalities in bone remodeling produce loss of the material and structural properties that no longer keep bone “just right”. High remodeling reduces the mineral content of bone tissue resulting in loss of stiffness. Sex hormone deficiency increases the volume of bone resorbed and reduces the volume of bone formed in each BMU. The contributions made by differences in material composition (tissue mineral content, collagen type, and cross linking) and structure (bone size, cortical thickness and porosity, trabecular number, thickness, connectivity), and other factors (micro-damage burden, osteocyte density) to sex and racial difference bone fragility remain poorly defined. We do not know why and how bones break. The challenge is to measure these specific material and structural determinants of bone strength. Whether a combination of these material and structural properties will more accurately identify women likely to sustain fractures, or improve approaches to drug therapy, is unknown, but it is likely.

13.

14.

15.

16.

17.

18. 19.

20.

21.

References 1. Seeman, E. (1997). From density to structure: growing up and growing old on the surfaces of bone. J. Bone Min. Res. 12, 1–13. 2. Currey, J. D. (2002). Bones. In “Structure and Mechanics”. Princeton, UP, New Jersey, pp. 1– 380. 3. Zebaze, R. M., Jones, A., Welsh, F., Knackstedt, M., and Seeman, E. (2005). Femoral neck shape and the spatial distribution of its mineral

22.

23. 24.

mass varies with its size: Clinical and biomechanical implications. Bone 37(2), 243–252. Ruff, C. B., and Hayes, W. C. (1998). Sex differences in age-related remodeling of the femur and tibia. J. Orthop. Res. 6, 886–896. Duan, Y., Wang, X. F., Evans, A., and Seeman, E. (2005). Structural and biomechanical basis of racial and sex differences in vertebral fragility in Chinese and Caucasians. Bone 36, 987–998. Wang, X. F., Duan, Y., Beck, T., and Seeman, E. R. (2005). Varying contributions of growth and aging to racial and sex differences in femoral neck structure and strength in old age. Bone 36, 978–986. Parfitt, A. M., Travers, R., Rauch, F., and Glorieux, F. H. (2000). Structural and cellular changes during bone growth in healthy children. Bone 27, 487–494. Gilsanz, V., Roe, T. F., Stefano, M., Costen, G. and Goodman, W. G. (1991). Changes in vertebral bone density in black girls and white girls during childhood and puberty. New Engl. J. Med. 325, 1597–1600. Gilsanz, V., Gibbens, D. T., Roe, T. F., Carlson, M., and Senac, M. O. (1988). Vertebral bone density in children: effect of puberty. Radiology 166, 847–850. Murray, P. D. F., and Huxley, J. S. (1925). Self-differentiation in the grafted limb bud of the chick. J. Anatomy 59, 379–384. Seeman, E. (2002). An exercise in geometry. J. Bone Min. Res. 17, 373–380. Bonadio, J., Jepsen, K. J., Mansoura, M. K., Jaenisch, R., Kuhn, J. L., and Goldstein, S. A. (1993). A murine skeletal adaptation that significantly increases cortical bone mechanical properties. Implications for human skeletal fragility. J. Clin. Invest. 92, 1697–1705. Kozloff, K. M., Carden, A., Bergwitz, C., Forlino, A., Uveges, T. E., Morris, M. D., Marini, J. C., and Goldstein, S. A. Brittle IV (2004) mouse model for osteogenesis imperfect IV demonstrates postpubertal adaptations to improve whole bone strength. J. Bone Min. Res. 19, 614–622. McBride, D. J. Jr., Shapiro, J. R., and Dunn, M. G. (1998). Bone geometry and strength measurements in aging mice with the oim mutation. Calcif. Tissue Int. 62,172–176. Szulc, P., Seeman, E., and Delmas, P. D. (2000). Biochemical measurements of bone turnover in children and adolescents. Osteoporosis Int. 11, 281–294. Nishida, S., Endo, N., Yamagiwa, H., Tanizawa, T., and Takahashi, H. E. (1999). Number of osteoprogenitor cells in human bone marrow markedly decreases after skeletal maturation. J. Bone Miner. Metab. 17, 171–177. Stenderup, K., Justesen, J., Eriksen, E. F., Rattan, S. I., and Kassem, M. (2001). Number and proliferative capacity of osteogenic stem cells are maintained during aging and in patients with osteoporosis. J. Bone Miner. Res. 16, 1120–1129. Oreffo, R. O., Bord, S., and Triffitt, J. T. (1998). Skeletal progenitor cells and aging human populations. Clin. Sci. 94, 549–555. Lips, P., Courpron, P., and Meunier, P. J. (1978). Mean wall thickness of trabecular bone packets in the human iliac crest: changes with age. Calcif. Tissue Res. 10, 13–17. Vedi, S., Compston, J. E., Webb, A., and Tighe, J. R. (1984). Histomorphometric analysis of dynamic parameters of trabecular bone formation in the iliac crest of normal British subjects. Metabolic Bone Dis. Relat. Res. 5, 69–74. Gilsanz, V., Gibbens, D. T., Carlson, M., Boechat, I., Cann, C. E., and Schulz, E.S. (1987). Peak trabecular bone density: a comparison of adolescent and adult. Calcif. Tissue Int. 43, 260–262. Balena, R., Shih, M–S., and Parfitt, A. M. (1992). Bone resorption and formation on the periosteal envelope of the ilium: A histomorphometric study in healthy women. J. Bone Miner. Res. 7, 1475–1482. Seeman, E. (2003). Periosteal bone formation – a neglected determinant of bone strength. N. Eng. J. Med. 349, 320–323. Ahlborg, H. G., Johnell, O., Turner, C. H., Rannevik, G., and Karlsson, M. K. (2003). Bone loss and bone size after the menopause. N. Engl. J. Med. 349, 327–334.

220 25. Brown, J. P., Delmas, P. D., Arlot, M., and Meunier, P. J. (1987). Active bone turnover of the cortico-endosteal envelope in postmenopausal osteoporosis. J. Clin. Endocrinol. Metab. 64, 954–959. 26. Foldes, J., Parfitt, A. M., Shih, M–S., Rao, D. S., and Kleerekoper, M. (1991). Structural and geometric changes in iliac bone: relationship to normal aging and osteoporosis. J. Bone Miner. Res. 6, 759–766. 27. Aaron, J. E., Makins, N. B., and Sagreiy, K. (1987). The microanatomy of trabecular bone loss in normal aging men and women. Clin. Orth. RR. 215, 260–271. 28. Van der Linden, J.C., Homminga, J., Verhaar, J. A. N., and Weinans, H. (2001). Mechanical consequences of bone loss in cancellous bone. J. Bone Miner. Res. 16, 457–465.

EGO SEEMAN 29. Manolagas, S. C. (2000). Birth and death of bone cells: basic regulatory mechanisms and implications for the pathogenesis and treatment of osteoporosis. Endocr. Rev. 21, 115–137. 30. Duan, Y., Turner, C. H., Kim, B. T., and Seeman, E. (2001). Sexual dimorphism in vertebral fragility is more the results of gender differences in bone gain than bone loss. J. Bone Miner. Res. 16, 2267–2275. 31. Duan, Y., Beck, T. J., Wang, X–F., and Seeman, E. (2003). Structural and biomechanical basis of sexual dimorphism in femoral neck fragility has its origins in growth and aging. J. Bone Miner. Res. 18, 1766–1774.

Chapter 14

The Cells of Bone Jane B. Lian and Gary S. Stein

University of Massachusetts Medical School, Department of Cell Biology, 55 Lake Avenue North, Worcester, MA 01655, USA

V. The Osteoclast: A Functionally Unique Cell for Physiologically Regulated Resorption of Bone Mineral VI. Perspectives References

I. Abstract II. Introduction III. Developmental Signals for Cartilage and Bone Tissue Formation IV. Osteogenic Lineage Cells

I. ABSTRACT

that accommodate requirements for its central role in mineral homeostasis. The structural and metabolic properties of bone tissue arise from the organization and interactions of functionally distinct cell populations. The principal cells which develop the various bone structures and mediate the specialized functions of the mammalian skeleton are: mesenchymal stem cells recruited for the development and expansion of skeletal tissues, chondrocyte lineage cells which form the template for endochondral bone development and long bone growth, osteoblasts which synthesize the bone matrix, osteocytes throughout the mineralized bone matrix which support bone structure, the protective bone surface lining cells, and the multinucleated osteoclasts which are responsible for the resorption of calcified tissue. Throughout life, bone tissue undergoes remodeling; a continual process of resorption and renewal. Fidelity of bone formation and remodeling necessitates exchange of regulatory signals among these cell populations and their progenitors. The basis for our current understanding of bone development and homeostasis has been and continues to be principally derived from the properties of skeletal cells, characterization of expressed genes and identification of their functional roles through genetic analyses (mouse models and human diseases). Two distinct progenitor stem cell lineages contribute to formation of the skeleton: mesenchymal stem cells for

Bone formation requires an exquisite interplay among distinct populations of cells and tissue compartments to support the metabolic and structural functions of the skeleton. In this chapter, the properties of pluripotential skeletal progenitor cells, chondrocyte, osteoblast, and osteoclast cell lineage populations are described. Emphasis is placed on recent advances in understanding the cellular and molecular mechanism regulating the progression of stem cells to differentiated phenotypes during endochondral and intramembranous bone formation and the responsiveness of cells to physiologic regulation and mechanical forces. The dynamic integration of signaling pathways and novel transcriptional regulators of cell growth and differentiation that are coordinated for development of the skeleton and the control of bone turnover has led to new dimensions for improving the diagnosis and treatment of skeletal diseases.

II. INTRODUCTION The skeleton functions both as a connective tissue in response to mechanical forces to adapt its structural features and as an organ in response to physiologic signals Dynamics of Bone and Cartilage Metabolism

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chondrogenic and osteogenic cells and the hematopoietic stem cells which give rise to osteoclasts. It is becoming increasingly evident that both chondrocyte, osteoblast, and osteoclast differentiation require a multistep series of events modulated by an integrated cascade of gene expression that initially supports proliferation and the sequential expression of genes associated with each component of bone formation and resorption. Equally significant are the growth factor, mechanical stimuli, and hormone-responsive regulatory signals, which mediate developmental competency for expression of genes associated with skeletal cell proliferation, differentiation, and repair. In this chapter, the current concepts in understanding molecular and cellular mechanisms regulating the progression of osteoblast and osteoclast differentiation and the functional activities of distinct cell populations will be explored. Within this context, a basis can be provided for improved diagnosis of skeletal disease and treatment that is targeted to specific cells in bone tissue.

III. DEVELOPMENTAL SIGNALS FOR CARTILAGE AND BONE TISSUE FORMATION The complexity of the multiple signaling pathways that converge to form the skeleton is first appreciated by the formation of skeletal elements through two distinct developmental pathways. Intramembranous bone formation results from initial condensation of mesenchyme with direct differentiation of the cells into osteoblasts and gives rise to flat bones, for example, that comprise the cranium. Growth of this tissue continues from progenitor cells within the periosteum covering the bone surface or the cranial sutures. The process of endochondral bone formation (EBF) for the development and growth of long bones and vertebrae involves initial formation of a cartilage tissue from condensing mesenchyme. Perichondrium develops around a cartilage anlagen and provides a future source of progenitor cells for differentiation to chondrocytes. In the core of the anlagen, chondrocytes become hypertrophic producing a calcified cartilage matrix which subsequently is resorbed and replaced by bone tissue. Vascular invasion in hypertrophic cartilage leads to formation of the growth plates with clearly defined zones of proliferating and maturing chondrocytes. The growth plates spatially define growing ends of the bone and the marrow environment which houses hematopoietic tissue for blood cell differentiation and stromal cell progenitors for bone growth. Osteogenic cells are recruited to the cores of calcified cartilage for formation of trabecular bone spicules, concomitant

with the resorption of calcified cartilage. With induction of ossification, a periosteum develops over cortical bone to provide a continual source of progenitors for appositional growth to complement the endochondral long bone growth.

A. Inductive Events for Skeletal Development: Fibroblast Growth Factors, Bone Morphogenetic Proteins, and Wnt Signaling Different regions of the skeleton arise from three distinct embryonic zones, influenced by diffusible factors which induce the necessary cellular phenotypes for tissue formation [1, 2]. The major signaling proteins involved in segmentation of the vertebrate body are Hox genes which define the positions where bone structures will develop [3, 4]. Migration of neural crest cells from the ectoderm to the cephalic mesoderm provides progenitor cells for development of cranial and facial skeletal structures. Limb formation is initiated by proliferation of mesenchymal cells in the lateral plate mesodermal layer to form the appendicular skeleton. Cells of the paraxial mesoderm undergo condensation and segmentation to form somites under the regulation of the cell surface receptor Notch1 and its ligands, delta and serrate [5]. Sclerotome cells from somites are then induced to form cartilage by the cytokine sonic hedgehog (Shh) secreted by the notochord, producing the axial skeleton (spine, sternum, and ribs) [6, 7]. Importantly, Shh was shown to regulate mesenchymal cell recruitment into the osteogenic lineage in vitro [8]. Considering these different embryonic developmental programs of the mesoderm to form either flat bones directly (e.g., calvarium) or cartilage anlagen that will become bone (e.g., limbs and vertebrae), it should be recognized that a distinct osteoprogenitor may divert from a stem cell at these skeletal sites. The formation of skeletal tissues from the condensations of the mesenchymal progenitor cells at a specific site is determined by epithelial–mesenchymal interactions that control shape and size of the limbs through secretion of regulatory factors and target transcription factors. A group of epithelial cells, the apical ectodermal ridge (AER), caps the limb buds and secretes growth factors that pattern the limb. The clustered Dlx gene family encoding homeodomain proteins, for example, is expressed in the AER of the limb bud regulating the proximal–distal pattern of outgrowth as a regulatory network. These genes also specify the spatial and temporal formation of the craniofacial skeleton [9–11]. It is now being recognized that the regulatory factors which induce and control development of skeletal structures remain as key signals of bone renewal and repair. These include members of the TGFβ/BMP superfamily that induce

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mesenchyme condensation for bone formation, Wnt proteins which contribute to the formation of the bone axes and fibroblast growth factors (FGFs) that are essential for the earlier stages of limb bud outgrowth [12–17]. 1. Fibroblast Growth Factor Receptors and Ligands

The FGF receptors, FGFR1, FGFR2, and FGFR3 and the FGF ligands are expressed throughout skeletal cell populations and contribute to the regulation of progenitor and differentiated populations of cells during both intramembranous and endochondral bone formation [18–21]. FGFR1 is expressed in limb mesenchyme and in osteoprogenitor cells at the osteogenic front separating the nonosseous suture tissue between the ossification plates of the calvarial bone tissue. An FGFR1 activating mutation leads to premature fusion of craniofacial structure (craniosynostoses), while dominant-negative forms of FGFR1 will inhibit calvarial suture fusion [22, 23]. Mutations in FGFR2 are responsible for Apert syndrome, severe craniosynostoses [22], and Pfeiffer and Crouzon disorders [24]. FGFR2 is expressed as two variants, FGFR2b is required for limb outgrowth, while FGFR2c is required for osteoblast maturation [25]. Interestingly, different point mutations result in distinct phenotypic alterations in gene expression of the osteoblasts conveying the mutation [26, 27]. FGFR3 expression is initiated as chondrocytes differentiate long bone growth and functions as an important negative regulator of endochondral bone formation. Various knockin mutations of this gene lead to severe dwarfism [28]. Together with FGFR3, PTHrP signals coordinate cartilage and bone formation [29]. Complete ablation of FGFR3 leads to embryonic skeletal overgrowth [30], but also an osteopenia phenotype in the post-natal mouse [31]. Specific FGF ligands control limb outgrowth [14, 21, 32, 33]. Beginning in the lateral plate mesoderm, FGF10 in the presumptive limb regions induces FGF4 and FGF8 in ectodermal layers to become the AER. FGF2, FGF4, FGF8, and FGF9 all contribute to increasing proliferation of mesenchymal cells, but FGF2 and FGF18 appear to be important negative regulators of chondrocyte proliferation. Mice lacking FGF18 have defects in both chondrogenesis and osteogenesis [34, 35]. FGF2 was initially isolated from the cartilage matrix [36] and later identified in periosteal cells and osteoblasts [37]. FGF2 supports osteoblast growth and differentiation [38] and contributes to osteoblast survival [39, 40]. Mice lacking FGF2 have decreased bone mass [41]. These examples illustrate how FGF signaling through multiple receptors and ligands and with specific activities on different cell populations control expansion of embryonic cartilaginous tissue for endochondral bone formation (illustrated in Figure 1).

Figure 1 Regulatory factors controlling chondrogenesis and endochondral bone formation. The condensation of mesenchymal cells committed to chondrocyte differentiation is characterized by expression of the transcription factors Sox5 and Sox6. PTHrP expressed in the perichondrium stimulates chondrocyte proliferation. The increase in growth rate delays maturation of the hypertrophic chondrocytes which secrete IHH, thereby negatively regulating IHH. IHH, which diffuses into the epiphysis, stimulates PTHrP activity, but also increases bone collar formation in conjunction with BMPs. In this manner, growth plate events are coordinated with bone formation during development. Runx2 which is expressed in MSCs, must be down-regulated, mediated by Nkx3.2 for recruitment of progenitors into the chondrogenic lineage. Runx2 is then up-regulated later during endochondral bone formation in hypertrophic chondrocytes. Runx2 transcriptional activity is tightly regulated by co-regulatory protein interactions, such as the negative regulator histone deacetylase-4 (HDAC4). This interaction may be required to attenuate Runx2 activity and expression of its target genes (VEGF and MMP9) during FGF. RZ, resting zone; PZ, proliferating zone; HZ, hypertophic zone.

2. The BMP Family of Proteins and Skeletal Development

Bone morphogenetic proteins (BMP) are multifunctional growth and differentiation factors (GDFs) that support the development of many tissues including cartilage and bone. BMP signaling is required at an early stage of skeletal development for formation of the AER and dorsal–ventral patterning of the limb [42]. Their specific roles in the skeleton and regulation of BMP activity through antagonists have been recently reviewed [13, 43]. Briefly, BMPs are members of the TGFβ superfamily which function through specific receptors that are distinct but have overlapping functions [16, 44]. Genetic studies that have modified BMP receptors 1A and 1B have identified distinct activities. For example, constitutively active forms of BMP1A and BMP1B promote chondrogenesis and osteogenesis, but only the dominant-negative form of BMPR1B inhibits these events [45]. Complete null mutation of the BMPR1B Type I receptor revealed defects mainly in the

224 appendicular skeleton, with marked reduction in proliferation of the prechondrocytic population and subsequent chondrocyte differentiation [46]. The receptor signal is transduced by phosphorylating intracellular BMP- and TGFβ-specific receptor Smad proteins (R-Smads), which results in dissociation from the receptor, followed by formation of heterodimers with a DNA-binding Smad4 (a common-mediator Smad) that enters the nucleus and is competent to either target gene promoters or form multimeric complexes on gene promoters with other transcription factors. Antagonistic or inhibitory Smads (I-Smads) contribute to regulating this pathway (reviewed in reference [47]). BMP ligands are expressed in mesenchymal condensations functioning in chondrogenesis and later are expressed in specific cartilaginous regions of the limb (reviewed in reference [46]). BMP2, 4, and 7 are expressed in the perichondrium, while BMP6 is found in the prehypertrophic and hypertrophic zones of the growth plate. Characterizing specific BMP activities has been challenging due to early embryonic lethality as shown for BMP2 and BMP4. However, conditional knockouts are being characterized. BMP7 (also designated OP-1, osteogenic protein-1) is essential for appendicular skeletal development and is a strong anabolic factor for cartilage [46, 48]. GDF-5 is essential for normal joint formation and development of the appendicular skeleton [49]. In the postnatal skeleton, several BMPs continue to function in stimulating chondrocyte proliferation (e.g., BMP7), but will inhibit terminal differentiation and require regulation by BMP antagonists [50]. BMP activity is regulated by several antagonists, one being Noggin which is highly expressed in cartilage tissue. Disruption of this control results in early skeletal malformations in the mouse [51, 52]. BMP2 and BMP6 can stimulate cartilage differentiation to the hypertrophic phenotype [53]. BMP2 and BMP4 are potent osteoinductive growth factors, a property mediated by the induction of the Dlx and Runx transcription factors as immediate early targets of BMP2 [54–57]. A significant finding relevant to the osteogenic effects of BMPs is the interaction of Smad complexes with Runx2 (Cbfa1/AML3), a transcription factor essential for bone formation [58–60]. Recent studies indicate an important positive regulatory loop between BMP2 and the Runx2 transcription factor. Runx2 is induced within a few hours of BMP2 treatment [54]. BMP2 and Runx2 together then have synergistic effects in promoting osteogenesis [61, 62]. Runx2 is expressed in early embryogenesis [63, 64], followed by an up-regulation of Runx2 in late stages of bone development [65, 66], suggesting that this factor may be important in early specification of the phenotype, as well as having a significant role for sustaining osteoblast differentiation

JANE B. LIAN AND GARY S. STEIN

(see Section IV). How crosstalk between the BMP and Runx2 signals is regulated remains a question, but such regulation provides an example of “feed forward” loops necessary to support development of the skeleton. BMPs induce a spectrum of other transcription factors essential for differentiation of skeletal tissues, including Sox genes which are required for chondrogenesis [67, 68], homeodomain transcription factors which pattern the skeleton and promote bone formation [54, 69, 70], and Osterix, which, like Runx2, is essential for bone formation [71]. Thus, the chondrogenic and the osteogenic activity of BMPs are related to induction of specialized transcriptional regulators of cell differentiation. 3. Wnt Signaling and Skeletal Development

The Wnt family of 19 secreted cysteine-rich glycoproteins regulate numerous developmental processes including cell polarity, cell differentiation, and migration [72]. The Wnt signaling pathway is a recently appreciated key regulator of early skeletal development. Wnt proteins bind to and activate receptor complexes consisting of the Frizzled family of G protein-coupled receptors and the low-density lipoprotein (LDL) receptor-related proteins. Wnt signaling is transduced through three pathways [72]. Activation of the canonical pathway results in stabilization of β-catenin by inhibiting its phosphorylation involving casein kinase 1 and glycogen synthase kinase 3 within a protein complex, which prevents the targeting of β-catenin for ubiquitination/ proteasome degradation. Subsequently, β-catenin is translocated into the nucleus to form heterodimers with the TCF1 or LEF transcription factors for expression of Wntresponsive genes. In the absence of nuclear β-catenin, TCF/LEF is associated with transcriptional corepressors and suppresses Wnt target genes. β-catenin is thus a critical regulatory step of canonical Wnt signaling and for embryogenesis. The other two pathways, the planar cell polarity and the calcium pathways are not as well defined in the mammalian tissues in the skeleton. The Wnt signaling pathway is regulated by several antagonists. DKK (Dickkopf) interacts with the frizzled receptor complex preventing transduction of the Wnt signal, while a class of secreted frizzled related proteins (SFRPs) interacts with Wnt proteins sequestering them from interaction with frizzled receptors, as does WIF (Wnt inhibitory factor-1) and Cerberus [73]. While specific Wnt proteins that activate either the canonical β-catenin pathway or noncanonical pathways have been identified, additional studies are necessary to clarify the specific roles of Wnt factors that may operate through multiple pathways. Both gain- and loss-of-function mutations in specific Wnt proteins, as well as forced expression of Wnt factors in limb bud cultures, have revealed the significance

Chapter 14 The Cells of Bone

of Wnt signaling in regulating skeletal development [17, 72, 74]. Wnt10a misexpression in the developing chick limb identified its importance for AER formation. Wnt3a knockout mice have a skeletal phenotype as Wnt3a is required for somite formation [75]. Recently, a rare human genetic disorder tetra-amaelia, characterized by absence of all the limbs has been linked to mutation in Wnt3 [76]. The expression of several Wnts (Wnt4, 5a, 5b, 6, 11 and 14) during limb development in the chick suggests key functions in initial stages of skeletal development [77, 78]. Misexpression of Wnt14 identified its role in induction of joint interzone [79]. Wnt5a and Wnt5b coordinate the pace and transitions between chondrocyte zones [80–82]. Wnt5b, which is expressed in the prehypertrophic chondrocyte zone, as well as in joints and perichondrium, delays hypertrophy, while Wnt4 blocks initiation of chondrogenesis, but accelerates hypertrophy [83–85]. Therefore, the developing limb and endochondral bone formation which both rely on the regulation of the proliferation, maturation, and spatial organization of chondrocytes, are processes highly dependent on Wnt signals. The importance of canonical Wnt signaling in skeletal development and postnatal bone formation is now established. By conditional ablation in mice, β-catenin has been identified as a key regulator of formation of the apical ectodermal ridge (AER) and of the dorsal–ventral axis of the limb [86]. In a series of genetic studies, inactivation of β-catenin in mesenchymal lineage cells results in severe loss of bone from inhibited osteoblast maturation and increases osteoclast differentiation [87–89]. By increasing Wnt signaling, e.g., by ectopic or by expressing a stabilized form of β-catenin, produces enhanced ossification and suppression of chondrogenesis [88–90]. More direct effects on formation and turnover of the mature skeleton have been revealed by an activating mutation (gain-of-function) in the Wnt co-receptor LRP5 (Low-density lipoprotein Related Protein 5), resulting in the high bone mass trait in humans [91, 92], a phenotype reproduced in the mouse model [93, 94]. Consistent with this phenotype, the LRP5 loss-of-function mutation leads to an osteopenia accompanied by fractures in humans causing osteoporosis pseudoglioma syndrome (OPPG) [95, 96]. This low bone mass phenotype is recapitulated in the mouse [97]. Inactivation of the Wnt antagonist sFRP1 in mouse, enabling increased Wnt signaling, also resulted in a high bone mass in the mouse adult skeleton with a delay in bone loss in ages 7–9 months [98]. These findings underscore the significance of Wnt signaling in maintaining bone mass in the adult skeleton. Expression studies suggest direct roles of several Wnts in bone formation and maintaining bone mass. Expression of Wnt10b in mouse marrow results in increased metaphyseal bone volume, and bone that is

225 mechanically stronger. In addition, the female mice are protected from bone loss due to estrogen deficiency [99]. Consistent with this observation, the null mutation of Wnt10b revealed decreased trabecular bone in the Wnt10b−/− mouse. Wnt10b is expressed in pre-adipocytes and the mechanism of increased osteogenesis by Wnt10b relates to the inhibition in development of fat tissue in vitro and in vivo in the transgenic mice, which had 50% less white fat and virtually no brown fat that generates body heat [100]. These studies suggest that Wnt signaling is not only necessary for bone development, but supports maintenance of bone tissue functions in the adult skeleton through several mechanisms. We have discussed above three key signaling centers for development of the skeleton. It is important to recognize that the molecular mechanisms underlying the signaling pathways induced by secreted factors for skeletal development point to specific transcriptional regulators that control pattern formation and/or guide the mesenchymal cell to the chondrogenic versus the osteoblast lineage (selected key transcription factors are shown in Table I). Furthermore, the importance of coordinated activities among these pathways is being realized and will be the focus of new investigations. Interactions between FGF, BMP, and Wnt signals occur primarily at the level of regulating secreted factors for coordinating the timing of developmental events.

B. Regulation of Endochondral Bone Formation The coordination of chondrogenic differentiation with bone formation events is tightly regulated through multiple signaling pathways as discussed above. Both selective expression of secreted factors in chondrogenic subpopulations and feedback loops control the proliferation and maturation of chondrocytes and the pace of endochondral ossification [101]. Two essential regulatory proteins include Indian hedgehog (Ihh), a mammalian homolog of the Drosphila hedgehog secreted factor and parathyroid hormone-related peptide (PTHrP) which is secreted by many tissues, but with specialized autocrine and paracrine activities for regulating endochondral bone formation. PTHrP is abundantly expressed in the periarticular perichondrium and diffuses towards the prehypertrophic zones. PTHrP functions through the PTH(PTHrP) G protein-coupled receptor which is expressed at high levels in pre- and hypertrophic chondrocyte zones. These events are illustrated in Figure 1. Human mutations in this receptor characterize several chondrodysplasias. In mice deficient in PTHrP, chondrocyte maturation is accelerated while overexpression of PTHrP

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Transcription factor Pax1, Pax 2 Nkx3.2

T-box genes (Tbx4, Tbx5) Sox5, Sox6 HIF-1α Sox9

Transcriptional Regulators of Early Skeletal Development Role

• • • • • • • • • •

Reference

Normal digit formation Sclerotome formation Chondrogenesis Axial skeletal development Limb bud outgrowth Growth plate organization Notocord ECM sheath formation and nucleus pulposus development Required for epiphyseal cartilage ECM synthesis Chondrogenic lineage determination in the cranial neural crest Essential for cartilage function

Foxc1 Homeodomain Proteins (Msx1, Msx2; Dlx1–6) Gli factors

• Progression of intramembranous bone formation • Patterning and cell differentiation

Runx2

• • • •

TCF/LEF-1

• Mediator of Shh for sclerotome development • Target of Ihh and PTHrP signal • Required for normal endochondral bone formation Present in pre-mesenchymal condensations, but function unknown Expressed in chondrocytes at the growth plate Mediator of canonical Wnt signaling Essential for differentiation of hypertrophic chondrocytes and osteoblast

stimulates chondrocyte proliferation, thereby inhibiting cartilage differentiation and delaying bone formation [102–104]. These studies identify PTHrP as a negative regulator of the hypertrophic chondrocyte phenotype (i.e., terminal differentiation). Ihh secretion in the hypertrophic zones serves multiple functions in coordinating the events of expansion of the proliferative zone and regulating osteoprogenitor differentiation for maturation of the bone collar [105]. Ihh increases PTHrP expression and consequently chondrocyte proliferation. Ihh therefore maintains the chondrocyte phenotype in the resting and proliferative zones [101, 104, 106, 107]. PTHrP at the growth plate by inhibiting the differentiation of proliferating chondrocytes to hypertrophic chondrocytes inhibits Ihh synthesis [108]. This negative loop maintains a steady rate of chondrocyte maturation and ensures a synchronous timing of long bone growth. In addition to PTHrP, prostaglandin, thyroid hormone, retinoic acid, and growth hormone influence chondrocyte proliferation and maturation to the hypertrophic cell. Reserve zone and proliferating chondrocytes appear to be the target of T3 in vivo. T3 inhibits clonal expression of chondrocytes in vitro, but induces markers of hypertrophic resting chondrocytes [109, 110]. The prostaglandin, PGE2, also inhibits chondrocyte differentiation [111]. Retinoic acid (RA) is required for normal development [112, 113] and, interestingly, for repeated regeneration of deer antler tissue [114]. However, nonphysiologic levels of RA are detrimental to endochondral bone formation as a result of

Lu et al., 1999 [492] Herbrand et al., 2002 [493] Murtaugh et al., 2001 [121] Tribioli & Lufkin, 1999 [117] Takeuchi et al., 2003 [494] Smits et al., 2004 [495] Smits & Lefebvre, 2003 [496] Pfander et al., 2003 [497] Mori-Akiyama et al., 2003 [68] Akiyama et al., 2002 [124] Bi et al., 1999 [498] Rice et al., 2003 [499] Bendall & Abate-Shen, 2000 [500] Depew et al., 2002 [9] Miao et al., 2004 [501] Buttitta et al., 2003 [502] Vortkamp et al., 1996 [503] Jemtland et al., 2003 [504] Smith et al., 2005 [340] Stricker et al., 2002 [130] Logan and Nusse, 2004 [72] Hsu et al., 1998 [505]

RA inhibition of chondrogenesis [115]. In contrast, growth hormone enhances chondrogenesis and osteogenesis, but oversecretion will result in tissue with inferior mechanical properties [116]. Thus, EBF requires not only developmental signals, but also hormones and cytokines that promote transcriptional control of target genes for regulation of growth plate events. Skeletal development and progression of endochondral bone growth at the hypertrophic stage of cartilage maturation producing a calcified matrix involve coupled positive and negative transcriptional control of gene expression (Table I). This is illustrated by two transcription factors expressed in undifferentiated mesenchymal cells, Nkx3.2, the gene associated with the bagpipe mutation in mouse [117] and Runx2/Cbfa1 [118]. Runx2, the transcription factor essential for osteoblast differentiation, is expressed in prechondrogenic mesenchymes [66, 119]. Nkx3.2 is a strong repressor transcription factor and one of the earliest mediators of chondrocyte commitment [120, 121]. Repression of Nkx3.2 is critical for development of the axial skeleton and position of the jaw joint [117, 122, 123]. Through a series of derepression events, Nkx3.2 allows for activation of Sox 9, a requirement for chondrocyte differentiation [124]. As the mesenchymal cells are recruited into the chondrogenic lineage by Nkx3.2, Runx2 becomes down-regulated through direct transcriptional control mediated by an Nkx3.2 response element in the Runx2 gene [125]. Thus Nkx3.2 initiates a cascade of events for both suppressing osteogenesis and activating chondrogenesis.

Chapter 14 The Cells of Bone

In situ hybridization studies in mouse models have led Helms and colleagues to propose that cell fates for cartilage and bone are specified by a hierarchy of Sox9 and Runx2 expression [126]. Runx2 is then re-expressed in the hypertrophic zone of the growth plate following the down-regulation of Nkx3.2. Runx2 activates vascular endothelial growth factor and matrix metalloproteinase 9, both essential factors for vascular invasion and recruitment of both resorbing cells to remove the cartilage matrix and osteoprogenitors [119, 127–129]. These findings suggest Runx2 may coordinate calcification with recruitment blood vessels and resorption/degradation of the calcified cartilage matrix. Forced expression of Runx2 in chondrocytes in transgenic mouse models shows that Runx2 supports hypertrophic chondrocyte differentiation [130–134], thus confirming the skeletal abnormalities of Runx2 null models as well as mutations in the Runx2 DNA-binding partner, CBFβ [63, 64, 66, 135, 136]. Evidence is accumulating for a key regulatory role of Runx2 to support chondrocyte maturation, perichondral invasion, and osteogenesis in cartilaginous tissue ([137] and reviewed in [138]). Synthesis of a cartilage extracellular matrix competent to calcify is a pivotal stage of long bone growth. Hypertrophic chondrocytes have unique properties defined by morphology, function, and by the expression of proteins with specific consequences restricted to this cell for production of a calcified matrix, such as type X collagen, small proteoglycans, and several noncollagenous proteins that are abundant in bone. The mineralized calcified cartilage matrix is requisite for resorption by chondroclasts/osteoclasts. Hypertrophic chondrocytes have a limited life span and are destined for apoptosis. These phenotypic changes in the chondrocytes are necessitated by the requirements for tissue organization, where the mineralized matrix at the growth plate eventually recruits the bone marrow stromal cells which will produce trabecular bone to accommodate lengthening of the bone during growth.

IV. OSTEOGENIC LINEAGE CELLS A. Mesenchymal Stem Cells: The Promise for Treating Skeletal Disorders 1. Stem Cell Populations and Properties

Advances in stem cell- and bone marrow-derived mesenchymal stem cell biology are increasing utilization of these cells for correcting genetic disorders and tissue repair.

227 Embryonic stem cells (ESC) are derived from the inner cell mass of blastula-stage embryos and express genes characteristic of the early blastocyst [139]. ESCs are defined by their infinite self-renewal capacity and will give rise to nearly every tissue and cell type in the body dependent on the appropriate in vivo environment and stimulus. Intact embryoid body tissue or micromass culture of ESCs will undergo chondrogenesis [140] and ESCs can be directed into osteogenic cells in the presence of ascorbate, 1,25(OH)2D3, β-glycerol phosphate [140–143]. Procedures have been detailed in the Methods in Enzymology series [144–146]. In the adult, stem celllike populations are being isolated from various tissues. Bone marrow contains the hematopoietic stem cell, a wellcharacterized self-renewing population that gives rise to every blood cell type. Within the bone marrow is a rare and rather elusive population of multipotential adult progenitor cells (MAPC) that can form endodermal, mesodermal, and ectodermally derived cell types; for example, endothelial cells, hepatocytes, and neurons in vitro [147, 148]. When these cells are injected into blastocytes, they contribute to tissues that include brain, long muscle, kidney, intestine, blood, skin, retina, spleen, and bone marrow in mice [149]. A side population (SP) of cells is characterized by their ability to extrude Hoechst dye and are enriched for hematopoietic stem cells ( HSC) and will differentiate into chondrocytes and osteoblasts [150]. Another population of stem-like cells of mesenchymal origin persists in the marrow. This is the mesenchymal stromal cell (MSC) which is an adherent cell with a limited lifespan (no more than 50 doublings) and has the ability to differentiate into multiple lineages, including adipocytes, chondrocytes, and osteoblasts. Osteogenic progenitor cells can also be derived from the periosteum. The stem cell-like properties of cells isolated from adult bone marrow and tissues are a question of recent debate as there is evidence for their plasticity (that is, the ability to acquire a mature phenotype by cell–cell fusion) or to transdifferentiate when exposed to a different microenvironment [151]. Progression of the most primitive pluripotent cell to the undifferentiated multipotential mesenchymal cell is not understood. Assurance of their stem cell activity must be defined by transplantation to a recipient animal examining their differentiation long term [152]. Using characterization of hematopoietic stem cells as a paradigm, several cell surface markers as well as antibodies generated to bone marrow stromal cell populations are used to identify presumptive stem cells from bone marrow. These reagents, including for example Stro-1, Sca−/+, SB10, have the potential for both recognition and purification of skeletal-related progenitor cells [153–158].

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Maintaining the stem cell-like properties in vitro has been challenging. Stem cells by their nature are generally in a noncycling (G0) stage of the cell cycle. ES cells can be propagated in culture on a feeder layer of mouse embryonic fibroblasts or without feeders in the presence of leukemia inhibitory factor (LIF), required for maintenance of mouse, but not human, ES cells [139]. Oct4 (a POU homeodomain protein) and nanog (a new homeodomain protein) are also requirements for a self-renewal of ES cells [159, 160]. Progenitor cells are responsive to a broad spectrum of regulatory signals that both maintain their pluripotent properties and mediate commitment to osteoblast differentiation and progression of phenotype development. When suspensions of marrow cells are plated in vitro, clonal colonies of adherent fibroblasts are formed; each derived from the single cell which has been designated as the colony-forming fibroblastic unit or CFU/F formation [161]. A proportion of these cells have high proliferative and differentiation capacity and exhibit characteristics of stem cells when transplanted in the closed environment of a diffusion chamber [162]. The CFUs are readily differentiated to osteogenic colonies in appropriate medium. The osteoprogenitor appears to have limited selfrenewal capacity and also, in contrast to the stem cell, has a capacity for extensive proliferation. Formation of CFUs requires the presence of hematopoietic cells [163] and recent studies have raised provocative implications of a direct influence of immune cells in contributing to osteogenic differentiation [164]. 2. From Stem Cell to Osteoprogenitor

Commitment of colony-forming units to the osteoprogenitor phenotype is regulated by several signaling proteins that include cytokines, growth factors, and hormones which influence growth and differentiation. Leukemia inhibitory factor, which maintains stem cell populations by inhibiting their differentiation, has also been reported to have osteogenic activity by enhancing differentiation of preosteoblasts [165, 166]. Platelet-derived growth factor and epidermal growth factor have been identified as important in stimulating expansion of the CFU/F [163, 167] and contribute to osteoprogenitor cell survival [168]. The fibroblast growth factor (bFGF) and transforming growth factor β1 (TGF-β1) are potent mitogens for periosteal osteoprogenitors and marrow stromal cells [169–171] and are expressed and produced by osteoblast lineage cells. These growth factors are stored in the bone extracellular matrix and thus provide a local mechanism for stimulating proliferation of progenitors in the bone microenvironment [172, 173]. The osteoinductive effects of the bone morphogenetic proteins (BMPs)

are complex and dependent on the specific BMP, concentration dependency, and the progenitor cell phenotype [43]. The activity of BMPs is regulated by production of inhibitors. Noggin, for example, is a glycoprotein that binds selectively to BMPs and inhibits stromal cell differentiation in vitro [174]. Tob proteins also inhibit BMP activity, but through a different mechanism, by enhancing the inhibitory Smad–receptor interactions to repress BMP signaling [175, 176]. BMP-2 rapidly induces osteoblast differentiation in marrow stromal cells [177], a number of pluripotent cell lines [178], and is competent to transdifferentiate the mouse myogenic cell line C2C12 into osteoblasts [179]. BMP2 is most widely used as an inducer of osteogenesis and is currently in clinical trials to augment fracture repair [180]. Among the hormones most influencing early osteogenesis are parathyroid hormone (PTH) which stimulates growth of osteoprogenitor populations [181], and glucocorticoids that have been shown to stimulate the growth and differentiation of CFUs and osteoprogenitors from human and rat, but not the mouse [182]. However, glucocorticoids have a marked inhibitory effect on bone formation by decreasing the number of mature osteoblasts and their activity through inhibition of insulin-like growth factor I [183, 184]. Increased bone formation by intermittent PTH is due to the stimulation of proliferation and differentiation of osteoprogenitor cells in bone marrow (reviewed in reference [185]). Recently it was shown that PTH strongly up-regulated amphiregulin, an epidermal growth factor family member [186]. Amphiregulin is a potent growth factor for preosteoblasts, although this growth factor inhibits osteoblast differentiation. In addition to growth-stimulating effects, PTH arrests cell cycle progression [187]; thereby, PTH promotes osteoblast maturation. Thus, PTH contributes to progenitor cell expansion in part through induction of the growth factors and further functions as an anabolic bone agent countering the inhibitory effects of growth factors on osteoblast differentiation. Parathyroid hormone-related peptide (PTHrP) that also binds to the PTH receptor, functions as a cellular cytokine regulating osteoblast growth for differentiation in development (reviewed in reference [188]). The significance of many of these growth factors and morphogens in regulating progression of the pluripotent stem cell and multipotential mesenchymal cell to the committed osteoprogenitor and finally to recognizable osteoblasts, is appreciated from mouse models, in which the PTHrP gene and the PTH/PTHrP receptor for these proteins have been ablated with severe consequences on formation of the skeleton [104, 189, 190]. Recent studies revealed PTHR-1R as a signature for definitive osteoprogenitors [191].

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Chapter 14 The Cells of Bone 3. Molecular Mechanisms Regulating Allocation to Osteoblastic Phenotype

Much attention has been given to lineage allocation of mesenchymal stem cells in marrow between the osteoblast and adipocyte bone. While debate still occurs as to whether the inherent osteogenic potential of the MSC osteoprogenitor number in marrow declines with aging, factors influencing lineage direction have been clearly defined. Notch, an important signaling receptor that controls cell fate determination during development, has opposing effects on mesenchymal cells, suppressing chondrogenic differentiation [192], but stimulating osteoblast differentiation [193]. Menin, a product of the multiple endocrine neoplasia type 1 (MEN1) gene, was recently identified as a requirement for MSC commitment to osteoblasts in the null mouse [194]. PTHrP interacts with the BMP2 by enhancing BMP1A receptor expression pathway for osteoblastogenesis, but decreases adipogenesis [195]. Commitment of a stem cell to a phenotype is regulated by cell shape and cytoskeleton changes which involve Rho GTPase activity. A dominant-negative RhoA promotes a round shape leading to adipocyte differentiation,

Figure 2

while a constitutively active RhoA induced the osteogenic phenotype independent of cell shape [196, 197]. Physical forces on the MSC appear to be a significant component for osteoblast allocation as microgravity inhibits osteoblast colony formation of human MSCs and increases adipocytes [198, 199]. Finally, transcriptional regulators of gene expression have potent and direct effects in modifying cellular phenotypes. A number of key studies have defined master genes that direct a pluripotent cell to different lineages. Adipogenesis is promoted through the activities of PPARγ and CEBPα [200, 201], chondrogenesis requires Sox9 [202] and osteoblast maturation requires Runx2 [65, 66] and Osterix [71, 203] (Fig. 2). Inhibitory transcription factors, such as GILZ or retinoic acid, can block adipogenesis [204], thereby increasing a pool of progenitors for osteoblast differentiation. The potency of Runx2 in directing osteogenic commitment is provided by numerous studies that show Runx2 expression can activate bone phenotypic genes in pluripotent cells and redirect a committed premuscle cell into the osteoblast lineage [57, 205] or inhibit the adipogenic phenotype [206]. Conversely, activation of PPARγ in osteoblasts will down-regulate

Regulation of osteoblastogenesis. (A) Selected stages of osteogenic lineage cells are illustrated indicating secreted factors (BMPs, TGFβ, FGFs, and Wnt proteins) and hormones most influencing progression of the osteocyte from the earliest stem cell. In the lower panel of (A), transcriptional control of lineage allocation of the mesenchymal stem cell to nonosseous phenotypes is shown. (B) Markers frequently used to characterize stages of maturation include collagen type I, alkaline phosphatase, bone sialoprotein, osteopontin, and osteocalcin. (C) Examples of transcription factors regulating progression of osteoblast differentiation are indicated by direction of the arrows.

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Runx2-mediated transcription of bone phenotypic genes [207]. Even more significant, PPARγ-deficient ES cells not only failed to become adipocytes, but spontaneously differentiated to osteoblasts [208]. The Runx2 transcription factor is expressed in the early pluripotent mesenchymal cell prior to expression of the osteoblast phenotype, and increases during osteoblast differentiation, consistent with its genetic role in maturation of the osteoblast phenotype [63, 65]. Runx2 protein is also retained during mitosis and may provide a mechanism for regulating genes that retain the osteogenic lineage properties of dividing cells [209]. The in vivo significance of Runx2 in early commitment to the osteoblast lineage is indicated by the evidence that mesenchymal progenitor cells from Runx2-null mice differentiate more towards chondrocytes and adipocytes [210]. Importantly, Runx2 may function as an inhibitor of proliferation of progenitors, thus providing a mechanism for regulating the transition from growth to a post-proliferative stage as a component of cellular commitment to osteogenic lineage cells [211]. We are now beginning to reach an understanding of the complexity of factors required to support expansion of a progenitor cell and the signals which must be initiated for stem cells to acquire an osteogenic property. With the onset of new discoveries, considerations for how the different regulatory proteins can be applied for a therapeutic strategy must take into account their effects on a spectrum of diverse activities within different pathways.

B. Osteoblasts: The Bone-forming Cells and Gatekeepers of Hormonal Activity The mature surface osteoblast is defined by the biosynthesis and organization of the bone extracellular matrix. The primary function of osteoblasts on active bone-forming surfaces is the production of a bone matrix, contributing to expansion of bone volume by laying down osteoid and secreting factors that facilitate mineral deposition. An equally important function of osteoblasts and the preosteoblasts which lie in close proximity to the bone surface is their responsiveness to endocrine factors and the production of paracrine and autocrine factors for the recruitment of osteoprogenitors, the growth of preosteoblasts, and the regulation of osteoclastic resorption of the mineralized bone matrix. Osteoblasts and osteocytes have receptors for Key regulators of bone turnover, which include cytokines, parathyroid hormone, 1,25(OH)2D3, and sex steroids. These features provide mechanisms for mediating the coupling of osteoblast and osteoclast activities (see Section V).

The in vitro study of primary osteoblasts from several species has allowed molecular analysis of expressed genes and transcription factors regulating the differentiation of osteoblasts in relation to progressive formation of the bone matrix [212]. These studies, carried out over a period of several weeks, provide a definition of discrete stages of maturation of the osteoblasts (Fig. 2). The proliferating preosteoblast stage is engaged in synthesis of growth factors and matrix components of woven bone. When the osteoprogenitor or preosteoblast ceases to proliferate, a key signaling event occurs for development of the large cuboidal surface osteoblast from the spindle-shaped osteoprogenitor. The osteoblast vectorially secretes type I collagen and specialized bone matrix proteins towards the mineralizing front of the tissue. The matrix maturation stage marks the postproliferative osteoblast expressing maximal levels of alkaline phosphatase to render the matrix competent for mineralization. Type I collagen continues to accumulate, together with the specialized classes of calcium-binding proteins, osteopontin, bone sialoprotein, and osteocalcin. The proteins are up-regulated at the mineralization stage with initial deposition of hydroxyapatite in the ECM. Both in vivo and in vitro osteoblasts at sites of bone formation and in the mineralized nodule undergo apoptosis, reflecting a biological mechanism for triaging those osteoblasts recruited to produce matrix but cannot all be accommodated in the mineralized matrix as osteocytes [213–217]. The increase in collagenase detected in cells at the mineralization stage in vitro may reflect the reorganization of the extracellular matrix necessary for maturation of osteoblasts to the osteocytes. Guiding the transitions from the osteoprogenitor to the osteoblast and osteocyte are the cell–matrix and cell–cell interactions, presumed to be important for progression of differentiation. Functional studies establishing requirement of the type I collagen and other ECM components in promoting osteoblast/osteocyte differentiation have been carried out by modifying production of osteoblast-secreted products or culturing osteoblasts on various matrices [218–221]. Indeed, the ECM composition changes during maturation of the osteoprogenitor cells. Osteoprogenitor cells synthesize collagen type III and large proteoglycans, versican and hyaluronan; whereas in preosteoblasts and osteoblasts, these proteins are replaced by collagen types I and V and small chrondroitin sulfate proteoglycans, as decorin and biglycan [222]. ECM-integrin signaling is an important pathway for activation of osteoblast gene expression. Osteoblasts appear to use β1 integrins to adhere to the full range of RGD-containing bone matrix proteins [223]. The intracellular kinase pathways that become activated through integrin signaling can modify a spectrum of factors

Chapter 14 The Cells of Bone

promoting osteoblast differentiation. For example, the Runx2 transcription factor essential for bone formation becomes phosphorylated in response to ECM-kinase signaling, enhancing its functional activity [224]. Cell–cell adhesion proteins also characterize osteoblast maturation. Several members of the cadherin family are expressed in osteoblasts including cadherin-11, cadherin-4, N-cadherin, and OB-cadherin [225]. N-cadherin is present on proliferative preosteoblastic cells, but is lost as they become osteocytic [226]. In contrast, OB-cadherin is barely detected in osteoprogenitor cells and is up-regulated in alkaline phosphatase-expressing cells [227]. The activated leukocyte cell-adhesion molecule (ALCAM; CD166) is highly expressed on the surface of MSCs and becomes down-regulated in concert with changes in morphology and detection of alkaline phosphatase activity as periosteal progenitors migrate and develop into an osteoblast [153, 228]. Signaling pathways from the extracellular matrix through the cytoskeleton and finally to the nucleus, which allow expression and up-regulation of bone-specific and bone-related genes, are being investigated. For example, β-catenin, the intracellular regulator of the canonical Wnt pathway, also colocalizes and coprecipitates with cadherins [225]. Another potential candidate is CD44, the hyaluronate receptor which is a nonintegrin adhesion receptor that is linked to the cytoskeleton. CD44 is enriched during osteocyte differentiation [229, 230] and has become a useful marker for this stage [231]. The temporal expression of proteins and enzymes involved in cell adherence and the initial production and modifications of the extracellular matrix, provides competency for mineral deposition and the final stage of osteocyte phenotype development in the mineralization period. The induced expression of subclasses of osteoblastic genes (shown in Fig. 2, osteocalcin, osteopontin, bone sialoprotein) are used as markers of the stages of osteoblast maturation, but may reflect selective functions of osteoblasts dependent on their location in bone. This concept is supported by the analysis of these expressed genes and protein levels at single-cell levels in osteoblast cultures developing a tissue organization (the bone nodule) and in bone sections [191, 232, 233]. The variations in levels of expressed gene products reveal subtle yet important differences in functional properties that relate to the establishment and maintenance of bone structure. Although there are likely to be more than the major stages in development of a mature osteoblast (depicted in Fig. 2), this model of differentiation provides a better understanding of cellular properties and selective responses to growth factors [234–236] and hormones [182, 237–240]. For example, TGFβ stimulates the replication of progenitor cells and directly stimulates collagen

231 synthesis, but the mitogenic effects of TGFβ are not apparent on mature postproliferative osteoblasts. The fibroblast growth factors (FGF-1, FGF-2) also stimulate bone cell replication in cells capable of collagen synthesis. However, FGF2 does not directly affect the differentiated functions, Indeed, in mature osteoblasts, FGF both inhibits type I collagen synthesis and increases collagenase expression [235]. The insulin-like growth factors (IGF-1 and IGF-2), which are synthesized by osteoblasts, are key regulators of osteoblast anabolic activities [241]. IGF-1 directly increases type I collagen synthesis and complementarily decreases the activity of the collagenase 3 (MMP-13) enzyme, reflecting its role in maintenance of the bone matrix. Anabolic effects of IGF-1 and IGF-2 in bone remodeling are further underscored by regulation of these proteins by hormones [242]). Their essential requirement for bone mineralization is established by the osteoblastspecific mutation of the IGF receptor [243]. Osteoblast differentiation and activity is strongly influenced by the steroid hormones glucocorticoid [182, 244], 1,25(OH)2D3 [237, 238, 245], estrogen [239], and androgens [246]. All have selective effects at specific stages of osteogenesis. Glucocorticoids and 1,25(OH)2D3 have antiproliferative effects to promote differentiation of the cells at early stages of maturation, but inhibit anabolic activities and promote resorptive properties of the osteoblast at later stages. Estrogens exert both antiapoptotic effects on bone-forming cells mediated by activation of the Src/Shc signal transduction pathway [247] and proapoptotic effects on mesenchymal cells and osteoclasts through classical ER regulation of target genes [248, 249]. The thyroid hormone receptor is expressed in all osteogenic lineage cells and affects bone formation and resorption [250]. Thyroid hormone (T3) is essential for linear growth and maintenance of bone mass (reviewed in reference [251]). Mice deficient in thyroid hormone receptors exhibit decreased bone mineral density and increased adipogenesis [252]. Differential regulation of specific gene expression as a function of differentiation by polypeptide hormones has also been documented. For example, PTH increases the resorption promoting RANKL cytokine to a greater extent in mature osteoblasts, but inhibits expression of the antiresorptive osteoprotegerin factor at all stages of osteoblast differentiation [253]. PTHrP has bidirectional effects on phosphorylation of ERK between mesenchymal cells and differentiated mouse osteoblast lines implicating MAPK signaling in regulating the selective effects of this hormone [254]. Taken together, our current understanding is one of selective and combinatorial responses of osteogenic cells to growth factors, hormones, and physical forces dependent on the structural and metabolic requirements of the cells in a particular tissue environment.

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C. The Osteocyte: Differentiation Supports Bone Structure, Viability and Physiologic Responsiveness The final stage of osteoblastogenesis begins with deposition of mineral in the ECM by several mechanisms (reviewed in reference [255, 256]). Osteoblasts on boneforming surfaces have several fates. As mineralization of the matrix envelopes the osteoblast, morphologic changes are induced in the cell when the osteoblast progresses to the fully differentiated osteocyte. Alternatively, on a quiescent bone surface, the osteoblast develops into a flattened bone-lining cell of a single layer forming the endosteum against the marrow and underlying the periosteum directly on the mineralized surfaces. This cell layer lining protects the bone from the extracellular fluid space. Lining cells function to (1) maintain the bone’s own environment through synthesis and resorptive-like activities, (2) protect themselves from cell death, and (3) respond to mechanical forces on the skeleton. Osteoblasts receive the majority of systemic and local signals and can transmit these to osteocytes, while the osteocytes perceive mechanical forces on the bone and transmit the regulatory information to the surface-lining cells. Osteocytes represent the most abundant cell in the skeleton. They are in direct communication with the bone-lining cell and with each other within the mineralized matrix through cellular processes that lie within and are tethered to the canaliculi channels [257]. Thus, the osteocytes and surface-lining cells form a continuum, or syncytium, by connection of their cell processes through gap junctions. The osteocyte phenotype develops during embedment of the osteoblast in mineralized bone, although the signaling mechanisms for how the cuboidal osteoblast transitions to the stellate osteocyte remain unknown. Osteoblasts that do not differentiate to the osteocyte or bone-lining cell undergo programmed cell death [216, 258]. However, TGFβ activation has been reported to maintain osteoblast survival during their differentiation into osteocytes [259]. Intermittent PTH (as opposed to continuous PTH treatment) also prevents osteoblast apoptosis increasing the lifespan of mature osteoblasts [260, 261]. The osteocyte is a terminally differentiated cell and when isolated directly from bone tissue, retains its morphologic features in vitro. In culture, the cell does not divide and upon attachment to surface, forms characteristic cytoplasmic extrusions in all directions [262]. The characterization of osteocytic cell lines has recently advanced our understanding of their phenotypic properties [263]. Osteocytes have the capacity to synthesize certain matrix molecules, as abundantly as osteoblasts, visualized by in situ studies. The detection of osteocalcin, for example, is particularly robust, supporting the relevance of in vitro

models for osteoblast differentiation which show maximal levels of osteocalcin in mineralized nodules. The dentin matrix protein 1 (DMP1) was found to be expressed 20-fold higher in the differentiated osteocyte and stimulated further by mechanical loading [264, 265]. The DMP1-null mice have reduced bone mineral [266]. Interestingly, a mouse model expressing 8 kb of the DMP1 promoter-GFP transgene showed responsiveness to increased mechanical loading in osteocytes [267]. Thus, highly specialized proteins are likely to be synthesized by osteocytes to support the integrity of mineralized bone tissue. Survival of the single osteocyte, resident in lacunae and in contact with other cells, is critical for bone to function as a viable organ and structural connective tissue. Disruption of cell–cell contact leads to cell death. In vivo osteocytes are long-lived cells, but, with aging, empty lacunae from cellular apoptosis have been observed [213, 214, 217, 258]. Several mechanisms protecting the osteocyte against apoptosis are being established. The CD40 antigen is expressed in an osteocyte cell line and adding CD40 ligand to cultured cells inhibited apoptosis induced by TNFα and glucocorticoids [268]. In vivo the loss of cell viability, as occurs in areas of microcracks, is coupled to targeting and initiation of new remodeling activity [214, 269–272]. However, the specific mediators of recruitment of cells to the microcracks need to be identified. Interestingly, a higher osteocyte lacunae density was quantified in woven compared to lamellar bone, further implicating the osteocyte with bone remodeling [273]. In vitro apoptosis of osteoblasts and osteocytes is associated with the mineralized nodule [216], a finding which may be analogous to in vivo bone formation where osteoblasts at bone-forming surfaces do not all mature to an osteocyte [215]. Alternatively, in vitro osteocytes may lack requisite factors for survival, as observed in vivo [217]. Bone as a structural connective tissue can adapt its architecture to meet physical demands on the skeleton. The osteocyte syncytium is highly responsive to loading forces [271, 274]. Mechanical forces on the bone result in stress-generated electric potentials produced by strain in the organic components (piezo-electric potential) or result in electrolyte fluid flow produced by deformation of the bone (streaming potential) ([268, 275] and reviewed in reference [276]). After osmotic pressure and vascularderived pressure gradients respond to the mechanical loading, the fluid movement which is produced through bone tissue is regulated by permeability of the tissue controlled by hyaluronan [277, 278]. Mechanisms which contribute to osteocyte function as the mechanosensor of skeletal tissues are being identified [279, 280]. Mechanical strain and fluid flow regulate their communication through GAP junctions [281]. An important component in establishing a viable bone tissue is an

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obligatory cell–cell contact mediated by GAP junctions and formed primarily by connexin 43, between osteoblast and osteocytes within a mineralized matrix [282]. Both delayed osteoblast differentiation and increased osteoblast apoptosis characterize the connexin 43-null mouse. Thus, GAP junctions and the connexin 43 hemichannels maintain cell survival [283]. Other antiapoptotic signals continued to be identified; for example, the CD40 ligand known to function as an antiapoptotic factor in dendritic cells. Fluid shear forces were reported to promote cell survival by increasing the antiapoptotic factor Bcl2 and decreasing the proapoptotic Bax protein, while disuse resulted in apoptosis [284]. Importantly, GAP junctions allow the osteocytes to be metabolically and electrically coupled to surface osteoblasts. Rapid fluxes of bone calcium across these junctions facilitate transmission of information between osteoblasts on the bone surface and osteocytes within the structure of bone itself [285]. At a molecular level, it is now realized that the GAP junctions respond to mechanical strain on the bone through PGE2 and the prostaglandin EP2 receptor [263, 281, 286] and by increasing connexin 43 expression [287]. Mechanical strain is also likely to influence cell– matrix interactions through integrin signaling. Integrins are tightly coupled to the cytoskeleton [288] and together the integrin–cytoskeleton complex facilitates the transduction of mechanical signals that may ultimately lead to modifications in gene expression. The extracellular matrix receptors, such as integrins and CD44 receptors, are thought to mediate cellular sensing of mechanical loads [289]. Mechanical strain induces changes in attachment of cellular processes to the canalicular wall [257] and focal adhesion kinase activity [290]. Therefore, several components of osteocyte cellular architecture will respond to mechanical strain. While physiologic mechanical strain can maintain bone mass, skeletal unloading has negative effects on bone, for example, skeletal unloading alleviates the anabolic action of intermittent PTH [291]. The molecular mediators of osteocyte mechanotransduction signals are being identified, increasing our understanding of osteocyte functions and responses. Modifications in signaling factors, enzymes, and transcriptional regulators have been reported [292–294]. Direct evidence that osteocytes sense mechanical loading [295] has been demonstrated by rapid changes in metabolic activity by either 3H-uridine uptake [296], increased glucose-6-phosphate dehydrogenase activity [297], activation of the kinase signal transduction pathway [290], and increased IGF-1 expression [298]. Osteocytes produce IGF-1 and release prostaglandins in response to stress [299]. Signals induced by fluid flow include prostaglandin PGE-2, cAMP, and nitric oxide (NO), all potent stimulators of bone formation and bone resorption consistent

with mechanical stimulation of bone turnover [263, 293, 300, 301]. Mechanical strain activates the endothelial nitric oxide synthase (eNOS) in osteocytes producing NO [302]. NO is a highly reactive molecule and NO produced by the eNOS isoform is essential for osteoblast function [303], while the inducible isoform (iNOS) produces cytokines that act on osteoclasts [303–306]. Recently, the neuronal NO synthase has been implicated in bone turnover [307]. Interestingly, osteocytes have the capacity to retain responses and the mechanisms contributing to their longterm potential are being identified. Skerry and colleagues considered the involvement of the NMDA and AMPA glutamate receptors, which account for the cell memory in the central nervous system in osteocyte activities [308]. Glutamate, an intercellular signaling agent, was found to be activated in response to mechanical loading, and the glutamate receptor 2 shown to be important for osteoblast survival [309]. In conclusion, the structural organization of the osteocyte in bone and its direct contact with active osteoblast or surface lining cells is consistent with the concept that bone cells respond to varying physiological signals through multiple pathways to communicate their responses.

D. Transcriptional Control of Osteoblast Differentiation The progression of osteoblast differentiation requires the sequential activation and suppression of genes that encode phenotypic proteins and regulatory factors. Signaling molecules (morphogens, growth factors) indirectly result in a cascade of gene expression through induction of transcription factors that directly engage in protein–DNA as well as protein–protein interactions [212]. Human mutations, mouse genetics, gene profiling, and differential display strategies in response to osteoinductive factors, for example, BMP2, have all contributed to an explosion in the identification of many transcriptional regulators and signaling proteins that control bone formation. Several regulatory factors have been shown to be necessary for normal skeletal development and bone formation based on genetic models [310]. Examples of recently identified factors that play key roles in regulating maturation of osteoblasts, as well as maintenance of bone architecture proven from genetic models, are presented in Table II. From these new findings, several important concepts have emerged for understanding direct effects on gene expression that integrate physiological responses. The requirements for gene expression during osteoblast differentiation are now being understood to involve (1) both negative and positive transcriptional regulatory cascades

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Table II.

Transcriptional Regulators of Osteoblast Differentiation

Promote osteoblast differentiation Runx2 JunB Fra1 䉭FOSB DLX3 DLX5

Osterix ATF4 C/EBP

CHOP C/EBP/LAP Inhibit osteoblast differentiation STAT1

C-fos

MSX2

TWIST

HEY1

Lian et al., 2004 [516] Hess et al., 2003 [517] Jochum et al., 2000 [518] Kveiborg et al., 2004 [519] Sabatakos et al., 2000 [520] Hassan et al., 2004 [326] Lee et al., 2003 [521] Miyama et al., 1999 [522] Tadic et al., 2002 [523] Tai et al., 2004 [524] Nakashima et al., 2002 [71] Yang et al., 2004 [525] Harrison et al. 2004 [363] Gutierrez et al., 2002 [361] Umayahara et al., 1999 [526] Pereira et al., 2004 [362] Iyer et al., 2004 [527] Kim et al., 2003 [506] Xiao et al., 2004 [507] Takayanagi et al., 2004 [508] Grigoriadis et al., 1993 [318] Sunters et al., 2004 [509] McCabe et al., 1996 [316] Ichida et al., 2004 [510] Dodig et al., 1999 [511] Yoshizawa et al., 2004 [512] Cheng et al., 2003 [513] Lee et al., 1999 [314] Bialek et al., 2004 [315] Ishii et al., 2003 [514] Zamurovic et al., 200 [515]

for supporting osteoblast phenotype development; (2) the role of co-regulatory protein interactions in response to physiologic factors for achieving gene-specific activation and suppression of transcription at appropriate times in development of a cell phenotype; and (3) combinatorial control mechanisms involving the integration of signal transduction with transcriptional control of gene expression, as well as interactions between transcription factors for cooperative effects on a gene promoter. This section will focus on selected transcription factors to illustrate these concepts. The Runx2 protein has provided a paradigm for understanding these regulatory mechanisms. 1. Regulatory Networks for Repression and Activation of Gene Transcription: AP-1 and Homeodomain Proteins

We have learned that factors with identified roles in specifying spatial differentiation and pattern formation during embryogenesis are re-expressed at selective stages of osteoblast maturation. As bone remodels in the postnatal

animal, there is a continuing requirement to re-establish and sustain a complex tissue organization. A concept is emerging that expression of many transcription factors expressed in the undifferentiated mesenchymal cell, although classified as inhibitors of osteoblast differentiation, are requisite for normal bone formation. The helix-loop-helix (HLH) family of proteins (Twist, Inhibitor of differentiation (Id), Scleraxis) are examples of negative transcriptional regulators at early stages of osteoblast development, but necessary for osteoprogenitor cell expansion [311]. They are expressed in mesoderm tissue of developing embryo, but not detected at the onset of ossification [312, 313]. The restricted expression of these factors in proliferating osteoblasts in vitro is consistent with their required downregulation in order for differentiation to proceed [314]. Another mechanism of negative regulation of osteoblastogenesis has recently been defined for twist inhibition of osteogenesis. Twist proteins transiently inhibit Runx2mediated transcription by interacting with the Runx2 DNAbinding domain. Thus, in vivo deficiency of Twist proteins results in premature osteoblast differentiation [315]. A typical molecular mechanism for control of gene expression involves a change in heterodimerization proteins to form either a transactivating or repression complex transcription of the gene at specific stages of osteoblast maturation. AP-1 factors include fos (c-fos, fra1, fra2) and jun (c-jun, jun-D, jun-B) oncogene-encoded transcription factors. These proteins regulate cell cycle and differentiation-related genes in cartilage and bone by forming homo- or heterodimeric complexes with fos, jun, and ATF member proteins at AP-1 regulatory elements. Through heterodimer formation with different partners, cellular levels of transcription can be finely modulated or selectively repressed or enhanced, as occurs on the osteocalcin gene where c-fos/c-jun down-regulates and fra2/ junD up-regulates transcription [316]. Another example revealing a novel function for an AP-1 factor is the binding of JunD to menin (product of the MEN1 tumor suppressor gene) which changes JunD from a growth suppressor to a growth-promoting factor when complexed with menin [317]. AP-1 elements are found in most genes and are responsive to growth factors, polypeptide hormones and other physiological signals. Fos- and jun-related proteins exhibit developmental stage-specific expression and activities during osteoblast differentiation in vivo and in vitro constituting a regulatory network [318–320]. Thus, in part, cellular levels of transcription factors are a determinant to which factors occupy regulatory elements. Genetic studies have established that several AP-1 family members are essential for normal bone development and osteoblast differentiation (Table I). In conclusion, the modifications observed in the representation and activities of family

Chapter 14 The Cells of Bone

members of these classes of transcription factors during osteoblast differentiation reflect linkage to transcriptional control for bone formation. The concept of a regulatory network of transcription factors for osteoblast differentiation is best illustrated by the Msx and Dlx homeodomain proteins. The importance of homeodomain proteins for normal bone formation is realized by skeletal abnormalities that result from mutations or misexpression of these factors [10, 321–324] (see also section III and Table I). The most well-studied during osteoblast differentiation are Msx2, Dlx3, and Dlx5 [56, 325, 326]. To ensure maintenance of the osteoblast phenotype, these related family members of specific classes of proteins are expressed in specific osteoblast subpopulations and temporally expressed during osteoblast differentiation with Msx2 restricted to proliferating and apoptotic cells, followed by up-regulation of Dlx3 and Dlx5 expression in postproliferative cells [216, 326–330]. The chromatin immunoprecipitation assay has allowed direct analysis of specific homeodomain (HD) proteins associated with gene promoters during osteoblast differentiation. Studies of the osteocalcin (OC) gene, which contains an HD element that supports osteoblast-specific expression of OC, have identified two molecular switches controlled by different HD proteins to regulate its transcription. Msx2 is bound to the OC HD element in the growth stage when no mRNA is detected [326, 331, 332]. When transcripts are expressed, Runx2 and Dlx3 replace Msx2. A second switch in HD protein occupancy at the OC promoter occurs at the mineralization stage with replacement of Dlx3 by Dlx5 binding to the gene [326]. In conclusion, these studies have provided evidence for coordinated control of gene expression throughout bone formation by family members of a class of transcription factors that form a regulatory network. 2. Co-regulatory Protein Interactions with Transcription Factors Support Integration of Physiologic Signals and Gene-specific Expression: Role of Runx Factors

A fundamental mechanism of transcription factor activity is modulation by co-regulatory protein interactions for integrating and transducing physiological signals. This concept is best illustrated by the Runx2 (CBFA1/AML) transcription factor introduced earlier in this chapter. It is one member of a three mammalian gene family that are designated runt homology domain (RHD)-related core-binding factors. The shared RHD binds to the DNA motif ATGGCT/G in target genes and requires a partner protein CBFβ which does not contact the DNA. Genetic studies in human and mouse have

235 shown each Runx gene and CBFβ to be essential for organogenesis and embryogenesis. In human these factors were initially designated as AML because the first gene to be identified (Runx1/AML1B) is encoded by gene loci that are rearranged in acute myelogenous leukemia (AML) [333]. Runx1/Cbfa2/AML-1B/PEBP2B is critical for definitive hematopoiesis. Runx1 is also highly expressed in perichondrium/periosteum, and downregulated in mature osteoblasts [334, 335], although a critical role in chondrogenesis has not been identified as Runx1-null mice die at 11.5 days postcoital. Runx3/Cbfa3 (AML2/PEBP2C) exhibits more restricted expression, contributing to gut and nerve development with Runx3 gene mutations associated with gastric cancers. However, Runx3 is detected in the hypertrophic cartilage zone, but its absence (Runx3−/− mouse) only slightly delays maturation and vascular invasion. At birth, a normal skeleton is found [336]. Runx2/Cbfa1/AML-3/PEBPP2A is specifically required for bone formation demonstrated by the inhibition of bone tissue formation in the Cbfa1-null mutation mouse model [63, 337]. Although the skeleton forms through the initial chondrogenic stage, mineralization is blocked. Recent studies have identified all three Runx factors in premesenchymal condensations and specific skeletal elements, although their functions at this stage prior to bone formation are unknown [130, 334, 338–341]. Additionally, all factors are expressed as isoforms which arise from alternate usage of two promoters and multiple ATGs. For Runx2, the resulting proteins differ in their N-terminal sequence and the isoforms appear to have similar functions, but at different stages of embryonic development or cellular differentiation [340, 342–344]. All Runx factors have two key properties for their functional activity: [1] they are platform proteins that interact with a spectrum of co-regulatory proteins to integrate developmental signals; and [2] Runx proteins contain a unique nuclear matrix-targeting signal (NMTS) which mediates the intranuclear trafficking of Runx2 and their co-regulatory proteins to domains associated with the nuclear scaffold. Co-regulatory proteins include CBFβ, an essential binding partner protein that interacts with the runt homology DNA-binding domain to promote Runx2 interaction with ATGGCT/G motifs in gene promoters. Absence of functional CBFβ in osteoblasts results in bone abnormalities in the mouse [135, 345, 346]. Other partner proteins contribute to gene structure by either promoting increasing transcription by acetylation (e.g., by p300) or inhibiting target gene transcription through deacetylation of histone proteins by histone deacetylase (HDAC) enzymes. Runx2 thereby has a function in modifying chromatin organization of target genes to control accessibility

236 of regulatory factors on target gene promoters. This has been clearly documented for the activation/repression of the osteocalcin gene [347–351]. Such modifications in chromatin structure involving Runx2 are critical for osteoblast differentiation [62] and normal bone development as shown by null mutation or overexpression of HDAC4, leading respectively to premature ossification or inhibition of chondrocyte maturation [352]. The C-terminus activation domain of Runx2 interacts with a variety of signaling proteins including, for example, Smads that mediate BMP/TGFB [58–60]; the Yesassociated protein (YAP) that mediates c-Src signaling [353], homeodomain (HD) proteins [326, 354]; TAZ, a transcriptional co-activator [355]; Groucho/TLE, a developmental signal which suppresses Runx2 activity [356], while a Groucho homolog Grg5 enhances Runx2 transcription and is particularly critical for growth plate development [357]. These co-regulatory factors are organized with Runx as multimeric complexes on target genes in subnuclear domains visualized as punctuate foci. Interestingly, the Runx2 nuclear matrix targeting sequence (~32 amino acids) (NMTS) is also the interacting domain for several of these co-regulatory proteins (Smads, homeodomain proteins, and YAP), emphasizing the importance of organizing gene regulatory complexes in subnuclear domains for tissue-specific control of gene expression [59, 326, 353]. The requirement for this architectural organization of Runx2 transcriptional complexes for bone formation was established in vivo by a knock-in mutation of Runx2 in the mouse [358] and also by several in vitro functional studies. Deletion of the C-terminus targeting and co-regulatory protein interaction domain results in a lethal phenotype and absence of mineralized skeleton analogous to the Runx2-null mouse. There is a requirement for Runx2 recruitment of Smad co-regulatory factors to Runx domains to execute the BMP/TGFβ transcriptional signal for target gene regulation [58, 59]. Of further significance for this critical property of Runx2 is the observation that Runx2 subnuclear targeting deficient mutant proteins, when expressed in metastatic breast cancer cells, inhibit the expression of Runx2 target genes and block the osteolytic properties of metastatic cell lines in bone [359, 360]. Thus these specialized properties of Runx2 provide a basis for controlling accessibility of promoter elements to cognate transcription factors by chromatin remodeling and supporting integration of activities in response to physiologic signals. Taken together, these findings have demonstrated a unique mechanism for tissue-specific control of gene expression through the formation of transcriptional complexes in nuclear microenvironments.

JANE B. LIAN AND GARY S. STEIN 3. Cooperative Control of Gene Transcription Through Interactions at Distinct Promoter Elements: Contribution of C/EBP and Steroid Response Elements

CAAT enhancer-binding proteins include a family of transcription factors that bind to specific motifs and function in cell phenotype differentiation. C/EBPα plays an essential role in adipogenesis, while C/EBPβ is now appreciated to function as a key component of transcriptional control for bone formation. C/EBPβ is up-regulated during osteoblast differentiation [361] and expression of a homologous C/EBP enhancer-binding protein, DDIT3, was shown to induce osteoblast differentiation [362]. Significantly, targeted expression of a truncated nonfunctional C/EBP isoform in bone results in osteopenia [363]. C/EBP and Runx2 regulatory elements are often located in gene promoters in close proximity to each that support physiologic responsiveness for bone formation. C/EBP and Runx2 proteins can form complexes by a direct protein–protein interaction that results in a 30–50% synergistic increase in osteocalcin expression [361]. This mechanism supports osteoblast-specific expression of other genes, such as sclerostin [364]. Other examples of cooperative effects between transcription factors are the requirements of Runx2 and cfos interaction for PTH responsiveness on the MMP13 promoter [365]. Large multimeric complexes are also formed at steroid hormone response elements, including VDRE and ERE motifs. The vitamin D and estrogen receptor complexes recruit chromatin remodeling factors that can bridge with transcriptional complexes at neighboring elements. In this regard, the Runx2 site adjacent to the VDRE interacts with p300 to facilitate vitamin Dmediated enhancement of osteocalcin gene expression [351]. Runx2 also mediates estrogen responses at ERE motifs [366]. In this manner, physiologic control of gene expression is coordinated among distinct regulatory elements on promoters and in a tissue-specific manner.

V. THE OSTEOCLAST: A FUNCTIONALLY UNIQUE CELL FOR PHYSIOLOGICALLY REGULATED RESORPTION OF BONE MINERAL The osteoclast is a multinucleated cell with the ability to degrade mineralized tissues dependent upon cellular attachment to the bone substratum. Osteoclasts are large cells (~100 µm) and appear in histologic sections of bone generally in resorption pits called Howships lacunae. Nuclei numbers are variable and in normal human osteoclasts in

Chapter 14 The Cells of Bone

the range of 4–10 can be seen. Nuclei numbers appear to reflect osteoclast activity and can number as many as 20–50 nuclei in Paget’s disease [367]. It appears from a limited number of studies that all osteoclast nuclei are equivalently transcriptionally active with the components of the transcriptional machinery, including RNA processing and splicing functions organized in subnuclear domains in each nucleus [368, 369]. The osteoclast is a highly polarized cell raising questions as to whether specific nuclei in osteoclasts express subsets of genes reflecting different functional activities or whether they have equivalent functions. Two components of the bone microenvironment are essential for osteoclastogenesis: (1) the mineralized bone tissue is necessary for recruitment, attachment, and activation of a functional osteoclast; and (2) osteoblast lineage cells are required for fusion of the mononuclear precursor to the multinucleated osteoclast and are responsive cells to the calcitrophic hormones which promote osteoclastic resorption. Microfractures that need repair or mechanical forces that signal adaptive changes in bone architecture result in a change in stress lines that provide a road map for homing of osteoclasts to the appropriate site [299]. The requirement for stromal osteoprogenitors or osteoblasts in mediating osteoclast differentiation is linked to the production of cytokines, CSF-1/M-CSF, RANKL, OPG, and several interleukins essential for regulating development of pre-osteoclasts and the multinucleated phenotype.

A. Cellular Features of Bone Resorption The highly specialized morphological features of the osteoclast accommodate its unique function to resorb bone and calcified cartilage. The structural properties of the osteoclast are recognized only when the cell is activated, a process that requires attachment to the mineralized surface, which induces a distinct organization of the cytoskeleton [370–372]. Characteristic features of the osteoclast that result from attachment are: (1) appearance of the sealing zone which is a podosome or focal adhesion complex, a specialized cell–extracellular matrix adhesion structure which allows the osteoclast to form a tight seal against the bone surface; (2) formation of a highly convoluted plasma membrane, the “ruffled border” on the apical bone surface (between the sealing zones); (3) a distinct organization of the cytoskeleton; (4) projections of the basal lateral membrane for secretion of minerals into the vascular space; (5) polarization of the osteoclast nuclei at the basolateral surface; and (6) numerous mitochondria to support activity of the proton pumps, as well

237 as primary lysosomal organelles for secretion of enzymes that will degrade the ECM and secondary lysosomal particles removing matrix debris. Mechanisms contributing to these functional activities in relation to osteoclast cell structure will be briefly described. Genetic and in vitro studies have defined the molecular components of the signaling pathways controlling osteoclast activity which have been recently reviewed in great detail [373, 374]. The adhesion of recruited osteoclasts to the bone surface involves classical integrin receptors αVβ3 and results in the activation of several signaling cascades and distinct morphological features of resorbing osteoclasts. Disruption of integrin mediated adhesion results in inhibition of bone resorption [375, 376]. Following αVβ3 interaction with RGD matrix proteins (vitronectin, osteopontin), small GTPases (Rho and Rac) regulate cytoskeletal re-organization which leads to cell spreading, formation of the sealing zone, and definition of the ruffled membrane [377, 378]. The αVβ3 integrin initially is randomly distributed on the plasma membrane prior to osteoclast activation, then translocates to the bone matrix attachment site, and associates with the filamentous actin in a complex forming a podosome analogous to a focal adhesion structure. The podosome structures completely surround the ruffled membrane rich in F-actin filaments that are oriented perpendicular to the bone surface forming a ring. The actin ring can be used to distinguish between resorbing osteoclasts and nonresorbing cells. Upon osteoclast adhesion to the bone surface, the proto-oncogene c-Src selectively binds the β3 integrin for transduction of the c-Src-dependent polarization signal separating the apical and basal lateral membranes [378–380]. C-Src is a nonreceptor tyrosinase kinase enzyme having an essential role in activating quiescent osteoclasts for bone resorption by formation of the ruffled membrane and polarization of the nuclei in the activated osteoclast [380]. The c-Src−/− mutant mice have many multinucleated cells present in bone with some properties of the osteoclast, but lack function because they are incapable of forming a ruffled border [381, 382]. Both c-Src dependent tyrosine phosphorylation and activation of c-Src result in formation of a complex with a proline-rich tyrosine kinase 2 (Pyk2), a member of the focal adhesion kinase (FAK) family. A major adhesion-induced kinase in osteoclasts is formed with Src, Pyk2, and c-Cbl, a requirement for osteoclast motility [383]. These components of the podosome complex form a positive and negative regulatory loop to regulate cytoskeletal rearrangement, migration, and polarization of the osteoclast [384]. c-Src binds to and phosphorylates c-Cbl which is a β3 ubiquitin ligase and in

238 turn will inhibit Src kinase activity [380]. Therefore, osteoclast migration is controlled by degradation of the podosome complex through the proteasome pathway upon polyubiquitinylation of c-Src. Formation of the complex will occur once again upon osteoclast adhesion [373]. Through these mechanisms, cyclic events of osteoclast activation and migration allow for bone formation at the resorbing site to maintain bone mass. Thus, the initial integrin-mediated adhesion attachment of the osteoclast involves a highly regulated and coordinated series of signaling events for cell attachment and structural modifications that mediate osteoclast activation and migration. The active process of bone resorption requires additional cell structural changes. Attachment of the osteoclast to bone also results in tubulin polymerization for movement of intracellular acidic vesicles along the microtubules to be targeted to the ruffled membrane. Insertion of the H+-ATPase-containing proton pump-bearing vesicles into the apical membrane leads to the infoldings that characterize the ruffled membrane, the actual resorbing component facing the resorption lacunae. The ruffled membrane is compartmentalized with respect to transcytotic vesicles forming at the center of the lacunae and endocytotic vesicles located more peripherally [385]. The tight seal of the surrounding plasma membrane to the bone surface, described above, is necessary to form a proton-impermeable acidic environment in the lacunae for degradation of mineralized bone. Acidification of the resorption lacunae is accomplished by the activities of carbonic anhydrase II and protons generated through the membrane bound H+-ATPase [386, 387]. Mice deficient in the ATP6i complex develop severe osteopetrosis due to the loss of acidification [388]. Various ion channels, chloride/carbonate, and sodium/proton antiporters, maintain cellular electroneutrality (reviewed in references [389, 390]). Osteoclasts are rich in matrix-degrading enzymes, particularly cathespin K and matrix metalloproteinases (MMPs) and a tartrate-resistant acid phosphatase (TRAP) isoenzyme which provides a histochemical marker commonly used to identify osteoclast activity. Genetic studies have identified their critical functions in osteoclastic resorption of bone. Cathepsin K-null mice not only have excessive trabeculae, but an altered ultrastructure of osteoclasts. Recently a human mutation in cathepsin K was identified in a syndrome of pychondystosis [391, 392]. TRAP-null mice revealed that cytoplasmic vesicles accumulate in TRAP−/− osteoclasts suggesting a functional role in modulating vesicular transport [393]. The MMP9null mouse exhibits a delay in osteoclast recruitment [394]. Enrichment of the matrix metalloproteinase MMP9 is a marker of chondroclasts and stimulates MMP13 in hypertrophic chondrocytes which more efficiently degrades

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unmineralized cartilage proteins. MMP9 facilitates bioavailability of the vascular endothelial growth factor, thus recruiting osteoclasts for normal development and long bone growth in conjunction with vascularization of the growth plate [395]. This complexity of osteoclast cellular features is required and coordinated for recruitment of the multinucleated cell to the bone surface for its boneresorbing function.

B. Molecular Mechanisms Mediating Osteoclast Ontogeny The multinucleated osteoclast forms only through the fusion of mononuclear precursors. Proliferation of osteoclasts has never been documented. The naturally occurring animal models of osteopetroses [396], null mutation mouse models [160], and bone marrow transplant or splenic parabiosis studies have firmly established the hematopoietic origin of the mononuclear precursors. Studies that determined expression of cell-surface antigens and phenotypic markers of pre-osteoclast lineage cells, as well as those studies which defined factors regulating commitment to a pre-osteoclast phenotype and maturation to the osteoclast [397, 398], show that the process of osteoclast differentiation involves a number of steps (Fig. 3). An osteoclast-specific cell line is formed at some point in time after formation of the colony-forming unit (CFU) for granulocytes and macrophages. Recently, however, consideration has been given to the formation of osteoclasts from pro-B cells [164]. Historically the generation of osteoclasts in vitro required co-culture of hematopoietic marrow or spleen cells with marrow stromal cells or an osteoblastic cell line. Stromal osteoblastic cells are the major source of two cytokines essential for osteoclastogenesis, the colonystimulating factor M-CSF (also referred to a CSF-1) and a membrane-bound TNF-related cytokine RANKL (receptor activator of NF-κB ligand)/TRANCE/TNFSF11 (tumor necrosis factor ligand superfamily member 11) (a type II transmembrane protein). Both factors are potent stimulators of bone resorption and participate in osteoclastogenesis at early and late stages. M-CSF interacts with the c-Fms receptor on hematopoietic lineage cells initiating a cascade of events for preosteoclast expansion and fusion. c-Fms is a tyrosine kinase receptor which recruits c-Src and phosphatidylinositol-3 kindase (PI-3K), resulting in proliferation and survival of osteoclast precursor cells [399]. The significance of M-CSF for osteoclast function was first established by neutralizing antibodies against M-CSF, which completely inhibited multinucleated cell formation in mouse bone marrow cultures and later by

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Figure 3 Regulation of osteoclastogenesis. (A) Stages in progression of osteoclast differentiation are controlled by transcription factors (Pu.1, cFos , NFAT, NF-κB, FRA1), cytokines (interleukins, TGFβ, RankL), hormones (PTH, 1,25(OH)2 D3, E2) and extra- and intracellular signaling factors (integrins, c-Src, Traf6). (B) Markers frequently used to identify the phenotype of osteoclast lineage cells. CT-R, calcitonin receptor; TRAP, tartrate-resistant acid phosphatase; Cl C-7, specific chloride channel; CAII, carbonic anhydrase.

the finding that recombinant M-CSF injections cure the osteopetrosis in the OP/OP mouse, which is characterized by production of a nonfunctional CSF-1 protein [389, 400–402]. The M-CSF-1/c-Fms receptor interaction signals through the intracellular p38 MAPK, which leads to phosphorylation of the microphalmia-associated transcription factor (MITF) for transcription of target genes that include carbonic anhydrase II and tartrate-resistant acid phosphatase, markers of mature osteoclasts [403, 404]. Spontaneous Mitf mutations in quail and null mutation in mouse (Mitf Mi/Mi) result in an absence of osteoclasts and osteopetrosis [405, 406]. M-CSF and αvβ3 integrin interactions also occur, resulting in phosphorylation of p130(Cas) and c-cbl and consequent p130(Cas) association with Pyk2, thereby contributing to osteoclast attachment and activation [407–409]. The discoveries of RANKL which interacts with RANK on osteoclast precursors and the osteoclast inhibitory factor, osteoprotegerin (OPG), provided insight into mechanisms for the essential role of osteoblasts in mediating osteoclastogenesis. RANKL is produced by osteoblastic stromal cells and, notably, by activated T lymphocytes thereby contributing to bone resorption in inflammatory disease states [410]. RANKL was demonstrated to have competency for inducing osteoclast formation from hematopoietic cells in the absence of stromal cells, allowing

osteoclasts to be generated in vitro from precursor cells without co-culture. Mice deficient in either RANK (encoded by the gene TNFR-SFIIA) or RANKL, and human mutations in the RANK receptor exhibit the same phenotype, osteopetrosis (reviewed in reference [373]. RANKL activation of RANK signaling is transduced by the binding of TRAFs (TNF receptor associated cytoplasmic factors) to the cytoplasmic domain of RANK (TNFRSFIIA). Although multiple TRAFs are present in the cells, only the TRAF6-null mutation results in osteopetrosis [411–413]. RANKL interaction with RANK on osteoclast precursors activates gene expression through signal transduction pathways resulting mainly in NF-κB- and AP-1-mediated transcriptional events (illustrated in Fig. 4). Together then, the RANK/RANKL and cFms/ CSF-1 receptor/ligand pairs represent factors required for coupling stromal/osteoblastic cells to the formation of osteoclasts, while OPG attenuates resorption by inhibiting osteoclastogenesis. Regulation of osteoclastogenesis is achieved at multiple levels. Osteoprotegerin (OPG) encoded by TNFR-SIIB is a soluble decoy protein with strong homology to RANK receptors expressed and secreted by several tissues, including bone, cartilage, kidney, blood vessels, and osteoblast lineage cells [414, 415]. OPG functions as a soluble decoy receptor for RANKL and thereby blocks RANKL-mediated

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Figure 4

Signaling pathways and transcriptional targets of osteoclast differentiation. The extracellular signal is shown in bold and transcription factor targets in grey. Not all intermediates are shown. See text for details. Several extracellular signals can be transduced through common intracellular pathways. PI3k/Akt signaling leads to cell survival, ERK signaling also results in cell survival and proliferation, while the IKK/NF-κB pathway supports osteoclast resorption. ITAM, immunoreceptor tyrosine based activation motif; SHIP, Src homology 2-containing inositol phosphatase negatively regulates osteoclast formation [528].

osteoclast formation and survival of pre-existing osteoclasts [416]. Overexpression of OPG in transgenic mice resulted in severe osteopetrosis as a result of inhibition of the terminal stages of osteoclast differentiation [414, 415, 417, 418]. A spectrum of cytokines also regulates osteoclast differentiation and activity. The stromal cell-derived growth factor (SDF-1) recruits osteoclast precursors by inducing MMP and chemotaxis [419]. The prostaglandin E[2] cytokine, also produced by stromal cells, acts as a potent stimulator of bone resorption and signals through the PGE2 receptor that exists as four subtypes. Null mutation of the EP2 and Ep4 receptors confirms the importance of PGE2 for osteoclast formation [420, 421]. The cytokines II-6 and II-11, like TNFα and IL-1 that are predominantly derived from monocytes, have mitogenic effects on the mononuclear osteoclast precursor. TNFα also promotes osteoclast survival by stimulating phosphorylation of the Akt and ERK, thereby increasing the activity of these pathways [422]. In addition, IL-1 and TNF feed back on stromal cells to regulate their production of CSF-1 and IL-11 (reviewed in references [389, 423–425]). IL-6 and related cytokines that include (but are not limited to) IL-11, oncostatin M, and leukuemia-inhibitory factors, which all enhance osteoclast differentiation. The selective effects of these cytokines occur through the same signal transduction pathway [389, 397, 426, 427]. The membrane-bound cell-surface receptor consists of two components, a specific ligand-binding subunit and a

second subunit that participates in the signaling cascade and is the transducing protein gp130 [424, 428]. The signal transduction cascade includes the nonreceptor tyrosine kinases known as Janus kinases (JAKs), which in turn phosphorylate components of the receptor kinases, and a series of cytoplasmic proteins designated signal transducers and activators of transcription (STATs). Specific cytokines also inhibit osteoclastogenesis [429]. IL-18, initially described as a T-cell cytokine that promotes interferon gamma and production by lymphocytes, was shown to be an important negative regulator of osteoclast formation. It is expressed in osteoblasts but functions via M-CSF [430, 431]. IL-7, a lymphokine secreted from T cells that stimulates production of M-CSF and RANKL, was thought to activate bone resorption. However, evidence from IL-7R-null mice indicates the cytokine is a direct inhibitor of osteoclast formation. The IL-7-deficient mice exhibit an increase in osteoclast number and confirmed by in vitro and ex vivo studies [429].

C. Hormonal Regulation and Transcriptional Control of Osteoclast Differentiation and Activity The calcitrophic hormones 1,25(OH)2D3 and parathyroid hormone (PTH) influence osteoclastogenesis at multiple stages of differentiation and activation through receptors on osteoblast lineage cells. It had been very well

Chapter 14 The Cells of Bone

established that 1,25(OH)2D3 induces differentiation of mouse leukemia cells into macrophages and promotes their activation and fusion [432]. However these cells cannot form resorption pits on dentin slices but required the presences of osteoblastic cells. It is now appreciated that 1,25(OH)2D3, as does PTH and numerous stimulators of bone resorption, including glucocorticoids, T4/T3, TNFα, IL-1, IL-11, and PGE-2, regulate osteoblast expression of RANKL. To further ensure osteoclastogenesis, the calcitrophic hormones 1,25(OH)2D3 and PTH, as well as glucocorticoids, also inhibit OPG production. These hormones also modulate the synthesis and secretion of CSF-1, IL-1, IL-6, and TNFα that have mitogenic effects on osteoclast progenitors. However, direct effects of hormones on osteoclasts should not be dismissed. For example, a VDR-binding protein TAFII-17 was recently found to be expressed by Pagetic osteoclast precursors, but not normal pre-osteoclasts [433]. Although Pagetic osteoclast lineages are characterized by the measles virus nucleocapsid protein (mVNP) responsible for the formation of the large size and numbers of osteoclasts, VDR-TAF II-17 interactions may contribute to the osteoclastogenesis in Paget’s disease. PTH is an essential regulator of calcium homeostasis, which induces a cAMP signal through G protein-coupled receptors (see Chapters 16 and 17). Interestingly, the PTH-related protein (PTHrP) influences osteoclastogenesis after birth and in rapidly turned-over tissues [434]. PTH and PTHrP have homologous N-terminal sequences (amino acids 8–13), which are required for binding to the shared PTH/PTHrP receptor. As such, PTHrP also serves as an inducer of bone resorption and hypercalcemia [435–437]. The majority of evidence supports an indirect activity of PTH and PTHrP on bone resorption, mediated through stromal and osteoblast lineage cells [438]. However, there are reports of osteoclasts expressing PTH/PTHrP receptors [246, 439–444]. The mechanisms by which PTH/PTHrP affects bone resorption at high catabolic concentrations versus osteoblast activity at low anabolic concentration are being identified [445]. Resorptive activity is controlled through regulation of RANKL and OPG [253, 446, 447], while anabolic activity is mediated by AP-1 factors [448] through an IGF-1 dependent mechanism, as described earlier [449, 450]. Estrogen has a broad range of effects in regulating bone resorption. Estrogen’s protective effects on bone are exerted by several mechanisms. Estrogen inhibits PTHinduced osteoclastogenesis and attachment to bone [444]. The suppression of PTH-stimulated osteoclast activity occurs both by blocking the cAMP-dependent PKA and calcium PKC pathways and by promoting osteoclast apoptosis [444]. Equally important, several cytokines that

241 prolong osteoclast survival, IL1, IL6, II7, TNFα, M-CSF, and RANKL are negatively regulated by estrogens and thereby estrogen increases osteoclast apoptosis [451–455]. At the same time, estrogen stimulates synthesis of antiresorptive factors OPG and TGFβ [456]. Thus, estrogen deficiency will lead to a marked increase in bone turnover. Estrogen effects are indirect functioning through the osteoblast lineage cells, mainly through the ERα, although the ERβ receptor also mediates responses [457] to activate osteoclasts. A few studies have reported that osteoclasts express estrogen receptors [442, 458, 459], while others have not found this result in developing bone [460]. Nonetheless, in isolated osteoclasts, estrogen induces a rapid response on inhibitory response via pp60src signaling [461]. The role of estrogen for bone health in men is being appreciated [443, 462]. Calcitonin, a polypeptide hormone, is secreted by the thyroid C-cells, but also by neuroendocrine cells, in response to hypercalcemia. Calcitonin binding to a G protein-coupled-type calcitonin receptor, which is a characteristic marker of the mature osteoblast, rapidly terminates resorption activity [463, 464]. The inhibition of osteoclast function induces rapid disappearance of the actin rings and subsequent loss of structural morphology. Calcitonin inhibits proton extrusion via PKA and cytoskeletal proteins become reorganized for osteoclast cell retraction [372, 465–468]. A surprising finding from the calcitonin/calcitonin gene-related peptide-null mouse is that absence of calcitonin does not affect bone resorption. A compensatory mechanism is the sensitization of the osteoclast to PTH upon calcitonin withdrawal [469, 470]. Calcitonin (salmon calcitonin) has long been used clinically as an antiresorptive agent and has an added analgesic benefit, although the mechanisms contributing to pain relief are unknown. Recently identified nonhormonal regulatory factors have roles in supporting the resorptive phase of bone remodeling by inhibiting bone formation. Sclerostin, for example, is secreted by osteoclasts and negatively regulates bone formation by functioning as a BMP antagonist [471]. c-Src signaling, essential for activation of osteoclast activity, is a negative regulator of Runx2-mediated osteoblast differentiation [58, 472]. Mechanisms regulating the limit of resorption in Howship lacunae are not well understood, but a combination of pathways is likely to be operative. Calcitonin exerts a rapid response for retraction of the osteoclast in response to physiologic Ca++ concentration [473], and the simultaneous elevated phosphate concentrations induce osteoclast apoptosis, as well as up-regulate OPG, thereby limiting resorptive activity by a combination of events [474]. Amylin, a calcitonin family member, was recently reported

242 to inhibit the fusion of osteoclast precursors through an unidentified receptor [475]. Numerous cytokines (e.g., TNFα, RANKL, and IL-1) prolong the ability of osteoclasts to go through successive rounds of resorption. However, the action of TGFβ and estrogen have a negative effect on osteoclast survival time [476]. In response to hormone- and cytokine-induced signal transduction pathways, the fusion of circulating mononuclear precursors to the multinucleated osteoclasts and osteoclast activity is ultimately regulated by the transcriptional targets of signal transduction pathways that control gene expression for commitment, and differentiation of the mononuclear precursors (Fig. 4). Our understanding of the transcriptional mediators that drive osteoclast differentiation and regulate their activation derive largely from characterization of murine animal models that manifest the osteopetrotic disorders of bone resorption [396, 477, 478]. Many nonsteroidal transcriptional regulators are essential for osteoclast differentiation and activity as illustrated in Figure 3. The required extracellular signals for osteoclastogenesis (M-CSF, RANKL, IL-1, αvβ3, Ca++/ calmodulin) are transduced via intracellular proteins to transcriptional control of target genes (illustrated in Fig. 4). The first transcription factor which was implicated in osteoclastogenesis, c-fos, was revealed by a marked osteopetrosis disorder in the null mutant mouse [479]. Bone sections revealed an abundance of macrophages but an absence of osteoclasts suggesting that c-fos promotes proliferation and differentiation of a specific mononuclear precursor committed to the osteoclast pathway [480, 481]. Transgenic mice in which the myeloid and B lymphoid transcription factor PU.1 is ablated, fail to generate macrophages or osteoclasts and also develop osteopetrosis [482]. The mutant mice are successfully rescued by marrow transplantation supporting the concept of the common lineage of osteoclasts and macrophages and that the PU.1 factor is involved in establishing one of the early myeloid cells. Additionally, PU.1 directs expression of M-CSF [483]. RANKL and II-1 signaling induce the transcription factor NF-κB. Mice deficient in both subunits (p50 and p52) of NF-κB result in osteopetrosis [484]. The nuclear factor of activated T cell cytoplasmic (NFAT2/NFATc) transcription factors have been identified as targets of RANKL (NFAT1/NFATc2) in promoting osteoclast differentiation [485–487] and calcium– calmodulin signaling (NFAT2/NFATc1). Transgenic mice overexpressing NFAT1 (NFAT2c) in osteoclasts result in decreased trabecular bone mass [488]. Upon dephosphorylation, these transcription factors translocate from the cytoplasm to the nucleus. The NFATs are now considered transcriptional key regulators of gene expression responsive to signaling pathways essential for osteoclast differentiation.

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The MI gene product, Mitf (mentioned earlier), encodes a member of the basic helix-loop-helix leucine zipper protein family that is one of the transcription factor targets of MAPK/p38 signaling. Mitf and the Tfe3 member of this family have redundant roles in osteoclast formation [489, 490]. MI/MI mice develop osteopetrosis, although many TRAP- positive osteoclasts are present in trabecular bones of these mice. However, the osteoclasts fail to form ruffle borders [491], similar to the Src−/− mice. In the final analyses of the regulatory controls for osteoclastogenesis, many levels are operative from the initial ECM microenvironment to the cellular stimulates and intracellular signal transduction pathways that induce transcriptional events.

VI. PERSPECTIVES Our understanding of the biological properties of bone cells is rapidly evolving. Complex regulatory mechanisms that are operative in the establishment of skeletal stem cell populations, as well as in commitment to the osteoblast or osteoclast phenotypes, are being clarified. Although systemic and local factors that promote and regulate differentiation of lineage-specific phenotypes are being identified, a goal for the future is a better understanding of the mechanisms that facilitate interactions between subpopulations of osteoblasts and osteoclasts to support maintenance of bone mass and mineral homeostasis. Pursuing the identity of cell type-specific proteins and transcription factors that regulate their expression has been instructive. As the signaling pathways that mediate expression of genes controlling proliferation and differentiation of bone cells are further clarified, a new generation of options for diagnosis and treatment of skeletal disorders will emerge.

Acknowledgments This work was supported by NIH grants AR39588, DE12528, P01 PO1 AR48818, and P30 DK32520. The contents of this manuscript are solely the responsibility of the authors and do not necessarily represent the official views of the National Institutes of Health.

References 1. Karaplis, A. (2002). Embryonic development of bone and the molecular regulation of intramembranous and endochondral bone formation. In “Principles of Bone Biology” (J. P. Bilezikian, L. G. Raisz, and G. A. Rodan, eds.), pp. 33–58. Academic Press, San Diego, CA. 2. Olsen, B. R., Reginato, A. M., and Wang, W. (2000). Bone development. Annu. Rev. Cell Dev. Biol. 16, 191–220.

Chapter 14 The Cells of Bone 3. Zakany, J., Kmita, M., and Duboule, D. (2004). A dual role for Hox genes in limb anterior-posterior asymmetry. Science 304, 1669–1672. 4. Goodman, F. R. (2002). Limb malformations and the human HOX genes. Am. J. Med. Genet. 112, 256–265. 5. Vargesson, N., Patel, K., Lewis, J., and Tickle, C. (1998). Expression patterns of Notch1, Serrate1, Serrate2 and Delta1 in tissues of the developing chick limb. Mech. Dev. 77, 197–199. 6. Yamaguchi, A., Komori, T., and Suda, T. (2000). Regulation of osteoblast differentiation mediated by bone morphogenetic proteins, hedgehogs, and Cbfa1. Endocr. Rev. 21, 393–411. 7. Harfe, B. D., Scherz, P. J., Nissim, S., Tian, H., McMahon, A. P., and Tabin, C. J. (2004). Evidence for an expansion-based temporal Shh gradient in specifying vertebrate digit identities. Cell 118, 517–528. 8. Spinella-Jaegle, S., Rawadi, G., Kawai, S., Gallea, S., Faucheu, C., Mollat, P., Courtois, B., Bergaud, B., Ramez, V., Blanchet, A. M., Adelmant, G., Baron, R., and Roman-Roman, S. (2001). Sonic hedgehog increases the commitment of pluripotent mesenchymal cells into the osteoblastic lineage and abolishes adipocytic differentiation. J. Cell Sci. 114, 2085–2094. 9. Depew, M. J., Lufkin, T., and Rubenstein, J. L. (2002). Specification of jaw subdivisions by Dlx genes. Science 298, 381–385. 10. Ferrari, D., Sumoy, L., Gannon, J., Sun, H., Brown, A. M., Upholt, W. B., and Kosher, R. A. (1995). The expression pattern of the Distal-less homeobox-containing gene Dlx-5 in the developing chick limb bud suggests its involvement in apical ectodermal ridge activity, pattern formation, and cartilage differentiation. Mech. Dev. 52, 257–264. 11. Robledo, R. F., Rajan, L., Li, X., and Lufkin, T. (2002). The Dlx5 and Dlx6 homeobox genes are essential for craniofacial, axial, and appendicular skeletal development. Genes Dev. 16, 1089–1101. 12. Thesleff, I., Vaahtokari, A., Vainio, S., and Jowett, A. (1996). Molecular mechanisms of cell and tissue interactions during early tooth development. Anat. Rec. 245, 151–161. 13. Yoon, B. S. and Lyons, K. M. (2004). Multiple functions of BMPs in chondrogenesis. J. Cell Biochem. 93, 93–103. 14. Boulet, A. M., Moon, A. M., Arenkiel, B. R., and Capecchi, M. R. (2004). The roles of Fgf4 and Fgf8 in limb bud initiation and outgrowth. Dev. Biol. 273, 361–372. 15. Marie, P. J. (2003). Fibroblast growth factor signaling controlling osteoblast differentiation. Gene 316, 23–32. 16. ten Dijke, P., Fu, J., Schaap, P., and Roelen, B. A. (2003). Signal transduction of bone morphogenetic proteins in osteoblast differentiation. J. Bone Joint Surg. Am. 85-A Suppl 3, 34–38. 17. Westendorf, J. J., Kahler, R. A., and Schroeder, T. M. (2004). Wnt signaling in osteoblasts and bone diseases. Gene 341, 19–39. 18. Iseki, S., Wilkie, A. O., and Morriss-Kay, G. M. (1999). Fgfr1 and Fgfr2 have distinct differentiation- and proliferation-related roles in the developing mouse skull vault. Development 126, 5611–5620. 19. Delezoide, A. L., Benoist-Lasselin, C., Legeai-Mallet, L., Le Merrer, M., Munnich, A., Vekemans, M., and Bonaventure, J. (1998). Spatio-temporal expression of FGFR 1, 2 and 3 genes during human embryo-fetal ossification. Mech. Dev. 77, 19–30. 20. Rice, D. P., Rice, R., and Thesleff, I. (2003). Molecular mechanisms in calvarial bone and suture development, and their relation to craniosynostosis. Eur. J. Orthod. 25, 139–148. 21. Sun, X., Mariani, F. V., and Martin, G. R. (2002). Functions of FGF signalling from the apical ectodermal ridge in limb development. Nature 418, 501–508. 22. Zhou, Y. X., Xu, X., Chen, L., Li, C., Brodie, S. G., and Deng, C. X. (2000). A Pro250Arg substitution in mouse Fgfr1 causes increased expression of Cbfa1 and premature fusion of calvarial sutures. Hum. Mol. Genet. 9, 2001–2008. 23. Hajihosseini, M. K., Lalioti, M. D., Arthaud, S., Burgar, H. R., Brown, J. M., Twigg, S. R., Wilkie, A. O., and Heath, J. K. (2004). Skeletal development is regulated by fibroblast growth factor receptor 1 signalling dynamics. Development 131, 325–335.

243 24. Rutland, P., Pulleyn, L. J., Reardon, W., Baraitser, M., Hayward, R., Jones, B., Malcolm, S., Winter, R. M., Oldridge, M., Slaney, S. F., Pook, M. D., and Wilkie, A. O. M. (1995). Identical mutations in the FGFR2 gene cause both Pfeiffer and Crouzon syndrome phenotypes. Nat. Genet. 9, 173–176. 25. Eswarakumar, V. P., Horowitz, M. C., Locklin, R., Morriss-Kay, G. M., and Lonai, P. (2004). A gain-of-function mutation of Fgfr2c demonstrates the roles of this receptor variant in osteogenesis. Proc. Natl Acad. Sci. USA 101, 12555–12560. 26. Baroni, T., Carinci, P., Lilli, C., Bellucci, C., Aisa, M. C., Scapoli, L., Volinia, S., Carinci, F., Pezzetti, F., Calvitti, M., Farina, A., Conte, C., and Bodo, M. (2005). P253R fibroblast growth factor receptor-2 mutation induces RUNX2 transcript variants and calvarial osteoblast differentiation. J. Cell Physiol. 202, 524–535. 27. Tanimoto, Y., Yokozeki, M., Hiura, K., Matsumoto, K., Nakanishi, H., Matsumoto, T., Marie, P. J., and Moriyama, K. (2004). A soluble form of fibroblast growth factor receptor 2 (FGFR2) with S252W mutation acts as an efficient inhibitor for the enhanced osteoblastic differentiation caused by FGFR2 activation in Apert syndrome. J. Biol. Chem. 279, 45926–45934. 28. Ornitz, D. M. and Marie, P. J. (2002). FGF signaling pathways in endochondral and intramembranous bone development and human genetic disease. Genes Dev. 16, 1446–1465. 29. Amizuka, N., Davidson, D., Liu, H., Valverde-Franco, G., Chai, S., Maeda, T., Ozawa, H., Hammond, V., Ornitz, D. M., Goltzman, D., and Henderson, J. E. (2004). Signalling by fibroblast growth factor receptor 3 and parathyroid hormone-related peptide coordinate cartilage and bone development. Bone 34, 13–25. 30. Deng, C., Wynshaw-Boris, A., Zhou, F., Kuo, A., and Leder, P. (1996). Fibroblast growth factor receptor 3 is a negative regulator of bone growth. Cell 84, 911–921. 31. Valverde-Franco, G., Liu, H., Davidson, D., Chai, S., ValderramaCarvajal, H., Goltzman, D., Ornitz, D. M., and Henderson, J. E. (2004). Defective bone mineralization and osteopenia in young adult FGFR3-/mice. Hum. Mol. Genet. 13, 271–284. 32. Abzhanov, A., and Tabin, C. J. (2004). Shh and Fgf8 act synergistically to drive cartilage outgrowth during cranial development. Dev. Biol. 273, 134–148. 33. Laufer, E., Nelson, C. E., Johnson, R. L., Morgan, B. A., and Tabin, C. (1994). Sonic hedgehog and Fgf-4 act through a signaling cascade and feedback loop to integrate growth and patterning of the developing limb bud. Cell 79, 993–1003. 34. Liu, Z., Xu, J., Colvin, J. S., and Ornitz, D. M. (2002). Coordination of chondrogenesis and osteogenesis by fibroblast growth factor 18. Genes Dev. 16, 859–869. 35. Ohbayashi, N., Shibayama, M., Kurotaki, Y., Imanishi, M., Fujimori, T., Itoh, N., and Takada, S. (2002). FGF18 is required for normal cell proliferation and differentiation during osteogenesis and chondrogenesis. Genes Dev. 16, 870–879. 36. Sullivan, R., and Klagsbrun, M. (1985). Purification of cartilagederived growth factor by heparin affinity chromatography. J. Biol. Chem. 260, 2399–2403. 37. Hurley, M. M., Abreu, C., Gronowicz, G., Kawaguchi, H., and Lorenzo, J. (1994). Expression and regulation of basic fibroblast growth factor mRNA levels in mouse osteoblastic MC3T3-E1 cells. J. Biol. Chem. 269, 9392–9396. 38. Tanaka, H., Ogasa, H., Barnes, J., and Liang, C. T. (1999). Actions of bFGF on mitogenic activity and lineage expression in rat osteoprogenitor cells: effect of age. Mol. Cell Endocrinol. 150, 1–10. 39. Debiais, F., Lefevre, G., Lemonnier, J., Le Mee, S., Lasmoles, F., Mascarelli, F., and Marie, P. J. (2004). Fibroblast growth factor-2 induces osteoblast survival through a phosphatidylinositol 3-kinase-dependent, -beta-catenin-independent signaling pathway. Exp. Cell Res. 297, 235–246. 40. Xiao, L., Liu, P., Sobue, T., Lichtler, A., Coffin, J. D., and Hurley, M. M. (2003). Effect of overexpressing fibroblast growth factor 2 protein

244

41.

42.

43.

44.

45.

46.

47. 48. 49.

50.

51.

52.

53.

54.

55.

56.

57.

isoforms in osteoblastic ROS 17/2.8 cells. J. Cell Biochem. 89, 1291–1301. Montero, A., Okada, Y., Tomita, M., Ito, M., Tsurukami, H., Nakamura, T., Doetschman, T., Coffin, J. D., and Hurley, M. M. (2000). Disruption of the fibroblast growth factor-2 gene results in decreased bone mass and bone formation. J. Clin. Invest. 105, 1085–1093. Ahn, K., Mishina, Y., Hanks, M. C., Behringer, R. R., and Crenshaw, E. B., III (2001). BMPR-IA signaling is required for the formation of the apical ectodermal ridge and dorsal-ventral patterning of the limb. Development 128, 4449–4461. Canalis, E., Economides, A. N., and Gazzerro, E. (2003). Bone morphogenetic proteins, their antagonists, and the skeleton. Endocr. Rev. 24, 218–235. Derynck, R., and Zhang, Y. E. (2003). Smad-dependent and Smadindependent pathways in TGF-beta family signalling. Nature 425, 577–584. Zhao, M., Harris, S. E., Horn, D., Geng, Z., Nishimura, R., Mundy, G. R., and Chen, D. (2002). Bone morphogenetic protein receptor signaling is necessary for normal murine postnatal bone formation. J. Cell Biol. 157, 1049–1060. Yi, S. E., Daluiski, A., Pederson, R., Rosen, V., and Lyons, K. M. (2000). The type I BMP receptor BMPRIB is required for chondrogenesis in the mouse limb. Development 127, 621–630. Wrana, J. L. (2000). Crossing Smads. Sci. STKE 2000, RE1. Chubinskaya, S., and Kuettner, K. E. (2003). Regulation of osteogenic proteins by chondrocytes. Int. J. Biochem. Cell Biol. 35, 1323–1340. Settle, S. H., Jr., Rountree, R. B., Sinha, A., Thacker, A., Higgins, K., and Kingsley, D. M. (2003). Multiple joint and skeletal patterning defects caused by single and double mutations in the mouse Gdf6 and Gdf5 genes. Dev. Biol. 254, 116–130. Zhang, D., Ferguson, C. M., O’Keefe, R. J., Puzas, J. E., Rosier, R. N., and Reynolds, P. R. (2002). A role for the BMP antagonist chordin in endochondral ossification. J. Bone Miner. Res. 17, 293–300. Wang, C. K., Omi, M., Ferrari, D., Cheng, H. C., Lizarraga, G., Chin, H. J., Upholt, W. B., Dealy, C. N., and Kosher, R. A. (2004). Function of BMPs in the apical ectoderm of the developing mouse limb. Dev. Biol. 269, 109–122. Archer, C. W., Dowthwaite, G. P., and Francis-West, P. (2003). Development of synovial joints. Birth Defects Res. C. Embryo Today 69, 144–155. Grimsrud, C. D., Romano, P. R., D’Souza, M., Puzas, J. E., Reynolds, P. R., Rosier, R. N., and O’Keefe, R. J. (1999). BMP-6 is an autocrine stimulator of chondrocyte differentiation. J. Bone Miner. Res. 14, 475–482. Balint, E., Lapointe, D., Drissi, H., van der Meijden, C., Young, D. W., van Wijnen, A. J., Stein, J. L., Stein, G. S., and Lian, J. B. (2003). Phenotype discovery by gene expression profiling: mapping of biological processes linked to BMP-2-mediated osteoblast differentiation. J. Cell Biochem. 89, 401–426. Banerjee, C., Javed, A., Choi, J.-Y., Green, J., Rosen, V., van Wijnen, A. J., Stein, J. L., Lian, J. B., and Stein, G. S. (2001). Differential regulation of the two principal Runx2/Cbfa1 N-terminal isoforms in response to bone morphogenetic protein-2 during development of the osteoblast phenotype. Endocrinology 142, 4026–4039. Harris, S. E., Guo, D., Harris, M. A., Krishnaswamy, A., and Lichtler, A. (2003). Transcriptional regulation of BMP-2 activated genes in osteoblasts using gene expression microarray analysis: role of Dlx2 and Dlx5 transcription factors. Front Biosci. 8, s1249–s1265. Lee, K. S., Kim, H. J., Li, Q. L., Chi, X. Z., Ueta, C., Komori, T., Wozney, J. M., Kim, E. G., Choi, J. Y., Ryoo, H. M., and Bae, S. C. (2000). Runx2 is a common target of transforming growth factor beta1 and bone morphogenetic protein 2, and cooperation between runx2 and smad5 induces osteoblast-specific gene expression in the pluripotent mesenchymal precursor cell line C2C12. Mol. Cell Biol. 20, 8783–8792.

JANE B. LIAN AND GARY S. STEIN 58. Zaidi, S. K., Sullivan, A. J., van Wijnen, A. J., Stein, J. L., Stein, G. S., and Lian, J. B. (2002). Integration of Runx and Smad regulatory signals at transcriptionally active subnuclear sites. Proc. Natl Acad. Sci. USA 99, 8048–8053. 59. Afzal, F., Pratap, J., Ito, K., Ito, Y., Stein, J. L., van Wijnen, A. J., Stein, G. S., Lian, J. B., and Javed, A. (2004). Smad function and intranuclear targeting share a Runx2 motif required for osteogenic lineage induction and BMP2 responsive transcription. J. Cell Physiol. [Epub ahead of print]. 60. Zhang, Y. W., Yasui, N., Ito, K., Huang, G., Fujii, M., Hanai, J., Nogami, H., Ochi, T., Miyazono, K., and Ito, Y. (2000). A RUNX2/ PEBP2αA/CBFA1 mutation displaying impaired transactivation and Smad interaction in cleidocranial dysplasia. Proc. Natl Acad. Sci. USA 97, 10549–10554. 61. Yang, S., Wei, D., Wang, D., Phimphilai, M., Krebsbach, P. H., and Franceschi, R. T. (2003). In vitro and in vivo synergistic interactions between the Runx2/Cbfa1 transcription factor and bone morphogenetic protein-2 in stimulating osteoblast differentiation. J. Bone Miner. Res. 18, 705–715. 62. Young, D. W., Pratap, J., Javed, A., Weiner, B., Ohkawa, Y., van Wijnen, A., Montecino, M., Stein, G. S., Stein, J. L., Imbalzano, A. N., and Lian, J. B. (2005). SWI/SNF chromatin remodeling complex is obligatory for BMP2-induced, Runx2-dependent skeletal gene expression that controls osteoblast differentiation. J. Cell Biochem. 94, 720–730. 63. Komori, T., Yagi, H., Nomura, S., Yamaguchi, A., Sasaki, K., Deguchi, K., Shimizu, Y., Bronson, R. T., Gao, Y.-H., Inada, M., Sato, M., Okamoto, R., Kitamura, Y., Yoshiki, S., and Kishimoto, T. (1997). Targeted disruption of Cbfa1 results in a complete lack of bone formation owing to maturational arrest of osteoblasts. Cell 89, 755–764. 64. Otto, F., Thornell, A. P., Crompton, T., Denzel, A., Gilmour, K. C., Rosewell, I. R., Stamp, G. W. H., Beddington, R. S. P., Mundlos, S., Olsen, B. R., Selby, P. B., and Owen, M. J. (1997). Cbfa1, a candidate gene for cleidocranial dysplasia syndrome, is essential for osteoblast differentiation and bone development. Cell 89, 765–771. 65. Banerjee, C., McCabe, L. R., Choi, J.-Y., Hiebert, S. W., Stein, J. L., Stein, G. S., and Lian, J. B. (1997). Runt homology domain proteins in osteoblast differentiation: AML-3/CBFA1 is a major component of a bone specific complex. J. Cell Biochem. 66, 1–8. 66. Ducy, P., Zhang, R., Geoffroy, V., Ridall, A. L., and Karsenty, G. (1997). Osf2/Cbfa1: a transcriptional activator of osteoblast differentiation. Cell 89, 747–754. 67. Chimal-Monroy, J., Rodriguez-Leon, J., Montero, J. A., Ganan, Y., Macias, D., Merino, R., and Hurle, J. M. (2003). Analysis of the molecular cascade responsible for mesodermal limb chondrogenesis: Sox genes and BMP signaling. Dev. Biol. 257, 292–301. 68. Mori-Akiyama, Y., Akiyama, H., Rowitch, D. H., and de Crombrugghe, B. (2003). Sox9 is required for determination of the chondrogenic cell lineage in the cranial neural crest. Proc. Natl Acad. Sci. USA 100, 9360–9365. 69. Semba, I., Nonaka, K., Takahashi, I., Takahashi, K., Dashner, R., Shum, L., Nuckolls, G. H., and Slavkin, H. C. (2000). Positionallydependent chondrogenesis induced by BMP4 is co-regulated by Sox9 and Msx2. Dev. Dyn. 217, 401–414. 70. Zhang, Z., Song, Y., Zhang, X., Tang, J., Chen, J., and Chen, Y. (2003). Msx1/Bmp4 genetic pathway regulates mammalian alveolar bone formation via induction of Dlx5 and Cbfa1. Mech. Dev. 120, 1469–1479. 71. Nakashima, K., Zhou, X., Kunkel, G., Zhang, Z., Deng, J. M., Behringer, R. R., and de Crombrugghe, B. (2002). The novel zinc finger-containing transcription factor osterix is required for osteoblast differentiation and bone formation. Cell 108, 17–29. 72. Logan, C. Y. and Nusse, R. (2004). The Wnt signaling pathway in development and disease. Annu. Rev. Cell Dev. Biol. 20, 781–810. 73. Kawano, Y. and Kypta, R. (2003). Secreted antagonists of the Wnt signalling pathway. J. Cell Sci. 116, 2627–2634.

245

Chapter 14 The Cells of Bone 74. Church, V. L., and Francis-West, P. (2002). Wnt signalling during limb development. Int. J. Dev. Biol. 46, 927–936. 75. Ikeya, M., and Takada, S. (2001). Wnt-3a is required for somite specification along the anteroposterior axis of the mouse embryo and for regulation of cdx-1 expression. Mech. Dev. 103, 27–33. 76. Niemann, S., Zhao, C., Pascu, F., Stahl, U., Aulepp, U., Niswander, L., Weber, J. L., and Muller, U. (2004). Homozygous WNT3 mutation causes tetra-amelia in a large consanguineous family. Am. J. Hum. Genet. 74, 558–563. 77. Lako, M., Strachan, T., Bullen, P., Wilson, D. I., Robson, S. C., and Lindsay, S. (1998). Isolation, characterisation and embryonic expression of WNT11, a gene which maps to 11q13.5 and has possible roles in the development of skeleton, kidney and lung. Gene 219, 101–110. 78. Loganathan, P. G., Nimmagadda, S., Huang, R., Scaal, M., and Christ, B. (2005). Comparative analysis of the expression patterns of Wnts during chick limb development. Histochem. Cell Biol. 79. Hartmann, C., and Tabin, C. J. (2001). Wnt-14 plays a pivotal role in inducing synovial joint formation in the developing appendicular skeleton. Cell 104, 341–351. 80. Yamaguchi, T. P., Bradley, A., McMahon, A. P., and Jones, S. (1999). A Wnt5a pathway underlies outgrowth of multiple structures in the vertebrate embryo. Development 126, 1211–1223. 81. Yang, Y., Topol, L., Lee, H., and Wu, J. (2003). Wnt5a and Wnt5b exhibit distinct activities in coordinating chondrocyte proliferation and differentiation. Development 130, 1003–1015. 82. Topol, L., Jiang, X., Choi, H., Garrett-Beal, L., Carolan, P. J., and Yang, Y. (2003). Wnt-5a inhibits the canonical Wnt pathway by promoting GSK-3-independent {beta}-catenin degradation. J. Cell Biol. 162, 899–908. 83. Hartmann, C., and Tabin, C. J. (2000). Dual roles of wnt signaling during chondrogenesis in the chicken limb. Development 127, 3141–3159. 84. Kawakami, Y., Wada, N., Nishimatsu, S. I., Ishikawa, T., Noji, S., and Nohno, T. (1999). Involvement of Wnt-5a in chondrogenic pattern formation in the chick limb bud. Dev. Growth Differ. 41, 29–40. 85. Church, V., Nohno, T., Linker, C., Marcelle, C., and Francis-West, P. (2002). Wnt regulation of chondrocyte differentiation. J. Cell Sci. 115, 4809–4818. 86. Parr, B. A., and McMahon, A. P. (1995). Dorsalizing signal Wnt-7a required for normal polarity of D-V and A-P axes of mouse limb. Nature 374, 350–353. 87. Holmen, S. L., Zylstra, C. R., Mukherjee, A., Sigler, R. E., Faugere, M. C., Bouxsein, M. L., Deng, L., Clemens, T. L., and Williams, B. O. (2005). Essential role of beta -catenin in post natal bone acquisition. J. Biol. Chem. [Epub ahead of print]. 88. Glass, D. A., Bialek, P., Ahn, J. D., Starbuck, M., Patel, M. S., Clevers, H., Taketo, M. M., Long, F., McMahon, A. P., Lang, R. A., and Karsenty, G. (2005). Canonical wnt signaling in differentiated osteoblasts controls osteoclast differentiation. Dev. Cell 8, 751–764. 89. Day, T. F., Guo, X., Garrett-Beal, L., and Yang, Y. (2005). Wnt/betacatenin signaling in mesenchymal progenitors controls osteoblast and chondrocyte differentiation during vertebrate skeletogenesis. Dev. Cell 8, 739–750. 90. Hill, T. P., Spater, D., Taketo, M. M., Birchmeier, W., and Hartmann, C. (2005). Canonical Wnt/beta-catenin signaling prevents osteoblasts from differentiating into chondrocytes. Dev. Cell 8, 727–738. 91. Boyden, L. M., Mao, J., Belsky, J., Mitzner, L., Farhi, A., Mitnick, M. A., Wu, D., Insogna, K., and Lifton, R. P. (2002). High bone density due to a mutation in LDL-receptor-related protein 5. N. Engl. J. Med. 346, 1513–1521. 92. Little, R. D., Recker, R. R., and Johnson, M. L. (2002). High bone density due to a mutation in LDL-receptor-related protein 5. N. Engl. J. Med. 347, 943–944. 93. Akhter, M. P., Wells, D. J., Short, S. J., Cullen, D. M., Johnson, M. L., Haynatzki, G. R., Babij, P., Allen, K. M., Yaworsky, P. J., Bex, F., and

94.

95.

96.

97.

98.

99.

100.

101.

102.

103.

104.

Recker, R. R. (2004). Bone biomechanical properties in LRP5 mutant mice. Bone 35, 162–169. Babij, P., Zhao, W., Small, C., Kharode, Y., Yaworsky, P. J., Bouxsein, M. L., Reddy, P. S., Bodine, P. V., Robinson, J. A., Bhat, B., Marzolf, J., Moran, R. A., and Bex, F. (2003). High bone mass in mice expressing a mutant LRP5 gene. J. Bone Miner. Res. 18, 960–974. Gong, Y., Slee, R. B., Fukai, N., Rawadi, G., Roman-Roman, S., Reginato, A. M., Wang, H., Cundy, T., Glorieux, F. H., Lev, D., Zacharin, M., Oexle, K., Marcelino, J., Suwairi, W., Heeger, S., Sabatakos, G., Apte, S., Adkins, W. N., Allgrove, J., Arslan-Kirchner, M., Batch, J. A., Beighton, P., Black, G. C., Boles, R. G., Boon, L. M., Borrone, C., Brunner, H. G., Carle, G. F., Dallapiccola, B., De Paepe, A., Floege, B., Halfhide, M. L., Hall, B., Hennekam, R. C., Hirose, T., Jans, A., Juppner, H., Kim, C. A., Keppler-Noreuil, K., Kohlschuetter, A., LaCombe, D., Lambert, M., Lemyre, E., Letteboer, T., Peltonen, L., Ramesar, R. S., Romanengo, M., Somer, H., Steichen-Gersdorf, E., Steinmann, B., Sullivan, B., Superti-Furga, A., Swoboda, W., van den Boogaard, M. J., van Hul, W., Vikkula, M., Votruba, M., Zabel, B., Garcia, T., Baron, R., Olsen, B. R., and Warman, M. L. (2001). LDL receptor-related protein 5 (LRP5) affects bone accrual and eye development. Cell 107, 513–523. Van Wesenbeeck, L., Cleiren, E., Gram, J., Beals, R. K., Benichou, O., Scopelliti, D., Key, L., Renton, T., Bartels, C., Gong, Y., Warman, M. L., de Vernejoul, M. C., Bollerslev, J., and van Hul, W. (2003). Six novel missense mutations in the LDL receptor-related protein 5 (LRP5) gene in different conditions with an increased bone density. Am. J. Hum. Genet. 72, 763–771. Kato, M., Patel, M. S., Levasseur, R., Lobov, I., Chang, B. H., Glass, D. A., Hartmann, C., Li, L., Hwang, T. H., Brayton, C. F., Lang, R. A., Karsenty, G., and Chan, L. (2002). Cbfa1-independent decrease in osteoblast proliferation, osteopenia, and persistent embryonic eye vascularization in mice deficient in Lrp5, a Wnt coreceptor. J. Cell Biol. 157, 303–314. Bodine, P. V., Zhao, W., Kharode, Y. P., Bex, F. J., Lambert, A. J., Goad, M. B., Gaur, T., Stein, G. S., Lian, J. B., and Komm, B. S. (2004). The Wnt antagonist secreted frizzled-related protein-1 is a negative regulator of trabecular bone formation in adult mice. Mol. Endocrinol. 18, 1222–1237. Bennett, C. N., Longo, K. A., Wright, W. S., Suva, L. J., Lane, T. F., Hankenson, K. D., and MacDougald, O. A. (2005). Regulation of osteoblastogenesis and bone mass by Wnt10b. Proc. Natl Acad. Sci. USA 102, 3324–3329. Longo, K. A., Wright, W. S., Kang, S., Gerin, I., Chiang, S. H., Lucas, P. C., Opp, M. R., and MacDougald, O. A. (2004). Wnt10b inhibits development of white and brown adipose tissues. J. Biol. Chem. 279, 35503–35509. Minina, E., Kreschel, C., Naski, M. C., Ornitz, D. M., and Vortkamp, A. (2002). Interaction of FGF, Ihh/Pthlh, and BMP signaling integrates chondrocyte proliferation and hypertrophic differentiation. Dev. Cell 3, 439–449. Schipani, E., Lanske, B., Hunzelman, J., Luz, A., Kovacs, C. S., Lee, K., Pirro, A., Kronenberg, H. M., and Juppner, H. (1997). Targeted expression of constitutively active receptors for parathyroid hormone and parathyroid hormone-related peptide delays endochondral bone formation and rescues mice that lack parathyroid hormone-related peptide. Proc. Natl Acad. Sci.USA 94, 13689–13694. Weir, E. C., Philbrick, W. M., Amling, M., Neff, L. A., Baron, R., and Broadus, A. E. (1996). Targeted overexpression of parathyroid hormonerelated peptide in chondrocytes causes chondrodysplasia and delayed endochondral bone formation. Proc. Natl Acad. Sci. USA 93, 10240–10245. Lanske, B., Karaplis, A. C., Lee, K., Luz, A., Vortkamp, A., Pirro, A., Karperien, M., Defize, L. H. K., Ho, C., Mulligan, R. C.,

246

105.

106.

107.

108.

109.

110.

111.

112.

113.

114.

115.

116.

117.

118.

119.

120.

121.

JANE B. LIAN AND GARY S. STEIN Abou-Samra, A. B., Juppner, H., Segre, G. V., and Kronenberg, H. M. (1996). PTH/PTHrP receptor in early development and Indian hedgehog-regulated bone growth. Science 273, 663–666. Long, F., Chung, U. I., Ohba, S., McMahon, J., Kronenberg, H. M., and McMahon, A. P. (2004). Ihh signaling is directly required for the osteoblast lineage in the endochondral skeleton. Development 131, 1309–1318. St Jacques, B., Hammerschmidt, M., and McMahon, A. P. (1999). Indian hedgehog signaling regulates proliferation and differentiation of chondrocytes and is essential for bone formation. Genes Dev. 13, 2072–2086. Kobayashi, T., Chung, U. I., Schipani, E., Starbuck, M., Karsenty, G., Katagiri, T., Goad, D. L., Lanske, B., and Kronenberg, H. M. (2002). PTHrP and Indian hedgehog control differentiation of growth plate chondrocytes at multiple steps. Development 129, 2977–2986. Schipani, E., and Provot, S. (2003). PTHrP, PTH, and the PTH/PTHrP receptor in endochondral bone development. Birth Defects Res. C. Embryo. Today 69, 352–362. Robson, H., Siebler, T., Stevens, D. A., Shalet, S. M., and Williams, G. R. (2000). Thyroid hormone acts directly on growth plate chondrocytes to promote hypertrophic differentiation and inhibit clonal expansion and cell proliferation. Endocrinology 141, 3887–3897. O’Shea, P. J., Harvey, C. B., Suzuki, H., Kaneshige, M., Kaneshige, K., Cheng, S. Y., and Williams, G. R. (2003). A thyrotoxic skeletal phenotype of advanced bone formation in mice with resistance to thyroid hormone. Mol. Endocrinol. 17, 1410–1424. Li, T. F., Zuscik, M. J., Ionescu, A. M., Zhang, X., Rosier, R. N., Schwarz, E. M., Drissi, H., and O’Keefe, R. J. (2004). PGE2 inhibits chondrocyte differentiation through PKA and PKC signaling. Exp. Cell Res. 300, 159–169. Vermot, J., Niederreither, K., Garnier, J. M., Chambon, P., and Dolle, P. (2003). Decreased embryonic retinoic acid synthesis results in a DiGeorge syndrome phenotype in newborn mice. Proc. Natl Acad. Sci. USA 100, 1763–1768. Niederreither, K., Vermot, J., Schuhbaur, B., Chambon, P., and Dolle, P. (2002). Embryonic retinoic acid synthesis is required for forelimb growth and anteroposterior patterning in the mouse. Development 129, 3563–3574. Allen, S. P., Maden, M., and Price, J. S. (2002). A role for retinoic acid in regulating the regeneration of deer antlers. Dev. Biol. 251, 409–423. Weston, A. D., Chandraratna, R. A., Torchia, J., and Underhill, T. M. (2002). Requirement for RAR-mediated gene repression in skeletal progenitor differentiation. J. Cell Biol. 158, 39–51. Tseng, K. F., and Goldstein, S. A. (1998). Systemic over-secretion of growth hormone in transgenic mice results in a specific pattern of skeletal modeling and adaptation. J. Bone Miner. Res. 13, 706–715. Tribioli, C., and Lufkin, T. (1999). The murine Bapx1 homeobox gene plays a critical role in embryonic development of the axial skeleton and spleen. Development 126, 5699–5711. Akazawa, H., Komuro, I., Sugitani, Y., Yazaki, Y., Nagai, R., and Noda, T. (2000). Targeted disruption of the homeobox transcription factor Bapx1 results in lethal skeletal dysplasia with asplenia and gastroduodenal malformation. Genes Cells 5, 499–513. Lengner, C. J., Drissi, H., Choi, J.-Y., van Wijnen, A. J., Stein, J. L., Stein, G. S., and Lian, J. B. (2002). Activation of the bone related Runx2/Cbfa1 promoter in mesenchymal condensations and developing chondrocytes of the axial skeleton. Mech. Dev. 114, 167–170. Zeng, L., Kempf, H., Murtaugh, L. C., Sato, M. E., and Lassar, A. B. (2002). Shh establishes an Nkx3.2/Sox9 autoregulatory loop that is maintained by BMP signals to induce somitic chondrogenesis. Genes Dev. 16, 1990–2005. Murtaugh, L. C., Zeng, L., Chyung, J. H., and Lassar, A. B. (2001). The chick transcriptional repressor Nkx3.2 acts downstream of Shh to promote BMP-dependent axial chondrogenesis. Dev. Cell 1, 411–422.

122. Wilson, J., and Tucker, A. S. (2004). Fgf and Bmp signals repress the expression of Bapx1 in the mandibular mesenchyme and control the position of the developing jaw joint. Dev. Biol. 266, 138–150. 123. Lettice, L. A., Purdie, L. A., Carlson, G. J., Kilanowski, F., Dorin, J., and Hill, R. E. (1999). The mouse bagpipe gene controls development of axial skeleton, skull, and spleen. Proc. Natl Acad. Sci. USA 96, 9695–9700. 124. Akiyama, H., Chaboissier, M. C., Martin, J. F., Schedl, A., and de Crombrugghe, B. (2002). The transcription factor Sox9 has essential roles in successive steps of the chondrocyte differentiation pathway and is required for expression of Sox5 and Sox6. Genes Dev. 16, 2813–2828. 125. Lengner, C. J., Hassan, M. Q., Serra, R. W., Lepper, C., van Wijnen, A. J., Stein, J. L., Lian, J. B., and Stein, G. S. (2005). Nkx3.2 mediated repression of RUNX2 promotes chondrogenic differentiation. J. Biol. Chem. 280, 15872–15879. 126. Eames, B. F., Sharpe, P. T., and Helms, J. A. (2004). Hierarchy revealed in the specification of three skeletal fates by Sox9 and Runx2. Dev. Biol. 274, 188–200. 127. Jimenez, M. J., Balbin, M., Alvarez, J., Komori, T., Bianco, P., Holmbeck, K., Birkedal-Hansen, H., Lopez, J. M., and Lopez-Otin, C. (2001). A regulatory cascade involving retinoic acid, Cbfa1, and matrix metalloproteinases is coupled to the development of a process of perichondrial invasion and osteogenic differentiation during bone formation. J. Cell Biol. 155, 1333–1344. 128. Zelzer, E., McLean, W., Ng, Y. S., Fukai, N., Reginato, A. M., Lovejoy, S., d’Amore, P. A., and Olsen, B. R. (2002). Skeletal defects in VEGF(120/120) mice reveal multiple roles for VEGF in skeletogenesis. Development 129, 1893–1904. 129. Pratap, J., Vashi, S., Zhang, J., Languino, L., van Wijnen, A. J., Stein, J. L., Lian, J. B., and Stein, G. S. (2004). Runx2 regulates transcription of gelatinases (MMP9) in metastatic cancer cell lines and functionally related to cell migration. J. Bone Miner. Res. 19[Suppl 1], S123. (Abstract). 130. Stricker, S., Fundele, R., Vortkamp, A., and Mundlos, S. (2002). Role of runx genes in chondrocyte differentiation. Dev. Biol. 245, 95–108. 131. Iwamoto, M., Kitagaki, J., Tamamura, Y., Gentili, C., Koyama, E., Enomoto, H., Komori, T., Pacifici, M., and Enomoto-Iwamoto, M. (2003). Runx2 expression and action in chondrocytes are regulated by retinoid signaling and parathyroid hormone-related peptide (PTHrP). Osteoarthritis Cartilage 11, 6–15. 132. Takeda, S., Bonnamy, J. P., Owen, M. J., Ducy, P., and Karsenty, G. (2001). Continuous expression of Cbfa1 in nonhypertrophic chondrocytes uncovers its ability to induce hypertrophic chondrocyte differentiation and partially rescues Cbfa1-deficient mice. Genes Dev. 15, 467–481. 133. Inada, M., Yasui, T., Nomura, S., Miyake, S., Deguchi, K., Himeno, M., Sato, M., Yamagiwa, H., Kimura, T., Yasui, N., Ochi, T., Endo, N., Kitamura, Y., Kishimoto, T., and Komori, T. (1999). Maturational disturbance of chondrocytes in Cbfa1-deficient mice. Dev. Dyn. 214, 279–290. 134. Kim, I. S., Otto, F., Zabel, B., and Mundlos, S. (1999). Regulation of chondrocyte differentiation by Cbfa1. Mech. Dev. 80, 159–170. 135. Kundu, M., Javed, A., Jeon, J. P., Horner, A., Shum, L., Eckhaus, M., Muenke, M., Lian, J. B., Yang, Y., Nuckolls, G. H., Stein, G. S., and Liu, P. P. (2002). Cbfbeta interacts with Runx2 and has a critical role in bone development. Nat. Genet. 32, 639–644. 136. Miller, J., Horner, A., Stacy, T., Lowry, C., Lian, J. B., Stein, G., Nuckolls, G. H., and Speck, N. A. (2002). The core binding factor beta subunit is required for bone formation and hematopoietic maturation. Nat Genet. 32, 645–649. 137. Takamoto, M., Tsuji, K., Yamashita, T., Sasaki, H., Yano, T., Taketani, Y., Komori, T., Nifuji, A., and Noda, M. (2003). Hedgehog signaling enhances core-binding factor a1 and receptor activator of nuclear factor-kappaB ligand (RANKL) gene expression in chondrocytes. J. Endocrinol. 177, 413–421.

Chapter 14 The Cells of Bone 138. Komori, T. (2003). Requisite roles of Runx2 and Cbfb in skeletal development. J. Bone Miner. Metab. 21, 193–197. 139. Ginis, I., Luo, Y., Miura, T., Thies, S., Brandenberger, R., GerechtNir, S., Amit, M., Hoke, A., Carpenter, M. K., Itskovitz-Eldor, J., and Rao, M. S. (2004). Differences between human and mouse embryonic stem cells. Dev. Biol. 269, 360–380. 140. Tanaka, H., Murphy, C. L., Murphy, C., Kimura, M., Kawai, S., and Polak, J. M. (2004). Chondrogenic differentiation of murine embryonic stem cells: effects of culture conditions and dexamethasone. J. Cell Biochem. 93, 454–462. 141. Sottile, V., Thomson, A., and McWhir, J. (2003). In vitro osteogenic differentiation of human ES cells. Cloning Stem Cells 5, 149–155. 142. zur Nieden, N. I., Kempka, G., and Ahr, H. J. (2003). In vitro differentiation of embryonic stem cells into mineralized osteoblasts. Differentiation 71, 18–27. 143. Bourne, S., Polak, J. M., Hughes, S. P., and Buttery, L. D. (2004). Osteogenic differentiation of mouse embryonic stem cells: differential gene expression analysis by cDNA microarray and purification of osteoblasts by cadherin-11 magnetically activated cell sorting. Tissue Eng. 10, 796–806. 144. Kramer, J., Hegert, C., and Rohwedel, J. (2003). In vitro differentiation of mouse ES cells: bone and cartilage. Methods Enzymol 365, 251–268. 145. Bronson, S. K. (2003). Bone nodule formation via in vitro differentiation of murine embryonic stem cells. Methods Enzymol 365, 241–251. 146. Tsuneto, M., Yamane, T., Okuyama, H., Yamazaki, H., and Hayashi, S. (2003). In vitro differentiation of mouse ES cells into hematopoietic, endothelial, and osteoblastic cell lineages: the possibility of in vitro organogenesis. Methods Enzymol 365, 98–114. 147. Petersen, B. E., Bowen, W. C., Patrene, K. D., Mars, W. M., Sullivan, A. K., Murase, N., Boggs, S. S., Greenberger, J. S., and Goff, J. P. (1999). Bone marrow as a potential source of hepatic oval cells. Science 284, 1168–1170. 148. Makino, S., Fukuda, K., Miyoshi, S., Konishi, F., Kodama, H., Pan, J., Sano, M., Takahashi, T., Hori, S., Abe, H., Hata, J., Umezawa, A., and Ogawa, S. (1999). Cardiomyocytes can be generated from marrow stromal cells in vitro. J. Clin. Invest. 103, 697–705. 149. Jiang, Y., Jahagirdar, B. N., Reinhardt, R. L., Schwartz, R. E., Keene, C. D., Ortiz-Gonzalez, X. R., Reyes, M., Lenvik, T., Lund, T., Blackstad, M., Du, J., Aldrich, S., Lisberg, A., Low, W. C., Largaespada, D. A., and Verfaillie, C. M. (2002). Pluripotency of mesenchymal stem cells derived from adult marrow. Nature 418, 41–49. 150. Olmsted-Davis, E. A., Gugala, Z., Camargo, F., Gannon, F. H., Jackson, K., Kienstra, K. A., Shine, H. D., Lindsey, R. W., Hirschi, K. K., Goodell, M. A., Brenner, M. K., and Davis, A. R. (2003). Primitive adult hematopoietic stem cells can function as osteoblast precursors. Proc. Natl Acad Sci. USA 100, 15877–15882. 151. Grove, J. E., Bruscia, E., and Krause, D. S. (2004). Plasticity of bone marrow-derived stem cells. Stem Cells 22, 487–500. 152. Dunbar, C. E., Tisdale, J., Yu, J. M., Soma, T., Zujewski, J., Bodine, D., Sellers, S., Cowan, K., Donahue, R., and Emmons, R. (1997). Transduction of hematopoietic stem cells in humans and in nonhuman primates. Stem Cells 15, 135–139 (Review). 153. Bruder, S. P., Ricalton, N. S., Boynton, R. E., Connolly, T. J., Jaiswal, N., Zaia, J., and Barry, F. P. (1998). Mesenchymal stem cell surface antigen SB-10 corresponds to activated leukocyte cell adhesion molecule and is involved in osteogenic differentiation. J. Bone Miner. Res. 13, 655–663. 154. Gronthos, S., Graves, S. E., Ohta, S., and Simmons, P. J. (1994). The STRO-1+ fraction of adult human bone marrow contains the osteogenic precursors. Blood 84, 4164–4173. 155. Haynesworth, S. E., Baber, M. A., and Caplan, A. I. (1992). Cell surface antigens on human marrow-derived mesenchymal cells are detected by monoclonal antibodies. Bone 13, 69–80.

247 156. Joyner, C. J., Bennett, A., and Triffitt, J. T. (1997). Identification and enrichment of human osteoprogenitor cells by using differentiation stage-specific monoclonal antibodies. Bone 21, 1–6. 157. Turksen, K., and Aubin, J. E. (1991). Positive and negative immunoselection for enrichment of two classes of osteoprogenitor cells. J. Cell Biol. 114, 373–384. 158. Otto, W. R., and Rao, J. (2004). Tomorrow’s skeleton staff: mesenchymal stem cells and the repair of bone and cartilage. Cell Prolif. 37, 97–110. 159. Chambers, I., and Smith, A. (2004). Self-renewal of teratocarcinoma and embryonic stem cells. Oncogene 23, 7150–7160. 160. Heng, B. C., Cao, T., Stanton, L. W., Robson, P., and Olsen, B. (2004). Strategies for directing the differentiation of stem cells into the osteogenic lineage in vitro. J. Bone Miner. Res. 19, 1379–1394. 161. Friedenstein, A. J., Chailakhjan, R. K., and Lalykina, K. S. (1970). The development of fibroblast colonies in monolayer cultures of guinea-pig bone marrow and spleen cells. Cell Tissue Kinet. 3, 393–403. 162. Friedenstein, A. J., Chailakhyan, R. K., and Gerasimov, U. V. (1987). Bone marrow osteogenic stem cells: in vitro cultivation and transplantation in diffusion chambers. Cell Tissue Kinet. 20, 263–272. 163. Friedenstein, A. J., Latzinik, N. V., Gorskaya Yu. F., Luria, E. A., and Moskvina, I. L. (1992). Bone marrow stromal colony formation requires stimulation by haemopoietic cells. Bone Miner. 18, 199–213. 164. Horowitz, M. C., and Lorenzo, J. A. (2004). The origins of osteoclasts. Curr. Opin. Rheumatol. 16, 464–468. 165. Malaval, L., and Aubin, J. E. (2001). Biphasic effects of leukemia inhibitory factor on osteoblastic differentiation. J. Cell Biochem. 81, 63–70. 166. Metcalf, D. (2003). The unsolved enigmas of leukemia inhibitory factor. Stem Cells 21, 5–14. 167. Owen, M. E., Cave, J., and Joyner, C. J. (1987). Clonal analysis in vitro of osteogenic differentiation of marrow CFU-F. J. Cell Sci. 87, 731–738. 168. Chaudhary, L. R., and Hruska, K. A. (2001). The cell survival signal Akt is differentially activated by PDGF-BB, EGF, and FGF-2 in osteoblastic cells. J. Cell Biochem. 81, 304–311. 169. Nakamura, T., Hanada, K., Tamura, M., Shibanushi, T., Nigi, H., Tagawa, M., Fukumoto, S., and Matsumoto, T. (1995). Stimulation of endosteal bone formation by systemic injections of recombinant basic fibroblast growth factor in rats. Endocrinology 136, 1276–1284. 170. Noda, M., and Camilliere, J. J. (1989). In vivo stimulation of bone formation by transforming growth factor-beta. Endocrinology 124, 2991–2994. 171. Hock, J. M., Canalis, E., and Centrella, M. (1990). Transforming growth factor-beta stimulates bone matrix apposition and bone cell replication in cultured fetal rat calvariae. Endocrinology 126, 421–426. 172. Hauschka, P. V., Mavrakos, A. E., Iafrati, M. D., Doleman, S. E., and Klagsbrun, M. (1986). Growth factors in bone matrix. J. Biol. Chem. 261, 12665–12674. 173. Long, M. W., Robinson, J. A., Ashcraft, E. A., and Mann, K. G. (1995). Regulation of human bone marrow-derived osteoprogenitor cells by osteogenic growth factors. J. Clin. Invest. 95, 881–887. 174. Gazzerro, E., Du, Z., Devlin, R. D., Rydziel, S., Priest, L., Economides, A. N., and Canalis, E. (2003). Noggin arrests stromal cell differentiation in vitro. Bone 32, 111–119. 175. Yoshida, Y., Tanaka, S., Umemori, H., Minowa, O., Usui, M., Ikematsu, N., Hosoda, E., Imamura, T., Kuno, J., Yamashita, T., Miyazono, K., Noda, M., Noda, T., and Yamamoto, T. (2000). Negative regulation of BMP/Smad signaling by Tob in osteoblasts. Cell 103, 1085–1097. 176. Yoshida, Y., von Bubnoff, A., Ikematsu, N., Blitz, I. L., Tsuzuku, J. K., Yoshida, E. H., Umemori, H., Miyazono, K., Yamamoto, T., and

248

177.

178.

179.

180.

181.

182.

183.

184.

185. 186.

187.

188.

189.

190.

191.

192.

JANE B. LIAN AND GARY S. STEIN Cho, K. W. (2003). Tob proteins enhance inhibitory Smad-receptor interactions to repress BMP signaling. Mech. Dev. 120, 629–637. Thies, R. S., Bauduy, M., Ashton, B. A., Kurtzberg, L., Wozney, J. M., and Rosen, V. (1992). Recombinant human bone morphogenetic protein-2 induces osteoblastic differentiation in W-20-17 stromal cells. Endocrinology 130, 1318–1324. Wang, E. A., Israel, D. I., Kelly, S., and Luxenberg, D. P. (1993). Bone morphogenetic protein-2 causes commitment and differentiation in C3H10T1/2 and 3T3 cells. Growth Factors 9, 57–71. Katagiri, T., Yamaguchi, A., Komaki, M., Abe, E., Takahashi, N., Ikeda, T., Rosen, V., Wozney, J. M., Fujisawa-Sehara, A., and Suda, T. (1994). Bone morphogenetic protein-2 converts the differentiation pathway of C2C12 myoblasts into the osteoblast lineage. J. Cell Biol. 127, 1755–1766. Einhorn, T. A., Majeska, R. J., Mohaideen, A., Kagel, E. M., Bouxsein, M. L., Turek, T. J., and Wozney, J. M. (2003). A single percutaneous injection of recombinant human bone morphogenetic protein-2 accelerates fracture repair. J. Bone Joint Surg. Am. 85-A, 1425–1435. Hock, J. M., Fonseca, J., Gunness-Hey, M., Kemp, B. E., and Martin, T. J. (1989). Comparison of the anabolic effects of synthetic parathyroid hormone-related protein (PTHrP) 1–34 and PTH 1–34 on bone in rats. Endocrinology 125, 2022–2027. Shalhoub, V., Aslam, F., Breen, E., van Wijnen, A., Bortell, R., Stein, G. S., Stein, J. L., and Lian, J. B. (1998). Multiple levels of steroid hormone-dependent control of osteocalcin during osteoblast differentiation: glucocorticoid regulation of basal and vitamin D stimulated gene expression. J. Cell Biochem. 69, 154–168. Delany, A. M., Durant, D., and Canalis, E. (2001). Glucocorticoid suppression of IGF I transcription in osteoblasts. Mol. Endocrinol. 15, 1781–1789. McCarthy, T. L., Ji, C., Chen, Y., Kim, K., and Centrella, M. (2000). Time- and dose-related interactions between glucocorticoid and cyclic adenosine 3′,5′-monophosphate on CCAAT/enhancer-binding proteindependent insulin-like growth factor I expression by osteoblasts. Endocrinology 141, 127–137. Rosen, C. J. (2003). The cellular and clinical parameters of anabolic therapy for osteoporosis. Crit. Rev. Eukaryot. Gene Expr. 13, 25–38. Qin, L., Tamasi, J., Raggatt, L., Li, X., Feyen, J. H., Lee, D. C., Dicicco-Bloom, E., and Partridge, N. C. (2005). Amphiregulin is a novel growth factor involved in normal bone development and in the cellular response to parathyroid hormone stimulation. J. Biol. Chem. 280, 3974–3981. Qin, L., Li, X., Ko, J. K., and Partridge, N. C. (2005). Parathyroid hormone uses multiple mechanisms to arrest the cell cycle progression of osteoblastic cells from G1 to S phase. J. Biol. Chem. 280, 3104–3111. Gensure, R. C., Gardella, T. J., and Juppner, H. (2005). Parathyroid hormone and parathyroid hormone-related peptide, and their receptors. Biochem. Biophys. Res. Commun. 328, 666–678. Karaplis, A. C., Luz, A., Glowacki, J., Bronson, R. T., Tybulewicz, V. L., Kronenberg, H. M., and Mulligan, R. C. (1994). Lethal skeletal dysplasia from targeted disruption of the parathyroid hormone-related peptide gene. Genes Dev. 8, 277–289. Miao, D., Li, J., Xue, Y., Su, H., Karaplis, A. C., and Goltzman, D. (2004). Parathyroid hormone-related peptide is required for increased trabecular bone volume in parathyroid hormone-null mice. Endocrinology 145, 3554–3562. Liu, F., Malaval, L., and Aubin, J. E. (2003). Global amplification polymerase chain reaction reveals novel transitional stages during osteoprogenitor differentiation. J. Cell Sci. 116, 1787–1796. Watanabe, N., Tezuka, Y., Matsuno, K., Miyatani, S., Morimura, N., Yasuda, M., Fujimaki, R., Kuroda, K., Hiraki, Y., Hozumi, N., and Tezuka, K. (2003). Suppression of differentiation and proliferation of early chondrogenic cells by Notch. J. Bone Miner. Metab. 21, 344–352.

193. Tezuka, K., Yasuda, M., Watanabe, N., Morimura, N., Kuroda, K., Miyatani, S., and Hozumi, N. (2002). Stimulation of osteoblastic cell differentiation by Notch. J. Bone Miner. Res. 17, 231–239. 194. Sowa, H., Kaji, H., Hendy, G. N., Canaff, L., Komori, T., Sugimoto, T., and Chihara, K. (2004). Menin is required for bone morphogenetic protein 2- and transforming growth factor beta-regulated osteoblastic differentiation through interaction with Smads and Runx2. J. Biol. Chem. 279, 40267–40275. 195. Chan, G. K., Miao, D., Deckelbaum, R., Bolivar, I., Karaplis, A., and Goltzman, D. (2003). Parathyroid hormone-related peptide interacts with bone morphogenetic protein 2 to increase osteoblastogenesis and decrease adipogenesis in pluripotent C3H10T 1/2 mesenchymal cells. Endocrinology 144, 5511–5520. 196. Settleman, J. (2004). Tension precedes commitment-even for a stem cell. Mol. Cell 14, 148–150. 197. McBeath, R., Pirone, D. M., Nelson, C. M., Bhadriraju, K., and Chen, C. S. (2004). Cell shape, cytoskeletal tension, and RhoA regulate stem cell lineage commitment. Dev. Cell 6, 483–495. 198. Zayzafoon, M., Gathings, W. E., and McDonald, J. M. (2004). Modeled microgravity inhibits osteogenic differentiation of human mesenchymal stem cells and increases adipogenesis. Endocrinology 145, 2421–2432. 199. Ontiveros, C., Irwin, R., Wiseman, R. W., and McCabe, L. R. (2004). Hypoxia suppresses runx2 independent of modeled microgravity. J. Cell Physiol. 200, 169–176. 200. Tang, Q. Q., Otto, T. C., and Lane, M. D. (2003). CCAAT/enhancerbinding protein beta is required for mitotic clonal expansion during adipogenesis. Proc. Natl Acad. Sci.USA 100, 850–855. 201. Mueller, E., Drori, S., Aiyer, A., Yie, J., Sarraf, P., Chen, H., Hauser, S., Rosen, E. D., Ge, K., Roeder, R. G., and Spiegelman, B. M. (2002). Genetic analysis of adipogenesis through peroxisome proliferatoractivated receptor gamma isoforms. J. Biol. Chem. 277, 41925–41930. 202. de Crombrugghe, B., Lefebvre, V., Behringer, R. R., Bi, W., Murakami, S., and Huang, W. (2000). Transcriptional mechanisms of chondrocyte differentiation. Matrix Biol. 19, 389–394. 203. Skillington, J., Choy, L., and Derynck, R. (2002). Bone morphogenetic protein and retinoic acid signaling cooperate to induce osteoblast differentiation of preadipocytes. J. Cell Biol. 159, 135–146. 204. Shi, X., Shi, W., Li, Q., Song, B., Wan, M., Bai, S., and Cao, X. (2003). A glucocorticoid-induced leucine-zipper protein, GILZ, inhibits adipogenesis of mesenchymal cells. EMBO Rep. 4, 374–380. 205. Lee, M. H., Javed, A., Kim, H. J., Shin, H. I., Gutierrez, S., Choi, J. Y., Rosen, V., Stein, J. L., van Wijnen, A. J., Stein, G. S., Lian, J. B., and Ryoo, H. M. (1999). Transient upregulation of CBFA1 in response to bone morphogenetic protein-2 and transforming growth factor β1 in C2C12 myogenic cells coincides with suppression of the myogenic phenotype but is not sufficient for osteoblast differentiation. J. Cell Biochem. 73, 114–125. 206. Gori, F., Thomas, T., Hicok, K. C., Spelsberg, T. C., and Riggs, B. L. (1999). Differentiation of human marrow stromal precursor cells: bone morphogenetic protein-2 increases OSF2/CBFA1, enhances osteoblast commitment, and inhibits late adipocyte maturation. J. Bone Miner. Res. 14, 1522–1535. 207. Jeon, M. J., Kim, J. A., Kwon, S. H., Kim, S. W., Park, K. S., Park, S. W., Kim, S. Y., and Shin, C. S. (2003). Activation of peroxisome proliferator-activated receptor-gamma inhibits the Runx2-mediated transcription of osteocalcin in osteoblasts. J. Biol. Chem. 278, 23270–23277. 208. Akune, T., Ohba, S., Kamekura, S., Yamaguchi, M., Chung, U. I., Kubota, N., Terauchi, Y., Harada, Y., Azuma, Y., Nakamura, K., Kadowaki, T., and Kawaguchi, H. (2004). PPARgamma insufficiency enhances osteogenesis through osteoblast formation from bone marrow progenitors. J. Clin. Invest. 113, 846–855. 209. Young, D. W., Zaidi, S. K., Furcinitti, P. S., Javed, A., van Wijnen, A. J., Stein, J. L., Lian, J. B., and Stein, G. S. (2004). Quantitative signature

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211.

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213. 214.

215.

216.

217.

218.

219.

220.

221.

222.

223.

224.

225.

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for architectural organization of regulatory factors using intranuclear informatics. J. Cell Sci. 117, 4889–4896. Kobayashi, H., Gao, Y., Ueta, C., Yamaguchi, A., and Komori, T. (2000). Multilineage differentiation of Cbfa1-deficient calvarial cells in vitro. Biochem. Biophys. Res. Commun. 273, 630–636. Pratap, J., Galindo, M., Zaidi, S. K., Vradii, D., Bhat, B. M., Robinson, J. A., Choi, J.-Y., Komori, T., Stein, J. L., Lian, J. B., Stein, G. S., and van Wijnen, A. J. (2003). Cell growth regulatory role of Runx2 during proliferative expansion of pre-osteoblasts. Cancer Res. 63, 5357–5362. Stein, G. S., Lian, J. B., Stein, J. L., van Wijnen, A. J., and Montecino, M. (1996). Transcriptional control of osteoblast growth and differentiation. Physiol. Rev. 76, 593–629. Hughes, D. E., and Boyce, B. F. (1997). Apoptosis in bone physiology and disease. Mol. Pathol. 50, 132–137. Noble, B. S., Stevens, H., Loveridge, N., and Reeve, J. (1997). Identification of apoptotic changes in osteocytes in normal and pathological human bone. Bone 20, 273–282. Parfitt, A. M., Villanueva, A. R., Foldes, J., and Rao, D. S. (1995). Relations between histologic indices of bone formation: implications for the pathogenesis of spinal osteoporosis. J. Bone Miner. Res. 10, 466–473. Lynch, M., Capparelli, C., Stein, J. L., Stein, G. S., and Lian, J. B. (1998). Apoptosis during bone-like tissue development in vitro. J. Cell Biochem. 68, 31–49. Tomkinson, A., Reeve, J., Shaw, R. W., and Noble, B. S. (1997). The death of osteocytes via apoptosis accompanies estrogen withdrawal in human bone. J. Clin. Endocrinol. Metab. 82, 3128–3135. Lynch, M. P., Stein, J. L., Stein, G. S., and Lian, J. B. (1995). The influence of type I collagen on the development and maintenance of the osteoblast phenotype in primary and passaged rat calvarial osteoblasts: modification of expression of genes supporting cell growth, adhesion, and extracellular matrix mineralization. Exp. Cell Res. 216, 35–45. Vukicevic, S., Luyten, F. P., Kleinman, H. K., and Reddi, A. H. (1990). Differentiation of canalicular cell processes in bone cells by basement membrane matrix components: regulation by discrete domains of laminin. Cell 63, 437–445. Andrianarivo, A. G., Robinson, J. A., Mann, K. G., and Tracy, R. P. (1992). Growth on type I collagen promotes expression of the osteoblastic phenotype in human osteosarcoma MG-63 cells. J. Cell Physiol. 153, 256–265. Franceschi, R. T., and Iyer, B. S. (1992). Relationship between collagen synthesis and expression of the osteoblast phenotype in MC3T3-E1 cells. J. Bone Miner. Res. 7, 235–246. Robey, P. G. (2002). Bone matrix proteoglycans and glycoproteins. In “Principles of Bone Biology” (J. P. Bilezikian, L. G. Raisz, and G. A. Rodan, eds.), pp. 225–237. Academic Press, San Diego, CA. Gronthos, S., Stewart, K., Graves, S. E., Hay, S., and Simmons, P. J. (1997). Integrin expression and function on human osteoblast-like cells. J. Bone Miner. Res. 12, 1189–1197. Xiao, G., Gopalkrishnan, R. V., Jiang, D., Reith, E., Benson, M. D., and Franceschi, R. (2002). Bone morphogenetic proteins, extracellular matrix, and mitogen-activated protein kinase signaling pathways are required for osteoblast-specific gene expression and differentiation in MC3T3-E1 cells. J. Bone Miner. Res. 17, 101–110. Cheng, S. L., Lecanda, F., Davidson, M. K., Warlow, P. M., Zhang, S. F., Zhang, L., Suzuki, S., St. John, T., and Civitelli, R. (1998). Human osteoblasts express a repertoire of cadherins, which are critical for BMP-2-induced osteogenic differentiation. J. Bone Miner. Res. 13, 633–644. Lee, Y. S., and Chuong, C. M. (1992). Adhesion molecules in skeletogenesis: I. Transient expression of neural cell adhesion molecules (NCAM) in osteoblasts during endochondral and intramembranous ossification. J. Bone Miner. Res. 7, 1435–1446.

249 227. Okazaki, M., Takeshita, S., Kawai, S., Kikuno, R., Tsujimura, A., Kudo, A., and Amann, E. (1994). Molecular cloning and characterization of OB-cadherin, a new member of cadherin family expressed in osteoblasts. J. Biol. Chem. 269, 12092–12098. 228. Arai, F., Ohneda, O., Miyamoto, T., Zhang, X. Q., and Suda, T. (2002). Mesenchymal stem cells in perichondrium express activated leukocyte cell adhesion molecule and participate in bone marrow formation. J. Exp. Med. 195, 1549–1563. 229. Jamal, H. H., and Aubin, J. E. (1996). CD44 expression in fetal rat bone: in vivo and in vitro analysis. Exp. Cell Res. 223, 467–477. 230. Hughes, D. E., Salter, D. M., and Simpson, R. (1994). CD44 expression in human bone: a novel marker of osteocytic differentiation. J. Bone Miner. Res. 9, 39–44. 231. Kato, Y., Windle, J. J., Koop, B. A., Mundy, G. R., and Bonewald, L. F. (1997). Establishment of an osteocyte-like cell line, MLO-Y4. J. Bone Miner. Res. 12, 2014–2023. 232. Pockwinse, S. M., Lawrence, J. B., Singer, R. H., Stein, J. L., Lian, J. B., and Stein, G. S. (1993). Gene expression at single cell resolution associated with development of the bone cell phenotype: ultrastructural and in situ hybridization analysis. Bone 14, 347–352. 233. Pockwinse, S. M., Stein, J. L., Lian, J. B., and Stein, G. S. (1995). Developmental stage-specific cellular responses to vitamin D and glucocorticoids during differentiation of the osteoblast phenotype: interrelationship of morphology and gene expression by in situ hybridization. Exp. Cell Res. 216, 244–260. 234. Breen, E. C., Ignotz, R. A., McCabe, L., Stein, J. L., Stein, G. S., and Lian, J. B. (1994). TGF beta alters growth and differentiation related gene expression in proliferating osteoblasts in vitro, preventing development of the mature bone phenotype. J. Cell Physiol. 160, 323–335. 235. Tang, K.-T., Capparelli, C., Stein, J. L., Stein, G. S., Lian, J. B., Huber, A. C., Braverman, L. E., and DeVito, W. J. (1996). Acidic fibroblast growth factor inhibits osteoblast differentiation in vitro: altered expression of collagenase, cell growth-related and mineralizationassociated genes. J. Cell Biochem. 61, 152–166. 236. Debiais, F., Hott, M., Graulet, A. M., and Marie, P. J. (1998). The effects of fibroblast growth factor-2 on human neonatal calvaria osteoblastic cells are differentiation stage specific. J. Bone Miner. Res. 13, 645–654. 237. Gerstenfeld, L. C., Zurakowski, D., Schaffer, J. L., Nichols, D. P., Toma, C. D., Broess, M., Bruder, S. P., and Caplan, A. I. (1996). Variable hormone responsiveness of osteoblast populations isolated at different stages of embryogenesis and its relationship to the osteogenic lineage. Endocrinology 137, 3957–3968. 238. Owen, T. A., Aronow, M. S., Barone, L. M., Bettencourt, B., Stein, G. S., and Lian, J. B. (1991). Pleiotropic effects of vitamin D on osteoblast gene expression are related to the proliferative and differentiated state of the bone cell phenotype: dependency upon basal levels of gene expression, duration of exposure, and bone matrix competency in normal rat osteoblast cultures. Endocrinology 128, 1496–1504. 239. Bodine, P. V. N., Henderson, R. A., Green, J., Aronow, M., Owen, T., Stein, G. S., Lian, J. B., and Komm, B. S. (1998). Estrogen receptorα is developmentally regulated during osteoblast differentiation and contributes to selective responsiveness of gene expression. Endocrinology 139, 2048–2057. 240. Ishizuya, T., Yokose, S., Hori, M., Noda, T., Suda, T., Yoshiki, S., and Yamaguchi, A. (1997). Parathyroid hormone exerts disparate effects on osteoblast differentiation depending on exposure time in rat osteoblastic cells. J. Clin. Invest. 99, 2961–2970. 241. Govoni, K. E., Baylink, D. J., and Mohan, S. (2005). The multifunctional role of insulin-like growth factor binding proteins in bone. Pediatr. Nephrol. 20, 261–268. 242. Delany, A. M., Pash, J. M., and Canalis, E. (1994). Cellular and clinical perspectives on skeletal insulin-like growth factor I. J. Cell Biochem. 55, 328–333.

250 243. Zhang, M., Xuan, S., Bouxsein, M. L., von Stechow, D., Akeno, N., Faugere, M. C., Malluche, H., Zhao, G., Rosen, C. J., Efstratiadis, A., and Clemens, T. L. (2002). Osteoblast-specific knockout of the insulin-like growth factor (IGF) receptor gene reveals an essential role of IGF signaling in bone matrix mineralization. J. Biol. Chem. 277, 44005–44012. 244. Pereira, R. M., Delany, A. M., and Canalis, E. (2001). Cortisol inhibits the differentiation and apoptosis of osteoblasts in culture. Bone 28, 484–490. 245. Van Leeuwen, J. P., van Driel, M., van den Bemd, G. J., and Pols, H. A. (2001). Vitamin D control of osteoblast function and bone extracellular matrix mineralization. Crit. Rev. Eukaryot. Gene Expr. 11, 199–226. 246. Orwoll, E. S. (2001). Androgens: basic biology and clinical implication. Calcif. Tissue Int. 69, 185–188. 247. Kousteni, S., Chen, J. R., Bellido, T., Han, L., Ali, A. A., O’Brien, C. A., Plotkin, L., Fu, Q., Mancino, A. T., Wen, Y., Vertino, A. M., Powers, C. C., Stewart, S. A., Ebert, R., Parfitt, A. M., Weinstein, R. S., Jilka, R. L., and Manolagas, S. C. (2002). Reversal of bone loss in mice by nongenotropic signaling of sex steroids. Science 298, 843–846. 248. Zhou, S., Zilberman, Y., Wassermann, K., Bain, S. D., Sadovsky, Y., and Gazit, D. (2001). Estrogen modulates estrogen receptor alpha and beta expression, osteogenic activity, and apoptosis in mesenchymal stem cells (MSCs) of osteoporotic mice. J. Cell Biochem. Suppl 36, 144–155. 249. Chen, J. R., Plotkin, L. I., Aguirre, J. I., Han, L., Jilka, R. L., Kousteni, S., Bellido, T., and Manolagas, S. C. (2004). Transient versus sustained phosphorylation and nuclear accumulation of ERKs underlie anti-versus Pro-apoptotic effects of estrogens. J. Biol. Chem. 280, 4632–4638. 250. Murphy, E., and Williams, G. R. (2004). The thyroid and the skeleton. Clin. Endocrinol.(Oxf) 61, 285–298. 251. Siebler, T., Robson, H., Shalet, S. M., and Williams, G. R. (2004). Thyroid hormone and the growth plate. Horm. Res. 62 Suppl 3, 116. 252. Kindblom, J. M., Gevers, E. F., Skrtic, S. M., Lindberg, M. K., Gothe, S., Tornell, J., Vennstrom, B., and Ohlsson, C. (2005). Increased adipogenesis in bone marrow but decreased bone mineral density in mice devoid of thyroid hormone receptors. Bone 36, 607–616. 253. Huang, J. C., Sakata, T., Pfleger, L. L., Bencsik, M., Halloran, B. P., Bikle, D. D., and Nissenson, R. A. (2004). PTH differentially regulates expression of RANKL and OPG. J. Bone Miner. Res. 19, 235–244. 254. Chen, C., Koh, A. J., Datta, N. S., Zhang, J., Keller, E. T., Xiao, G., Franceschi, R. T., D’Silva, N. J., and McCauley, L. K. (2004). Impact of the mitogen-activated protein kinase pathway on parathyroid hormone-related protein actions in osteoblasts. J. Biol. Chem. 279, 29121–29129. 255. Boskey, A. L., Wright, T. M., and Blank, R. D. (1999). Collagen and bone strength. J. Bone Miner. Res. 14, 330–335. 256. Glimcher, M. J. (1989). Mechanism of calcification: role of collagen fibrils and collagen-phosphoprotein complexes in vitro and in vivo. Anat. Rec. 224, 139–153. 257. Han, Y., Cowin, S. C., Schaffler, M. B., and Weinbaum, S. (2004). Mechanotransduction and strain amplification in osteocyte cell processes. Proc. Natl Acad. Sci. USA 101, 16689–16694. 258. Jilka, R. L., Weinstein, R. S., Bellido, T., Parfitt, A. M., and Manolagas, S. C. (1998). Osteoblast programmed cell death (apoptosis): modulation by growth factors and cytokines. J. Bone Miner. Res. 13, 793–802. 259. Karsdal, M. A., Larsen, L., Engsig, M. T., Lou, H., Ferreras, M., Lochter, A., Delaisse, J. M., and Foged, N. T. (2002). Matrix metalloproteinase-dependent activation of latent transforming growth factorbeta controls the conversion of osteoblasts into osteocytes by blocking osteoblast apoptosis. J. Biol. Chem. 277, 44061–44067. 260. Jilka, R. L., Weinstein, R. S., Bellido, T., Roberson, P., Parfitt, A. M., and Manolagas, S. C. (1999). Increased bone formation by prevention

JANE B. LIAN AND GARY S. STEIN

261.

262.

263.

264.

265.

266.

267.

268.

269.

270.

271.

272. 273. 274.

275.

of osteoblast apoptosis with parathyroid hormone. J. Clin. Invest. 104, 439–446. Bellido, T., Ali, A. A., Plotkin, L. I., Fu, Q., Gubrij, I., Roberson, P. K., Weinstein, R. S., O’Brien, C. A., Manolagas, S. C., and Jilka, R. L. (2003). Proteasomal degradation of Runx2 shortens parathyroid hormone-induced anti-apoptotic signaling in osteoblasts. A putative explanation for why intermittent administration is needed for bone anabolism. J. Biol. Chem. 278, 50259–50272. Nijweide, P. J., Burger, E. H., and Klein-Nulend, J. (2002). The osteocyte. In “Principles of Bone Biology” (J. P. Bilezikian, L. G. Raisz, and G. A. Rodan, eds.), pp. 93–107. Academic Press, San Diego, CA. Cheng, B., Kato, Y., Zhao, S., Luo, J., Sprague, E., Bonewald, L. F., and Jiang, J. X. (2001). PGE(2) is essential for gap junction-mediated intercellular communication between osteocyte-like MLO-Y4 cells in response to mechanical strain. Endocrinology 142, 3464–3473. Feng, J. Q., Huang, H., Lu, Y., Ye, L., Xie, Y., Tsutsui, T. W., Kunieda, T., Castranio, T., Scott, G., Bonewald, L. B., and Mishina, Y. (2003). The Dentin matrix protein 1 (Dmp1) is specifically expressed in mineralized, but not soft, tissues during development. J. Dent. Res. 82, 776–780. Gluhak-Heinrich, J., Ye, L., Bonewald, L. F., Feng, J. Q., MacDougall, M., Harris, S. E., and Pavlin, D. (2003). Mechanical loading stimulates dentin matrix protein 1 (DMP1) expression in osteocytes in vivo. J. Bone Miner. Res. 18, 807–817. Ye, L., MacDougall, M., Zhang, S., Xie, Y., Zhang, J., Li, Z., Lu, Y., Mishina, Y., and Feng, J. Q. (2004). Deletion of dentin matrix protein-1 leads to a partial failure of maturation of predentin into dentin, hypomineralization, and expanded cavities of pulp and root canal during postnatal tooth development. J. Biol. Chem. 279, 19141–19148. Yang, W., Lu, Y., Kalajzic, I., Guo, D., Harris, M. A., GluhakHeinrich, J., Kotha, S., Bonewald, L. F., Feng, J. Q., Rowe, D. W., Turner, C. H., Robling, A. G., and Harris, S. E. (2005). Dentin matrix protein 1 gene cis-regulation: Use in osteocytes to characterize local responses to mechanical loading in vitro and In vivo. J. Biol. Chem. 280, 20680–20690. Ahuja, S. S., Zhao, S., Bellido, T., Plotkin, L. I., Jimenez, F., and Bonewald, L. F. (2003). CD40 ligand blocks apoptosis induced by tumor necrosis factor alpha, glucocorticoids, and etoposide in osteoblasts and the osteocyte-like cell line murine long bone osteocyte-Y4. Endocrinology 144, 1761–1769. Tami, A. E., Nasser, P., Verborgt, O., Schaffler, M. B., and Knothe Tate, M. L. (2002). The role of interstitial fluid flow in the remodeling response to fatigue loading. J. Bone Miner. Res. 17, 2030–2037. Noble, B. S., Peet, N., Stevens, H. Y., Brabbs, A., Mosley, J. R., Reilly, G. C., Reeve, J., Skerry, T. M., and Lanyon, L. E. (2003). Mechanical loading: biphasic osteocyte survival and targeting of osteoclasts for bone destruction in rat cortical bone. Am. J. Physiol. Cell Physiol. 284, C934-C943. Colopy, S. A., Benz-Dean, J., Barrett, J. G., Sample, S. J., Lu, Y., Danova, N. A., Kalscheur, V. L., Vanderby, R., Jr., Markel, M. D., and Muir, P. (2004). Response of the osteocyte syncytium adjacent to and distant from linear microcracks during adaptation to cyclic fatigue loading. Bone 35, 881–891. Noble, B. (2003). Bone microdamage and cell apoptosis. Eur. Cell Mater. 6, 46–55. Hernandez, C. J., Majeska, R. J., and Schaffler, M. B. (2004). Osteocyte density in woven bone. Bone 35, 1095–1099. Knothe Tate, M. L., Steck, R., Forwood, M. R., and Niederer, P. (2000). In vivo demonstration of load-induced fluid flow in the rat tibia and its potential implications for processes associated with functional adaptation. J. Exp. Biol. 203 Pt 18, 2737–2745. Weinbaum, S., Cowin, S. C., and Zeng, Y. (1994). A model for the excitation of osteocytes by mechanical loading-induced bone fluid shear stresses. J. Biomech. 27, 339–360.

Chapter 14 The Cells of Bone 276. Cullinane, D. M., and Einhorn, T. A. (2002). Biomechanics of bone. In “Principles of Bone Biology” (J. P. Bilezikian, L. G. Raisz, and G. A. Rodan, eds.), pp. 17–32. Academic Press, San Diego, CA. 277. Noonan, K. J., Stevens, J. W., Tammi, R., Tammi, M., Hernandez, J. A., and Midura, R. J. (1996). Spatial distribution of CD44 and hyaluronan in the proximal tibia of the growing rat. J. Orthop. Res. 14, 573–581. 278. Knothe Tate, M. L. (2003). “Whither flows the fluid in bone?” An osteocyte’s perspective. J. Biomech. 36, 1409–1424. 279. Robling, A. G., and Turner, C. H. (2002). Mechanotransduction in bone: genetic effects on mechanosensitivity in mice. Bone 31, 562–569. 280. Skerry, T. M., and Suva, L. J. (2003). Investigation of the regulation of bone mass by mechanical loading: from quantitative cytochemistry to gene array. Cell Biochem. Funct. 21, 223–229. 281. Cherian, P. P., Cheng, B., Gu, S., Sprague, E., Bonewald, L. F., and Jiang, J. X. (2003). Effects of mechanical strain on the function of Gap junctions in osteocytes are mediated through the prostaglandin EP2 receptor. J. Biol. Chem. 278, 43146–43156. 282. Furlan, F., Lecanda, F., Screen, J., and Civitelli, R. (2001). Proliferation, differentiation and apoptosis in connexin43-null osteoblasts. Cell Commun. Adhes. 8, 367–371. 283. Plotkin, L. I., Manolagas, S. C., and Bellido, T. (2002). Transduction of cell survival signals by connexin-43 hemichannels. J. Biol. Chem. 277, 8648–8657. 284. Bakker, A., Klein-Nulend, J., and Burger, E. (2004). Shear stress inhibits while disuse promotes osteocyte apoptosis. Biochem. Biophys. Res. Commun. 320, 1163–1168. 285. Rubin, C. T., and Lanyon, L. E. (1987). Osteoregulatory nature of mechanical stimuli: function as a determinant for adaptive remodeling in bone. J. Orthop. Res. 5, 300–310. 286. Woodiel, F. N., Fall, P. M., and Raisz, L. G. (1996). Anabolic effects of prostaglandins in cultured fetal rat calvariae: structure-activity relations and signal transduction pathway. J. Bone Miner. Res. 11, 1249–1255. 287. Lohmann, C. H., Schwartz, Z., Liu, Y., Li, Z., Simon, B. J., Sylvia, V. L., Dean, D. D., Bonewald, L. F., Donahue, H. J., and Boyan, B. D. (2003). Pulsed electromagnetic fields affect phenotype and connexin 43 protein expression in MLO-Y4 osteocyte-like cells and ROS 17/2.8 osteoblast-like cells. J. Orthop. Res. 21, 326–334. 288. Hitt, A. L., and Luna, E. J. (1994). Membrane interactions with the actin cytoskeleton. Curr. Opin. Cell Biol. 6, 120–130. 289. Wang, N., Butler, J. P., and Ingber, D. E. (1993). Mechanotransduction across the cell surface and through the cytoskeleton. Science 260, 1124–1127. 290. Boutahar, N., Guignandon, A., Vico, L., and Lafage-Proust, M. H. (2004). Mechanical strain on osteoblasts activates autophosphorylation of focal adhesion kinase and proline-rich tyrosine kinase 2 tyrosine sites involved in ERK activation. J. Biol. Chem. 279, 30588–30599. 291. Tanaka, S., Sakai, A., Tanaka, M., Otomo, H., Okimoto, N., Sakata, T., and Nakamura, T. (2004). Skeletal unloading alleviates the anabolic action of intermittent PTH[1–34] in mouse tibia in association with inhibition of PTH-induced increase in c-fos mRNA in bone marrow cells. J. Bone Miner. Res. 19, 1813–1820. 292. Wang, F. S., Wang, C. J., Chen, Y. J., Chang, P. R., Huang, Y. T., Sun, Y. C., Huang, H. C., Yang, Y. J., and Yang, K. D. (2004). Ras induction of superoxide activates ERK-dependent angiogenic transcription factor HIF-1alpha and VEGF-A expression in shock wavestimulated osteoblasts. J. Biol. Chem. 279, 10331–10337. 293. Judex, S., Zhong, N., Squire, M. E., Ye, K., Donahue, L. R., Hadjiargyrou, M., and Rubin, C. T. (2005). Mechanical modulation of molecular signals which regulate anabolic and catabolic activity in bone tissue. J. Cell. Biochem. 94, 982–994. 294. Gross, T. S., Akeno, N., Clemens, T. L., Komarova, S., Srinivasan, S., Weimer, D. A., and Mayorov, S. (2001). Selected contribution: Osteocytes upregulate HIF-1alpha in response to acute disuse and oxygen deprivation. J. Appl. Physiol. 90, 2514–2519.

251 295. Warden, S. J., and Turner, C. H. (2004). Mechanotransduction in the cortical bone is most efficient at loading frequencies of 5–10 Hz. Bone 34, 261–270. 296. el Haj, A. J., Minter, S. L., Rawlinson, S. C., Suswillo, R., and Lanyon, L. E. (1990). Cellular responses to mechanical loading in vitro. J. Bone Miner. Res. 5, 923–932. 297. Skerry, T. M., Bitensky, L., Chayen, J., and Lanyon, L. E. (1989). Early strain-related changes in enzyme activity in osteocytes following bone loading in vivo. J. Bone Miner. Res. 4, 783–788. 298. Lean, J. M., Jagger, C. J., Chambers, T. J., and Chow, J. W. (1995). Increased insulin-like growth factor I mRNA expression in rat osteocytes in response to mechanical stimulation. Am. J. Physiol. 268, E318–E327. 299. Klein-Nulend, J., van der Plas, A., Semeins, C. M., Ajubi, N. E., Frangos, J. A., Nijweide, P. J., and Burger, E. H. (1995). Sensitivity of osteocytes to biomechanical stress in vitro. FASEB J. 9, 441–445. 300. Raisz, L. G. (1999). Prostaglandins and bone: physiology and pathophysiology. Osteoarthritis Cartilage 7, 419–421. 301. Burger, E. H., and Klein-Nulen, J. (1999). Responses of bone cells to biomechanical forces in vitro. Adv. Dent. Res. 13, 93–98. 302. Zaman, G., Pitsillides, A. A., Rawlinson, S. C., Suswillo, R. F., Mosley, J. R., Cheng, M. Z., Platts, L. A., Hukkanen, M., Polak, J. M., and Lanyon, L. E. (1999). Mechanical strain stimulates nitric oxide production by rapid activation of endothelial nitric oxide synthase in osteocytes. J. Bone Miner. Res. 14, 1123–1131. 303. Nomura, S., and Takano-Yamamoto, T. (2000). Molecular events caused by mechanical stress in bone. Matrix Biol. 19, 91–96. 304. Afzal, F., Polak, J., and Buttery, L. (2004). Endothelial nitric oxide synthase in the control of osteoblastic mineralizing activity and bone integrity. J. Pathol. 202, 503–510. 305. Aguirre, J., Buttery, L., O’Shaughnessy, M., Afzal, F., Fernandez, d. M., I, Hukkanen, M., Huang, P., MacIntyre, I., and Polak, J. (2001). Endothelial nitric oxide synthase gene-deficient mice demonstrate marked retardation in postnatal bone formation, reduced bone volume, and defects in osteoblast maturation and activity. Am. J. Pathol. 158, 247–257. 306. van’t Hof, R. J., Armour, K. J., Smith, L. M., Armour, K. E., Wei, X. Q., Liew, F. Y., and Ralston, S. H. (2000). Requirement of the inducible nitric oxide synthase pathway for IL-1-induced osteoclastic bone resorption. Proc. Natl Acad. Sci. USA 97, 7993–7998. 307. van’t Hof, R. J., Macphee, J., Libouban, H., Helfrich, M. H., and Ralston, S. H. (2004). Regulation of bone mass and bone turnover by neuronal nitric oxide synthase. Endocrinology 145, 5068–5074. 308. Gu, Y., Genever, P. G., Skerry, T. M., and Publicover, S. J. (2002). The NMDA type glutamate receptors expressed by primary rat osteoblasts have the same electrophysiological characteristics as neuronal receptors. Calcif. Tissue Int. 70, 194–203. 309. Genever, P. G., and Skerry, T. M. (2001). Regulation of spontaneous glutamate release activity in osteoblastic cells and its role in differentiation and survival: evidence for intrinsic glutamatergic signaling in bone. FASEB J. 15, 1586–1588. 310. Lian, J. B., Stein, G. S., and Aubin, J. E. (2003). Bone formation: maturation and functional activities of osteoblast lineage cells. In “Primer on the Metabolic Bone Dieseases and Disorders of Mineral Metabolism” (M. J. Favus, ed.), pp. 13–28. American Society for Bone and Mineral Research, Washington, D.C. 311. Jabs, E. W. (2001). A TWIST in the fate of human osteoblasts identifies signaling molecules involved in skull development. J. Clin. Invest. 107, 1075–1077. 312. Chen, Z. F., and Behringer, R. R. (1995). Twist is required in head mesenchyme for cranial neural tube morphogenesis. Genes Dev. 9, 686–699. 313. Cserjesi, P., Brown, D., Ligon, K. L., Lyons, G. E., Copeland, N. G., Gilbert, D. J., Jenkins, N. A., and Olson, E. N. (1995). Scleraxis: a basic

252

314.

315.

316.

317.

318.

319.

320.

321.

322.

323.

324.

325.

326.

327.

328.

JANE B. LIAN AND GARY S. STEIN helix-loop-helix protein that prefigures skeletal formation during mouse embryogenesis. Development 121, 1099–1110. Lee, M. S., Lowe, G. N., Strong, D. D., Wergedal, J. E., and Glackin, C. A. (1999). TWIST, a basic helix-loop-helix transcription factor, can regulate the human osteogenic lineage. J. Cell Biochem. 75, 566–577. Bialek, P., Kern, B., Yang, X., Schrock, M., Sosic, D., Hong, N., Wu, H., Yu, K., Ornitz, D. M., Olson, E. N., Justice, M. J., and Karsenty, G. (2004). A twist code determines the onset of osteoblast differentiation. Dev. Cell 6, 423–435. McCabe, L. R., Banerjee, C., Kundu, R., Harrison, R. J., Dobner, P. R., Stein, J. L., Lian, J. B., and Stein, G. S. (1996). Developmental expression and activities of specific fos and jun proteins are functionally related to osteoblast maturation: role of fra-2 and jun D during differentiation. Endocrinology 137, 4398–4408. Agarwal, S. K., Novotny, E. A., Crabtree, J. S., Weitzman, J. B., Yaniv, M., Burns, A. L., Chandrasekharappa, S. C., Collins, F. S., Spiegel, A. M., and Marx, S. J. (2003). Transcription factor JunD, deprived of menin, switches from growth suppressor to growth promoter. Proc. Natl Acad. Sci. USA 100, 10770–10775. Grigoriadis, A. E., Schellander, K., Wang, Z.-Q., and Wagner, E. F. (1993). Osteoblasts are target cells for transformation in c-fos transgenic mice. J. Cell Biol. 122, 685–701. Machwate, M., Jullienne, A., Moukhtar, M., and Marie, P. J. (1995). Temporal variation of c-fos proto-oncogene expression during osteoblast differentiation and osteogenesis in developing bone. J. Cell Biochem. 57, 62–70. McCabe, L. R., Kockx, M., Lian, J., Stein, J., and Stein, G. (1995). Selective expression of fos- and jun-related genes during osteoblast proliferation and differentiation. Exp. Cell Res. 218, 255–262. Lufkin, T., Mark, M., Hart, C. P., Dolle, P., LeMeur, M., and Chambon, P. (1992). Homeotic transformation of the occipital bones of the skull by ectopic expression of a homeobox gene. Nature 359, 835–841. Jabs, E. W., Muller, U., Li, X., Ma, L., Luo, W., Haworth, I. S., Klisak, I., Sparkes, R., Warman, M. L., Mulliken, J. B., Snead, M. L., and Maxson, R. (1993). A mutation in the homeodomain of the human MSX2 gene in a family affected with autosomal dominant craniosynostosis. Cell 75, 443–450. Mina, M., Gluhak, J., Upholt, W. B., Kollar, E. J., and Rogers, B. (1995). Experimental analysis of Msx-1 and Msx-2 gene expression during chick mandibular morphogenesis. Dev. Dynamics 202, 195–214. Zhao, G. Q., Zhao, S., Zhou, X., Eberspaecher, H., Solursh, M., and de Crombrugghe, B. (1994). rDlx, a novel distal-less-like homeoprotein is expressed in developing cartilages and discrete neuronal tissues. Dev. Biol. 164, 37–51. Orestes-Cardoso, S., Nefussi, J. R., Lezot, F., Oboeuf, M., Pereira, M., Mesbah, M., Robert, B., and Berdal, A. (2002). Msx1 is a regulator of bone formation during development and postnatal growth: in vivo investigations in a transgenic mouse model. Connect. Tissue Res. 43, 153–160. Hassan, M. Q., Javed, A., Morasso, M. I., Karlin, J., Montecino, M., van Wijnen, A. J., Stein, G. S., Stein, J. L., and Lian, J. B. (2004). Dlx3 transcriptional regulation of osteoblast differentiation: temporal recruitment of Msx2, Dlx3, and Dlx5 homeodomain proteins to chromatin of the osteocalcin gene. Mol. Cell Biol. 24, 9248–9261. Dodig, M., Kronenberg, M. S., Bedalov, A., Kream, B. E., Gronowicz, G., Clark, S. H., Mack, K., Liu, Y. H., Maxon, R., Pan, Z. Z., Upholt, W. B., Rowe, D. W., and Lichtler, A. C. (1996). Identification of a TAAT-containing motif required for high level expression of the COL1A1 promoter in differentiated osteoblasts of transgenic mice. J. Biol. Chem. 271, 16422–16429. Hoffmann, H. M., Catron, K. M., van Wijnen, A. J., McCabe, L. R., Lian, J. B., Stein, G. S., and Stein, J. L. (1994). Transcriptional control of the tissue-specific, developmentally regulated osteocalcin gene

329.

330.

331.

332.

333.

334.

335.

336.

337.

338.

339.

340.

341.

342.

343.

requires a binding motif for the Msx family of homeodomain proteins. Proc. Natl Acad. Sci. USA 91, 12887–12891. Ryoo, H.-M., Hoffmann, H. M., Beumer, T. L., Frenkel, B., Towler, D. A., Stein, G. S., Stein, J. L., van Wijnen, A. J., and Lian, J. B. (1997). Stage-specific expression of Dlx-5 during osteoblast differentiation: involvement in regulation of osteocalcin gene expression. Mol. Endocrinol. 11, 1681–1694. Towler, D. A., Rutledge, S. J., and Rodan, G. A. (1994). Msx-2/Hox 8.1: a transcriptional regulator of the rat osteocalcin promoter. Mol. Endocrinol. 8, 1484–1493. Hoffmann, H. M., Beumer, T. L., Rahman, S., McCabe, L. R., Banerjee, C., Aslam, F., Tiro, J. A., van Wijnen, A. J., Stein, J. L., Stein, G. S., and Lian, J. B. (1996). Bone tissue-specific transcription of the osteocalcin gene: role of an activator osteoblast-specific complex and suppressor hox proteins that bind the OC box. J. Cell Biochem. 61, 310–324. Bidder, M., Latifi, T., and Towler, D. A. (1998). Reciprocal temporospatial patterns of Msx2 and Osteocalcin gene expression during murine odontogenesis. J. Bone Miner. Res. 13, 609–619. Nimer, S. D., and Moore, M. A. (2004). Effects of the leukemiaassociated AML1-ETO protein on hematopoietic stem and progenitor cells. Oncogene 23, 4249–4254. Lian, J. B., Balint, E., Javed, A., Drissi, H., Vitti, R., Quinlan, E. J., Zhang, L., van Wijnen, A. J., Stein, J. L., Speck, N., and Stein, G. S. (2003). Runx1/AML1 hematopoietic transcription factor contributes to skeletal development in vivo. J. Cell Physiol. 196, 301–311. Levanon, D., Bettoun, D., Harris-Cerruti, C., Woolf, E., Negreanu, V., Eilam, R., Bernstein, Y., Goldenberg, D., Xiao, C., Fliegauf, M., Kremer, E., Otto, F., Brenner, O., Lev-Tov, A., and Groner, Y. (2002). The Runx3 transcription factor regulates development and survival of TrkC dorsal root ganglia neurons. EMBO J. 21, 3454–3463. Yoshida, C. A., Yamamoto, H., Fujita, T., Furuichi, T., Ito, K., Inoue, K., Yamana, K., Zanma, A., Takada, K., Ito, Y., and Komori, T. (2004). Runx2 and Runx3 are essential for chondrocyte maturation, and Runx2 regulates limb growth through induction of Indian hedgehog. Genes Dev. 18, 952–963. Mundlos, S., Otto, F., Mundlos, C., Mulliken, J. B., Aylsworth, A. S., Albright, S., Lindhout, D., Cole, W. G., Henn, W., Knoll, J. H. M., Owen, M. J., Mertelsmann, R., Zabel, B. U., and Olsen, B. R. (1997). Mutations involving the transcription factor CBFA1 cause cleidocranial dysplasia. Cell 89, 773–779. Levanon, D., Brenner, O., Negreanu, V., Bettoun, D., Woolf, E., Eilam, R., Lotem, J., Gat, U., Otto, F., Speck, N., and Groner, Y. (2001). Spatial and temporal expression pattern of Runx3 (Aml2) and Runx1 (Aml1) indicates non-redundant functions during mouse embryogenesis. Mech. Dev. 109, 413–417. Yamashiro, T., Aberg, T., Levanon, D., Groner, Y., and Thesleff, I. (2002). Expression of Runx1, -2 and -3 during tooth, palate and craniofacial bone development. Mech. Dev. 119, S107-S110. Smith, N., Dong, Y., Pratap, J., Lian, J. B., Kingsley, P., van Wijnen, A. J., Stein, J. L., Schwarz, E. M., O’Keefe, R. J., Stein, G. S., and Drissi, M. H. (2005). Overlapping expression of Runx1(Cbfa2) and Runx2(Cbfa1) transcription factors supports cooperative induction of skeletal development. J. Cell Pysiol. 203, 133–143. Yamashiro, T., Wang, X. P., Li, Z., Oya, S., Aberg, T., Fukunaga, T., Kamioka, H., Speck, N. A., Takano-Yamamoto, T., and Thesleff, I. (2004). Possible roles of Runx1 and Sox9 in incipient intramembranous ossification. J. Bone Miner. Res. 19, 1671–1677. Stewart, M., Terry, A., Hu, M., O’Hara, M., Blyth, K., Baxter, E., Cameron, E., Onions, D. E., and Neil, J. C. (1997). Proviral insertions induce the expression of bone-specific isoforms of PEBP2alphaA (CBFA1): evidence for a new myc collaborating oncogene. Proc. Natl Acad. Sci. USA. 94, 8646–8651. Ogawa, E., Maruyama, M., Kagoshima, H., Inuzuka, M., Lu, J., Satake, M., Shigesada, K., and Ito, Y. (1993). PEBP2/PEA2 represents

253

Chapter 14 The Cells of Bone

344.

345.

346.

347.

348.

349.

350.

351.

352.

353.

354.

355.

356.

357.

358.

a family of transcription factors homologous to the products of the Drosophila runt gene and the human AML1 gene. Proc. Natl Acad. Sci. USA 90, 6859–6863. Xiao, Z. S., Liu, S. G., Hinson, T. K., and Quarles, L. D. (2001). Characterization of the upstream mouse Cbfa1/Runx2 promoter. J. Cell Biochem. 82, 647–659. Yoshida, C. A., Furuichi, T., Fujita, T., Fukuyama, R., Kanatani, N., Kobayashi, S., Satake, M., Takada, K., and Komori, T. (2002). Corebinding factor beta interacts with Runx2 and is required for skeletal development. Nat. Genet. 32, 633–638. Miller, J., Horner, A., Stacy, T., Lowrey, C., Lian, J. B., Stein, G., Nuckolls, G. H., and Speck, N. A. (2002). The core-binding factor beta subunit is required for bone formation and hematopoietic maturation. Nat. Genet. 32, 645–649. Schroeder, T. M., Kahler, R. A., Li, X., and Westendorf, J. J. (2004). Histone deacetylase 3 interacts with runx2 to repress the osteocalcin promoter and regulate osteoblast differentiation. J. Biol. Chem. 279, 41998–42007. Westendorf, J. J., Zaidi, S. K., Cascino, J. E., Kahler, R., van Wijnen, A. J., Lian, J. B., Yoshida, M., Stein, G. S., and Li, X. (2002). Runx2 (Cbfa1, AML-3) interacts with histone deacetylase 6 and represses the p21(CIP1/WAF1) promoter. Mol. Cell Biol. 22, 7982–7992. Gutierrez, S., Liu, J., Javed, A., Montecino, M., Stein, G. S., Lian, J. B., and Stein, J. L. (2004). The vitamin D response element in the distal osteocalcin promoter contributes to chromatin organization of the proximal regulatory domain. J. Biol. Chem. 279, 43581–43588. Paredes, R., Arriagada, G., Cruzat, F., Olate, J., van Wijnen, A., Lian, J., Stein, G., Stein, J., and Montecino, M. (2004). The Runx2 transcription factor plays a key role in the 1α,25-dihydroxy vitamin D3-dependent upregulation of the rat osteocalcin (OC) gene expression in osteoblastic cells. J. Steroid Biochem. Mol. Biol. 89–90, 269–271. Paredes, R., Arriagada, G., Cruzat, F., Villagra, A., Olate, J., Zaidi, K., van Wijnen, A. J., Lian, J. B., Stein, G. S., Stein, J. L., and Montecino, M. (2004). The bone-specific transcription factor RUNX2 interacts with the 1α,25-dihydroxyvitamin D3 receptor to up-regulate rat osteocalcin gene expression in osteoblastic cells. Mol. Cell Biol. 24, 8847–8861. Vega, R. B., Matsuda, K., Oh, J., Barbosa, A. C., Yang, X., Meadows, E., McAnally, J., Pomajzl, C., Shelton, J. M., Richardson, J. A., Karsenty, G., and Olson, E. N. (2004). Histone deacetylase 4 controls chondrocyte hypertrophy during skeletogenesis. Cell 119, 555–566. Zaidi, S. K., Sullivan, A. J., Medina, R., Ito, Y., van Wijnen, A. J., Stein, J. L., Lian, J. B., and Stein, G. S. (2004). Tyrosine phosphorylation controls Runx2-mediated subnuclear targeting of YAP to repress transcription. EMBO J. 23, 790–799. Shirakabe, K., Terasawa, K., Miyama, K., Shibuya, H., and Nishida, E. (2001). Regulation of the activity of the transcription factor Runx2 by two homeobox proteins, Msx2 and Dlx5. Genes Cells 6, 851–856. Cui, C. B., Cooper, L. F., Yang, X., Karsenty, G., and Aukhil, I. (2003). Transcriptional coactivation of bone-specific transcription factor Cbfa1 by TAZ. Mol. Cell Biol. 23, 1004–1013. Javed, A., Guo, B., Hiebert, S., Choi, J.-Y., Green, J., Zhao, S.-C., Osborne, M. A., Stifani, S., Stein, J. L., Lian, J. B., van Wijnen, A. J., and Stein, G. S. (2000). Groucho/TLE/R-Esp proteins associate with the nuclear matrix and repress RUNX (CBFα/AML/PEBP2α) dependent activation of tissue-specific gene transcription. J. Cell Sci. 113, 2221–2231. Wang, W., Wang, Y. G., Reginato, A. M., Glotzer, D. J., Fukai, N., Plotkina, S., Karsenty, G., and Olsen, B. R. (2004). Groucho homologue Grg5 interacts with the transcription factor Runx2-Cbfa1 and modulates its activity during postnatal growth in mice. Dev. Biol. 270, 364–381. Choi, J.-Y., Pratap, J., Javed, A., Zaidi, S. K., Xing, L., Balint, E., Dalamangas, S., Boyce, B., van Wijnen, A. J., Lian, J. B., Stein, J. L., Jones, S. N., and Stein, G. S. (2001). Subnuclear targeting of

359.

360.

361.

362.

363.

364.

365.

366.

367. 368.

369.

370.

371.

372.

373. 374. 375.

376.

Runx/Cbfa/AML factors is essential for tissue-specific differentiation during embryonic development. Proc. Natl Acad. Sci. USA 98, 8650–8655. Barnes, G. L., Hebert, K. E., Kamal, M., Javed, A., Einhorn, T. A., Lian, J. B., Stein, G. S., and Gerstenfeld, L. C. (2004). Fidelity of Runx2 activity in breast cancer cells is required for the generation of metastases associated osteolytic disease. Cancer Res. 64, 4506–4513. Javed, A., Barnes, G. L., Pratap, J., Antkowiak, T., Gerstenfeld, L. C., van Wijnen, A. J., Stein, J. L., Lian, J. B., and Stein, G. S. (2005). Impaired intranuclear trafficking of Runx2 (AML3/CBFA1) transcription factors in breast cancer cells inhibits osteolysis in vivo. Proc. Natl Acad. Sci. USA 102, 1454–1459. Gutierrez, S., Javed, A., Tennant, D., van Rees, M., Montecino, M., Stein, G. S., Stein, J. L., and Lian, J. B. (2002). CCAAT/enhancerbinding proteins (C/EBP) β and δ Activate osteocalcin gene transcription and synergize with Runx2 at the C/EBP element to regulate bonespecific expression. J. Biol. Chem. 277, 1316–1323. Pereira, R. C., Delany, A. M., and Canalis, E. (2004). CCAAT/enhancer binding protein homologous protein (DDIT3) induces osteoblastic cell differentiation. Endocrinology 145, 1952–1960. Harrison, J. R., Huang, Y. F., Wilson, K. A., Kelly, P. L., Adams, D. J., Gronowicz, G. A., and Clark, S. H. (2005). Col1a1 promoter-targeted expression of p20C/EBPbeta , a truncated C/EBPbeta isoform, causes osteopenia in transgenic mice. J. Biol. Chem. 280, 8117–8124. Sevetson, B., Taylor, S., and Pan, Y. (2004). Cbfa1/RUNX2 directs specific expression of the sclerosteosis gene (SOST). J. Biol. Chem. 279, 13849–13858. Hess, J., Porte, D., Munz, C., and Angel, P. (2001). AP-1 and Cbfa/Runt physically interact and regulate PTH-dependent MMP13 expression in osteoblasts through a new OSE2/AP-1 composite element. J. Biol. Chem. 276, 20029–20038. McCarthy, T. L., Chang, W. Z., Liu, Y., and Centrella, M. (2003). Runx2 integrates estrogen activity in osteoblasts. J. Biol. Chem. 278, 43121–43129. Roodman, G. D. (1995). Osteoclast function in Paget’s disease and multiple myeloma. Bone 17, 57S-61S. Boissy, P., Saltel, F., Bouniol, C., Jurdic, P., and Machuca-Gayet, I. (2002). Transcriptional activity of nuclei in multinucleated osteoclasts and its modulation by calcitonin. Endocrinology 143, 1913–1921. Saltman, L. H., Javed, A., Ribadeneyra, J., Hussain, S., Young, D. W., Osdoby, P., Amcheslavsky, A., van Wijnen, A. J., Stein, J. L., Stein, G. S., Lian, J. B., and Bar-Shavit, Z. (2005). Organization of transcriptional regulatory machinery in osteoclast nuclei: Compartmentalization of Runx1. J. Cell Physiol. 204, 871–880. Lakkakorpi, P. T., and Vaananen, H. K. (1991). Kinetics of the osteoclast cytoskeleton during the resorption cycle in vitro. J. Bone Miner. Res. 6, 817–826. Vaananen, H. K., and Horton, M. (1995). The osteoclast clear zone is a specialized cell-extracellular matrix adhesion structure. J. Cell Sci. 108, 2729–2732. Tanaka, S., Takahashi, N., Udagawa, N., Murakami, H., Nakamura, I., Kurokawa, T., and Suda, T. (1995). Possible involvement of focal adhesion kinase, p125FAK, in osteoclastic bone resorption. J. Cell Biochem. 58, 424–435. Teitelbaum, S. L., and Ross, F. P. (2003). Genetic regulation of osteoclast development and function. Nat. Rev. Genet. 4, 638–649. Boyle, W. J., Simonet, W. S., and Lacey, D. L. (2003). Osteoclast differentiation and activation. Nature 423, 337–342. Engleman, V. W., Nickols, G. A., Ross, F. P., Horton, M. A., Griggs, D. W., Settle, S. L., Ruminski, P. G., and Teitelbaum, S. L. (1997). A peptidomimetic antagonist of the alpha(v)beta3 integrin inhibits bone resorption in vitro and prevents osteoporosis in vivo. J. Clin. Invest. 99, 2284–2292. Duong, L. T., Nakamura, I., Lakkakorpi, P. T., Lipfert, L., Bett, A. J., and Rodan, G. A. (2000). Inhibition of osteoclast function by adenovirus

254

377.

378.

379.

380.

381.

382.

383.

384.

385.

386.

387.

388.

389.

390. 391.

392.

393.

JANE B. LIAN AND GARY S. STEIN expressing antisense protein-tyrosine kinase 2. J. Biol. Chem. 276, 7484–7492. Chellaiah, M. A., Kizer, N., Biswas, R., Alvarez, U., Strauss-Schoenberger, J., Rifas, L., Rittling, S. R., Denhardt, D. T., and Hruska, K. A. (2003). Osteopontin deficiency produces osteoclast dysfunction due to reduced CD44 surface expression. Mol. Biol. Cell 14, 173–189. Faccio, R., Novack, D. V., Zallone, A., Ross, F. P., and Teitelbaum, S. L. (2003). Dynamic changes in the osteoclast cytoskeleton in response to growth factors and cell attachment are controlled by beta3 integrin. J. Cell Biol. 162, 499–509. Arias-Salgado, E. G., Lizano, S., Sarkar, S., Brugge, J. S., Ginsberg, M. H., and Shattil, S. J. (2003). Src kinase activation by direct interaction with the integrin beta cytoplasmic domain. Proc. Natl Acad. Sci. USA 100, 13298–13302. Miyazaki, T., Sanjay, A., Neff, L., Tanaka, S., Horne, W. C., and Baron, R. (2004). Src kinase activity is essential for osteoclast function. J. Biol. Chem. 279, 17660–17666. Soriano, P., Montgomery, C., Geske, R., and Bradley, A. (1991). Targeted disruption of the c-src proto-oncogene leads to osteopetrosis in mice. Cell 64, 693–702. Boyce, B. F., Yoneda, T., Lowe, C., Soriano, P., and Mundy, G. R. (1992). Requirement of pp60c-src expression for osteoclasts to form ruffled borders and resorb bone in mice. J. Clin. Invest. 90, 1622–1627. Chiusaroli, R., Sanjay, A., Henriksen, K., Engsig, M. T., Horne, W. C., Gu, H., and Baron, R. (2003). Deletion of the gene encoding c-Cbl alters the ability of osteoclasts to migrate, delaying resorption and ossification of cartilage during the development of long bones. Dev. Biol. 261, 537–547. Lakkakorpi, P. T., Bett, A. J., Lipfert, L., Rodan, G. A., and Duong, I. T. (2003). PYK2 autophosphorylation, but not kinase activity, is necessary for adhesion-induced association with c-Src, osteoclast spreading, and bone resorption. J. Biol. Chem. 278, 11502–11512. Mulari, M. T., Zhao, H., Lakkakorpi, P. T., and Vaananen, H. K. (2003). Osteoclast ruffled border has distinct subdomains for secretion and degraded matrix uptake. Traffic 4, 113–125. Chatterjee, D., Chakraborty, M., Leit, M., Neff, L., Jamsa-Kellokumpu, S., Fuchs, R., and Baron, R. (1992). Sensitivity to vanadate and isoforms of subunits A and B distinguish the osteoclast proton pump from other vacuolar H+ ATPases. Proc. Natl Acad. Sci. USA. 89, 6257–6261. Toyosawa, S., Ogawa, Y., Chang, C. K., Hong, S. S., Yagi, T., Kuwahara, H., Wakasa, K., and Sakurai, M. (1991). Histochemistry of tartrate-resistant acid phosphatase and carbonic anhydrase isoenzyme II in osteoclast-like giant cells in bone tumours. Virchows Arch. A. Pathol. Anat. Histopathol. 418, 255–261. Li, Y. P., Chen, W., Liang, Y., Li, E., and Stashenko, P. (1999). Atp6ideficient mice exhibit severe osteopetrosis due to loss of osteoclastmediated extracellular acidification. Nat. Genet. 23, 447–451. Teitelbaum, S. L., Tondravi, M. M., and Ross, F. P. (1996). Osteoclast biology. In “Osteoporosis” (R. Marcus, D. Feldman, and J. Kelsey, eds.), pp. 61–94. Academic Press, Inc., San Diego, CA. Teitelbaum, S. L. (2000). Bone resorption by osteoclasts. Science 289, 1504–1508. Saftig, P., Hunziker, E., Wehmeyer, O., Jones, S., Boyde, A., Rommerskirch, W., Moritz, J. D., Schu, P., and von Figura, K. (1998). Impaired osteoclastic bone resorption leads to osteopetrosis in cathepsin-K-deficient mice. Proc. Natl Acad. Sci. USA 95, 13453–13458. Saftig, P., Hunziker, E., Everts, V., Jones, S., Boyde, A., Wehmeyer, O., Suter, A., and von Figura, K. (2000). Functions of cathepsin K in bone resorption. Lessons from cathepsin K deficient mice. Adv. Exp. Med. Biol. 477, 293–303. Hollberg, K., Hultenby, K., Hayman, A., Cox, T., and Andersson, G. (2002). Osteoclasts from mice deficient in tartrate-resistant acid

394.

395.

396.

397. 398. 399.

400.

401.

402.

403.

404.

405.

406.

407.

408.

409.

410.

phosphatase have altered ruffled borders and disturbed intracellular vesicular transport. Exp. Cell Res. 279, 227–238. Bergers, G., Brekken, R., McMahon, G., Vu, T. H., Itoh, T., Tamaki, K., Tanzawa, K., Thorpe, P., Itohara, S., Werb, Z., and Hanahan, D. (2000). Matrix metalloproteinase-9 triggers the angiogenic switch during carcinogenesis. Nat. Cell Biol. 2, 737–744. Engsig, M. T., Chen, Q. J., Vu, T. H., Pedersen, A. C., Therkidsen, B., Lund, L. R., Henriksen, K., Lenhard, T., Foged, N. T., Werb, Z., and Delaisse, J. M. (2000). Matrix metalloproteinase 9 and vascular endothelial growth factor are essential for osteoclast recruitment into developing long bones. J. Cell Biol. 151, 879–890. Popoff, S. N., and Marks, S. C. Jr. (1995). The heterogeneity of the osteopetroses reflects the diversity of cellular influences during skeletal development. Bone 17, 437–445. Suda, T., Nakamura, I., Jimi, E., and Takahashi, N. (1997). Regulation of osteoclast function. J. Bone Miner. Res. 12, 869–879. Roodman, G. D. (1996). Advances in bone biology: the osteoclast. Endocr. Rev. 17, 308–332. Grey, A., Chen, Y., Paliwal, I., Carlberg, K., and Insogna, K. (2000). Evidence for a functional association between phosphatidylinositol 3- kinase and c-src in the spreading response of osteoclasts to colonystimulating factor-1. Endocrinology 141, 2129–2138. Felix, R., Cecchini, M. G., and Fleisch, H. (1990). Macrophage colony-stimulating factor restores in vivo bone resorption in the op/op osteopetrotic mouse. Endocrinology 127, 2592–2594. Yoshida, H., Hayashi, S., Kunisada, T., Ogawa, M., Nishikawa, S., Okamura, H., Sudo, T., and Shultz, L. D. (1990). The murine mutation osteopetrosis is in the coding region of the macrophage colony stimulating factor gene. Nature 345, 442–444. Takahashi, N., Udagawa, N., Akatsu, T., Tanaka, H., Isogai, Y., and Suda, T. (1991). Deficiency of osteoclasts in osteopetrotic mice is due to a defect in the local microenvironment provided by osteoblastic cells. Endocrinology 128, 1792–1796. Luchin, A., Purdom, G., Murphy, K., Clark, M. Y., Angel, N., Cassady, A. I., Hume, D. A., and Ostrowski, M. C. (2000). The microphthalmia transcription factor regulates expression of the tartrateresistant acid phosphatase gene during terminal differentiation of osteoclasts. J. Bone Miner. Res. 15, 451–460. Mansky, K. C., Sankar, U., Han, J., and Ostrowski, M. C. (2002). Microphthalmia transcription factor is a target of the p38 MAPK pathway in response to receptor activator of NF-kappa B ligand signaling. J. Biol. Chem. 277, 11077–11083. Weilbaecher, K. N., Motyckova, G., Huber, W. E., Takemoto, C. M., Hemesath, T. J., Xu, Y., Hershey, C. L., Dowland, N. R., Wells, A. G., and Fisher, D. E. (2001). Linkage of M-CSF signaling to Mitf, TFE3, and the osteoclast defect in Mitf(mi/mi) mice. Mol. Cell 8, 749–758. Kawaguchi, N., Ono, T., Mochii, M., and Noda, M. (2001). Spontaneous mutation in Mitf gene causes osteopetrosis in silver homozygote quail. Dev. Dyn. 220, 133–140. Lakkakorpi, P. T., Nakamura, I., Nagy, R. M., Parsons, J. T., Rodan, G. A., and Duong, L. T. (1999). Stable association of PYK2 and p130(Cas) in osteoclasts and their co-localization in the sealing zone. J. Biol. Chem. 274, 4900–4907. Nakamura, I., Lipfert, L., Rodan, G. A., and Duong, L. T. (2001). Convergence of alpha(v)beta(3) integrin- and macrophage colony stimulating factor-mediated signals on phospholipase Cgamma in prefusion osteoclasts. J. Cell Biol. 152, 361–374. Nakamura, I., Rodan, G. A., and Duong, l. T. (2003). Distinct roles of p130Cas and c-Cbl in adhesion-induced or macrophage colonystimulating factor-mediated signaling pathways in prefusion osteoclasts. Endocrinology 144, 4739–4741. Horwood, N. J., Kartsogiannis, V., Quinn, J. M., Romas, E., Martin, T. J., and Gillespie, M. T. (1999). Activated T lymphocytes support osteoclast formation in vitro. Biochem. Biophys. Res. Commun. 265, 144–150.

Chapter 14 The Cells of Bone 411. Lomaga, M. A., Yeh, W. C., Sarosi, I., Duncan, G. S., Furlonger, C., Ho, A., Morony, S., Capparelli, C., Van, G., Kaufman, S., van der, H. A., Itie, A., Wakeham, A., Khoo, W., Sasaki, T., Cao, Z., Penninger, J. M., Paige, C. J., Lacey, D. L., Dunstan, C. R., Boyle, W. J., Goeddel, D. V., and Mak, T. W. (1999). TRAF6 deficiency results in osteopetrosis and defective interleukin-1, CD40, and LPS signaling. Genes Dev. 13, 1015–1024. 412. Armstrong, A. P., Tometsko, M. E., Glaccum, M., Sutherland, C. L., Cosman, D., and Dougall, W. C. (2002). A RANK/TRAF6-dependent signal transduction pathway is essential for osteoclast cytoskeletal organization and resorptive function. J. Biol. Chem. 277, 44347–44356. 413. Naito, A., Azuma, S., Tanaka, S., Miyazaki, T., Takaki, S., Takatsu, K., Nakao, K., Nakamura, K., Katsuki, M., Yamamoto, T., and Inoue, J. (1999). Severe osteopetrosis, defective interleukin-1 signalling and lymph node organogenesis in TRAF6-deficient mice. Genes Cells 4, 353–362. 414. Simonet, W. S., Lacey, D. L., Dunstan, C. R., Kelley, M., Chang, M. S., Luthy, R., Nguyen, H. Q., Wooden, S., Bennett, L., Boone, T., Shimamoto, G., DeRose, M., Elliott, R., Colombero, A., Tan, H. L., Trail, G., Sullivan, J., Davy, E., Bucay, N., Renshaw-Gegg, L., Hughes, T. M., Hill, D., Pattison, W., Campbell, P., and Boyle, W. J. (1997). Osteoprotegerin: a novel secreted protein involved in the regulation of bone density. Cell 89, 309–319. 415. Wong, B. R., Josien, R., Lee, S. Y., Sauter, B., Li, H. L., Steinman, R. M., and Choi, Y. (1997). TRANCE (tumor necrosis factor [TNF]-related activation-induced cytokine), a new TNF family member predominantly expressed in T cells, is a dendritic cell-specific survival factor. J. Exp. Med. 186, 2075–2080. 416. Yasuda, H., Shima, N., Nakagawa, N., Mochizuki, S. I., Yano, K., Fujise, N., Sato, Y., Goto, M., Yamaguchi, K., Kuriyama, M., Kanno, T., Murakami, A., Tsuda, E., Morinaga, T., and Higashio, K. (1998). Identity of osteoclastogenesis inhibitory factor (OCIF) and osteoprotegerin (OPG): a mechanism by which OPG/OCIF inhibits osteoclastogenesis in vitro. Endocrinology 139, 1329–1337. 417. Yasuda, H., Shima, N., Nakagawa, N., Yamaguchi, K., Kinosaki, M., Mochizuki, S., Tomoyasu, A., Yano, K., Goto, M., Murakami, A., Tsuda, E., Morinaga, T., Higashio, K., Udagawa, N., Takahashi, N., and Suda, T. (1998). Osteoclast differentiation factor is a ligand for osteoprotegerin/osteoclastogenesis-inhibitory factor and is identical to TRANCE/RANKL. Proc. Natl Acad. Sci. USA 95, 3597–3602. 418. Lacey, D. L., Timms, E., Tan, H. L., Kelley, M. J., Dunstan, C. R., Burgess, T., Elliott, R., Colombero, A., Elliott, G., Scully, S., Hsu, H., Sullivan, J., Hawkins, N., Davy, E., Capparelli, C., Eli, A., Qian, Y. X., Kaufman, S., Sarosi, I., Shalhoub, V., Senaldi, G., Guo, J., Delaney, J., and Boyle, W. J. (1998). Osteoprotegerin ligand is a cytokine that regulates osteoclast differentiation and activation. Cell 93, 165–176. 419. Yu, X., Huang, Y., Collin-Osdoby, P., and Osdoby, P. (2003). Stromal cell-derived factor-1 (SDF-1) recruits osteoclast precursors by inducing chemotaxis, matrix metalloproteinase-9 (MMP-9) activity, and collagen transmigration. J. Bone Miner. Res. 18, 1404–1418. 420. Miyaura, C., Inada, M., Suzawa, T., Sugimoto, Y., Ushikubi, F., Ichikawa, A., Narumiya, S., and Suda, T. (2000). Impaired bone resorption to prostaglandin E2 in prostaglandin E receptor EP4-knockout mice. J. Biol. Chem. 275, 19819–19823. 421. Li, X., Okada, Y., Pilbeam, C. C., Lorenzo, J. A., Kennedy, C. R., Breyer, R. M., and Raisz, L. G. (2000). Knockout of the murine prostaglandin EP2 receptor impairs osteoclastogenesis in vitro. Endocrinology 141, 2054–2061. 422. Lee, S. E., Chung, W. J., Kwak, H. B., Chung, C. H., Kwack, K. B., Lee, Z. H., and Kim, H. H. (2001). Tumor necrosis factor-alpha supports the survival of osteoclasts through the activation of Akt and ERK. J. Biol. Chem. 276, 49343–49349. 423. Girasole, G., Passeri, G., Jilka, R. L., and Manolagas, S. C. (1994). Interleukin-11: a new cytokine critical for osteoclast development. J. Clin. Invest. 93, 1516–1524.

255 424. Romas, E., Udagawa, N., Zhou, H., Tamura, T., Saito, M., Taga, T., Hilton, D. J., Suda, T., Ng, K. W., and Martin, T. J. (1996). The role of gp130-mediated signals in osteoclast development: regulation of interleukin 11 production by osteoblasts and distribution of its receptor in bone marrow cultures. J. Exp. Med. 183, 2581–2591. 425. Elias, J. A., Tang, W., and Horowitz, M. C. (1995). Cytokine and hormonal stimulation of human osteosarcoma interleukin-11 production. Endocrinology 136, 489–498. 426. Suda, T., Udagawa, N., Nakamura, I., Miyaura, C., and Takahashi, N. (1995). Modulation of osteoclast differentiation by local factors. Bone 17, 87S-91S. 427. Devlin, R. D., Reddy, S. V., Savino, R., Ciliberto, G., and Roodman, G. D. (1998). IL-6 mediates the effects of IL-1 or TNF, but not PTHrP or 1, 25(OH)2D3, on osteoclast-like cell formation in normal human bone marrow cultures. J. Bone Miner. Res. 13, 393–399. 428. Kishimoto, T., Akira, S., Narazaki, M., and Taga, T. (1995). Interleukin-6 family of cytokines and gp130. Blood 86, 1243–1254. 429. Lee, S. K., Kalinowski, J. F., Jastrzebski, S. L., Puddington, L., and Lorenzo, J. A. (2003). Interleukin-7 is a direct inhibitor of in vitro osteoclastogenesis. Endocrinology 144, 3524–3531. 430. Udagawa, N., Horwood, N. J., Elliott, J., Mackay, A., Owens, J., Okamura, H., Kurimoto, M., Chambers, T. J., Martin, T. J., and Gillespie, M. T. (1997). Interleukin-18 (interferon-gamma-inducing factor) is produced by osteoblasts and acts via granulocyte/macrophage colony-stimulating factor and not via interferon-gamma to inhibit osteoclast formation. J. Exp. Med. 185, 1005–1012. 431. Horwood, N. J., Udagawa, N., Elliott, J., Grail, D., Okamura, H., Kurimoto, M., Dunn, A. R., Martin, T., and Gillespie, M. T. (1998). Interleukin 18 inhibits osteoclast formation via T cell production of granulocyte macrophage colony-stimulating factor. J. Clin. Invest. 101, 595–603. 432. Suda, T., Ueno, Y., Fujii, K., and Shinki, T. (2003). Vitamin D and bone. J. Cell Biochem. 88, 259–266. 433. Kurihara, N., Reddy, S. V., Araki, N., Ishizuka, S., Ozono, K., Cornish, J., Cundy, T., Singer, F. R., and Roodman, G. D. (2004). Role of TAFII-17, a VDR binding protein, in the increased osteoclast formation in Paget’s Disease. J. Bone Miner. Res. 19, 1154–1164. 434. Faucheux, C., Horton, M. A., and Price, J. S. (2002). Nuclear localization of type I parathyroid hormone/parathyroid hormone-related protein receptors in deer antler osteoclasts: evidence for parathyroid hormone-related protein and receptor activator of NF-kappaBdependent effects on osteoclast formation in regenerating mammalian bone. J. Bone Miner. Res. 17, 455–464. 435. Nakchbandi, I. A., Weir, E. E., Insogna, K. L., Philbrick, W. M., and Broadus, A. E. (2000). Parathyroid hormone-related protein induces spontaneous osteoclast formation via a paracrine cascade. Proc. Natl Acad. Sci. USA 97, 7296–7300. 436. Vilardaga, J. P., Lin, I., and Nissenson, R. A. (2001). Analysis of parathyroid hormone (PTH)/secretin receptor chimeras differentiates the role of functional domains in the pth/ pth-related peptide (PTHrP) receptor on hormone binding and receptor activation. Mol. Endocrinol. 15, 1186–1199. 437. Huang, Z., Chen, Y., Pratt, S., Chen, T. H., Bambino, T., Nissenson, R. A., and Shoback, D. M. (1996). The N-terminal region of the third intracellular loop of the parathyroid hormone (PTH)/PTH-related peptide receptor is critical for coupling to cAMP and inositol phosphate/Ca2+ signal transduction pathways. J. Biol. Chem. 271, 33382–33389. 438. Fuller, K., Owens, J. M., and Chambers, T. J. (1998). Induction of osteoclast formation by parathyroid hormone depends on an action on stromal cells. J. Endocrinol. 158, 341–350. 439. Gay, C. V., Zheng, B., and Gilman, V. R. (2003). Co-detection of PTH/PTHrP receptor and tartrate resistant acid phosphatase in osteoclasts. J. Cell Biochem. 89, 902–908.

256 440. Tong, H., Lin, H., Wang, H., Sakai, D., and Minkin, C. (1995). Osteoclasts respond to parathyroid hormone and express mRNA for its receptor. J. Bone Miner. Res. 10[Suppl. 1], S322. (Abstract). 441. Teti, A., Rizzoli, R., and Zambonin Zallone, A. (1991). Parathyroid hormone binding to cultured avian osteoclasts. Biochem. Biophys. Res. Commun. 174, 1217–1222. 442. Oreffo, R. O., Kusec, V., Virdi, A. S., Flanagan, A. M., Grano, M., Zambonin-Zallone, A., and Triffitt, J. T. (1999). Expression of estrogen receptor-alpha in cells of the osteoclastic lineage. Histochem. Cell Biol. 111, 125–133. 443. Orwoll, E. S. (2003). Men, bone and estrogen: unresolved issues. Osteoporos. Int. 14, 93–98. 444. Liu, B. Y., Wu, P. W., Bringhurst, F. R., and Wang, J. T. (2002). Estrogen inhibition of PTH-stimulated osteoclast formation and attachment in vitro: involvement of both PKA and PKC. Endocrinology 143, 627–635. 445. Swarthout, J. T., D’Alonzo, R. C., Selvamurugan, N., and Partridge, N. C. (2002). Parathyroid hormone-dependent signaling pathways regulating genes in bone cells. Gene 282, 1–17. 446. Fu, Q., Jilka, R. L., Manolagas, S. C., and O’Brien, C. A. (2002). Parathyroid hormone stimulates receptor activator of NFkappa B ligand and inhibits osteoprotegerin expression via protein kinase A activation of cAMP-response element-binding protein. J. Biol. Chem. 277, 48868–48875. 447. Locklin, R. M., Khosla, S., Turner, R. T., and Riggs, B. L. (2003). Mediators of the biphasic responses of bone to intermittent and continuously administered parathyroid hormone. J. Cell Biochem. 89, 180–190. 448. Demiralp, B., Chen, H. L., Koh, A. J., Keller, E. T., and McCauley, L. K. (2002). Anabolic actions of parathyroid hormone during bone growth are dependent on c-fos. Endocrinology 143, 4038–4047. 449. Bikle, D. D., Sakata, T., Leary, C., Elalieh, H., Ginzinger, D., Rosen, C. J., Beamer, W., Majumdar, S., and Halloran, B. P. (2002). Insulin-like growth factor I is required for the anabolic actions of parathyroid hormone on mouse bone. J. Bone Miner. Res. 17, 1570–1578. 450. Yamaguchi, M., Ogata, N., Shinoda, Y., Akune, T., Kamekura, S., Terauchi, Y., Kadowaki, T., Hoshi, K., Chung, U. I., Nakamura, K., and Kawaguchi, H. (2005). Insulin receptor substrate-1 is required for bone anabolic function of parathyroid hormone in mice. Endocrinology 146, 2620–2628. 451. Manolagas, S. C., Kousteni, S., and Jilka, R. L. (2002). Sex steroids and bone. Recent Prog. Horm. Res. 57, 385–409. 452. Srivastava, S., Toraldo, G., Weitzmann, M. N., Cenci, S., Ross, F. P., and Pacifici, R. (2001). Estrogen decreases osteoclast formation by down-regulating receptor activator of NF-kappa B ligand (RANKL)induced JNK activation. J. Biol. Chem. 276, 8836–8840. 453. Srivastava, S., Weitzmann, M. N., Kimble, R. B., Rizzo, M., Zahner, M., Milbrandt, J., Ross, F. P., and Pacifici, R. (1998). Estrogen blocks M-CSF gene expression and osteoclast formation by regulating phosphorylation of Egr-1 and its interaction with Sp-1. J. Clin.Invest. 102, 1850–1859. 454. Sunyer, T., Lewis, J., Collin-Osdoby, P., and Osdoby, P. (1999). Estrogen’s bone-protective effects may involve differential IL-1 receptor regulation in human osteoclast-like cells. J. Clin. Invest. 103, 1409–1418. 455. Shevde, N. K., Bendixen, A. C., Dienger, K. M., and Pike, J. W. (2000). Estrogens suppress RANK ligand-induced osteoclast differentiation via a stromal cell independent mechanism involving c-Jun repression. Proc. Natl Acad. Sci. USA 97, 7829–7834. 456. Bord, S., Ireland, D. C., Beavan, S. R., and Compston, J. E. (2003). The effects of estrogen on osteoprotegerin, RANKL, and estrogen receptor expression in human osteoblasts. Bone 32, 136–141. 457. Windahl, S. H., Hollberg, K., Vidal, O., Gustafsson, J. A., Ohlsson, C., and Andersson, G. (2001). Female estrogen receptor beta-/- mice are

JANE B. LIAN AND GARY S. STEIN

458.

459.

460.

461.

462. 463.

464.

465.

466.

467.

468.

469.

470.

471.

472.

473.

474.

partially protected against age-related trabecular bone loss. J. Bone Miner. Res. 16, 1388–1398. Robinson, J. A., Riggs, B. L., Spelsberg, T. C., and Oursler, M. J. (1996). Osteoclasts and transforming growth factor-beta: estrogenmediated isoform-specific regulation of production. Endocrinology 137, 615–621. van der Eerden, B. C., Lowik, C. W., Wit, J. M., and Karperien, M. (2004). Expression of estrogen receptors and enzymes involved in sex steroid metabolism in the rat tibia during sexual maturation. J. Endocrinol. 180, 457–467. Egerbacher, M., Helmreich, M., Rossmanith, W., and Haeusler, G. (2002). Estrogen receptor-alpha and estrogen receptor-beta are present in the human growth plate in childhood and adolescence, in identical distribution. Horm. Res. 58, 99–103. Pascoe, D., and Oursler, M. J. (2001). The Src signaling pathway regulates osteoclast lysosomal enzyme secretion and is rapidly modulated by estrogen. J. Bone Miner. Res. 16, 1028–1036. Khosla, S., Melton, L. J., III, and Riggs, B. L. (2001). Estrogens and bone health in men. Calcif. Tissue Int. 69, 189–192. Lee, S. K., Goldring, S. R., and Lorenzo, J. A. (1995). Expression of the calcitonin receptor in bone marrow cell cultures and in bone: a specific marker of the differentiated osteoclast that is regulated by calcitonin. Endocrinology 136, 4572–4581. Hattersley, G., and Chambers, T. J. (1989). Calcitonin receptors as markers for osteoclastic differentiation: correlation between generation of bone-resorptive cells and cells that express calcitonin receptors in mouse bone marrow cultures. Endocrinology 125, 1606–1612. Kajiya, H., Okamoto, F., Fukushima, H., and Okabe, K. (2003). Calcitonin inhibits proton extrusion in resorbing rat osteoclasts via protein kinase A. Pflugers Arch. 445, 651–658. Komarova, S. V., Shum, J. B., Paige, L. A., Sims, S. M., and Dixon, S. J. (2003). Regulation of osteoclasts by calcitonin and amphiphilic calcitonin conjugates: role of cytosolic calcium. Calcif. Tissue Int. 73, 265–273. Nakamura, I., Takahashi, N., Sasaki, T., Tanaka, S., Udagawa, N., Murakami, H., Kimura, K., Kabuyama, Y., Kurokawa, T., and Suda, T. (1995). Wortmannin, a specific inhibitor of phosphatidylinositol-3 kinase, blocks osteoclastic bone resorption. FEBS Lett. 361, 79–84. Lakkakorpi, P. T., and Vaananen, H. K. (1990). Calcitonin, prostaglandin E2, and dibutyryl cyclic adenosine 3′, 5′-monophosphate disperse the specific microfilament structure in resorbing osteoclasts. J. Histochem. Cytochem. 38, 1487–1493. Hoff, A. O., Catala-Lehnen, P., Thomas, P. M., Priemel, M., Rueger, J. M., Nasonkin, I., Bradley, A., Hughes, M. R., Ordonez, N., Cote, G. J., Amling, M., and Gagel, R. F. (2002). Increased bone mass is an unexpected phenotype associated with deletion of the calcitonin gene. J. Clin. Invest. 110, 1849–1857. Zaidi, M., Moonga, B. S., and Abe, E. (2002). Calcitonin and bone formation: a knockout full of surprises. J. Clin. Invest. 110, 1769–1771. Kusu, N., Laurikkala, J., Imanishi, M., Usui, H., Konishi, M., Miyake, A., Thesleff, I., and Itoh, N. (2003). Sclerostin is a novel secreted osteoclast-derived bone morphogenetic protein antagonist with unique ligand specificity. J. Biol. Chem. 278, 24113–24117. Marzia, M., Sims, N. A., Voit, S., Migliaccio, S., Taranta, A., Bernardini, S., Faraggiana, T., Yoneda, T., Mundy, G. R., Boyce, B. F., Baron, R., and Teti, A. (2000). Decreased c-Src expression enhances osteoblast differentiation and bone formation. J. Cell Biol. 151, 311–320. Zaidi, M., Moonga, B. S., and Huang, C. L. (2004). Calcium sensing and cell signaling processes in the local regulation of osteoclastic bone resorption. Biol. Rev. Camb. Philos. Soc. 79, 79–100. Kanatani, M., Sugimoto, T., Kano, J., Kanzawa, M., and Chihara, K. (2003). Effect of high phosphate concentration on osteoclast differentiation as well as bone-resorbing activity. J. Cell Physiol. 196, 180–189.

Chapter 14 The Cells of Bone 475. Dacquin, R., Davey, R. A., Laplace, C., Levasseur, R., Morris, H. A., Goldring, S. R., Gebre-Medhin, S., Galson, D. L., Zajac, J. D., and Karsenty, G. (2004). Amylin inhibits bone resorption while the calcitonin receptor controls bone formation in vivo. J. Cell Biol. 164, 509–514. 476. Kobayashi, Y., Hashimoto, F., Miyamoto, H., Kanaoka, K., MiyazakiKawashita, Y., Nakashima, T., Shibata, M., Kobayashi, K., Kato, Y., and Sakai, H. (2000). Force-induced osteoclast apoptosis in vivo is accompanied by elevation in transforming growth factor beta and osteoprotegerin expression. J. Bone Miner. Res. 15, 1924–1934. 477. McLean, W., and Olsen, B. R. (2001). Mouse models of abnormal skeletal development and homeostasis. Trends Genet. 17, S38-S43. 478. Marks, S. C., Jr., and Odgren, P. R. (2002). Structure and development of the skeleton. In “Principles of Bone Biology” (J. P. Bilezikian, L. G. Raisz, and G. A. Rodan, eds.), pp. 3–15. Academic Press, San Diego, CA. 479. Johnson, R. S., Spiegelman, B. M., and Papaioannou, V. (1992). Pleiotropic effects of a null mutation in the c-fos proto- oncogene. Cell 71, 577–586. 480. Grigoriadis, A. E., Wang, Z.-Q., Cecchini, M. G., Hofstetter, W., Felix, R., Fleisch, H. A., and Wagner, E. F. (1994). c-Fos: a key regulator of osteoclast-macrophage lineage determination and bone remodeling. Science 266, 443–448. 481. Wang, Z.-Q., Ovitt, C., Grigoriadis, A. E., Mohle-Steinlein, U., Ruther, U., and Wagner, E. F. (1992). Bone and haematopoietic defects in mice lacking c-fos. Nature 360, 741–745. 482. Tondravi, M. M., McKercher, S. R., Anderson, K., Erdmann, J. M., Quiroz, M., Maki, R., and Teitelbaum, S. L. (1997). Osteopetrosis in mice lacking haematopoietic transcription factor PU.1. Nature 386, 81–84. 483. Zhang, D. E., Hetherington, C. J., Chen, H. M., and Tenen, D. G. (1994). The macrophage transcription factor PU.1 directs tissue-specific expression of the macrophage colony-stimulating factor receptor. Mol. Cell Biol. 14, 373–381. 484. Xing, L., Franzoso, G., Siebenlist, U., and Boyce, B. (1997). Targeted disruption of the p50 and p52 subunits of NF-kB transcription factor leads to failure of osteoclast formation and osteopetrosis. J. Bone Miner. Res. 12, S136. (Abstract). 485. Ikeda, F., Nishimura, R., Matsubara, T., Tanaka, S., Inoue, J., Reddy, S. V., Hata, K., Yamashita, K., Hiraga, T., Watanabe, T., Kukita, T., Yoshioka, K., Rao, A., and Yoneda, T. (2004). Critical roles of c-Jun signaling in regulation of NFAT family and RANKL-regulated osteoclast differentiation. J. Clin. Invest. 114, 475–484. 486. Komarova, S. V., Pereverzev, A., Shum, J. W., Sims, S. M., and Dixon, S. J. (2005). Convergent signaling by acidosis and receptor activator of NF-kappaB ligand (RANKL) on the calcium/calcineurin/ NFAT pathway in osteoclasts. Proc. Natl Acad. Sci. USA 102, 2643–2648. 487. Ortega-Perez, I., Cano, E., Were, F., Villar, M., Vazquez, J., and Redondo, J. M. (2005). c-Jun NH2-terminal kinase (JNK) positively regulates NFATc2 transactivation through phosphorylation within the N-terminal regulatory domain. J. Biol. Chem. 280, 20867–20878. 488. Matsuo, K., Galson, D. L., Zhao, C., Peng, L., Laplace, C., Wang, K. Z., Bachler, M. A., Amano, H., Aburatani, H., Ishikawa, H., and Wagner, E. F. (2004). Nuclear factor of activated T-cells (NFAT) rescues osteoclastogenesis in precursors lacking c-Fos. J. Biol. Chem. 279, 26475–26480. 489. Steingrimsson, E., Tessarollo, L., Pathak, B., Hou, L., Arnheiter, H., Copeland, N. G., and Jenkins, N. A. (2002). Mitf and Tfe3, two members of the Mitf-Tfe family of bHLH-Zip transcription factors, have important but functionally redundant roles in osteoclast development. Proc. Natl Acad. Sci. USA 99, 4477–4482. 490. Hershey, C. L., and Fisher, D. E. (2004). Mitf and Tfe3: members of a b-HLH-ZIP transcription factor family essential for osteoclast development and function. Bone 34, 689–696.

257 491. Hodgkinson, C. A., Moore, K. J., Nakayama, A., Steingrimsson, E., Copeland, N. G., Jenkins, N. A., and Arnheiter, H. (1993). Mutations at the mouse microphthalmia locus are associated with defects in a gene encoding a novel basic-helix-loop-helix-zipper protein. Cell 74, 395–404. 492. Lu, M. F., Cheng, H. T., Lacy, A. R., Kern, M. J., Argao, E. A., Potter, S. S., Olson, E. N., and Martin, J. F. (1999). Paired-related homeobox genes cooperate in handplate and hindlimb zeugopod morphogenesis. Dev. Biol. 205, 145–157. 493. Herbrand, H., Pabst, O., Hill, R., and Arnold, H. H. (2002). Transcription factors Nkx3.1 and Nkx3.2 (Bapx1) play an overlapping role in sclerotomal development of the mouse. Mech. Dev. 117, 217–224. 494. Takeuchi, J. K., Koshiba-Takeuchi, K., Suzuki, T., Kamimura, M., Ogura, K., and Ogura, T. (2003). Tbx5 and Tbx4 trigger limb initiation through activation of the Wnt/Fgf signaling cascade. Development 130, 2729–2739. 495. Smits, P., Dy, P., Mitra, S., and Lefebvre, V. (2004). Sox5 and Sox6 are needed to develop and maintain source, columnar, and hypertrophic chondrocytes in the cartilage growth plate. J. Cell Biol. 164, 747–758. 496. Smits, P., and Lefebvre, V. (2003). Sox5 and Sox6 are required for notochord extracellular matrix sheath formation, notochord cell survival and development of the nucleus pulposus of intervertebral discs. Development 130, 1135–1148. 497. Pfander, D., Cramer, T., Schipani, E., and Johnson, R. S. (2003). HIF1alpha controls extracellular matrix synthesis by epiphyseal chondrocytes. J. Cell Sci. 116, 1819–1826. 498. Bi, W., Deng, J. M., Zhang, Z., Behringer, R. R., and de Crombrugghe, B. (1999). Sox9 is required for cartilage formation. Nat. Genet. 22, 85–89. 499. Rice, R., Rice, D. P., Olsen, B. R., and Thesleff, I. (2003). Progression of calvarial bone development requires Foxc1 regulation of Msx2 and Alx4. Dev. Biol. 262, 75–87. 500. Bendall, A. J., and Abate-Shen, C. (2000). Roles for Msx and Dlx homeoproteins in vertebrate development. Gene 247, 17–31. 501. Miao, D., Liu, H., Plut, P., Niu, M., Huo, R., Goltzman, D., and Henderson, J. E. (2004). Impaired endochondral bone development and osteopenia in Gli2-deficient mice. Exp. Cell Res. 294, 210–222. 502. Buttitta, L., Mo, R., Hui, C. C., and Fan, C. M. (2003). Interplays of Gli2 and Gli3 and their requirement in mediating Shh-dependent sclerotome induction. Development 130, 6233–6243. 503. Vortkamp, A., Lee, K., Lanske, B., Segre, G. V., Kronenberg, H. M., and Tabin, C. J. (1996). Regulation of rate of cartilage differentiation by Indian hedgehog and PTH-related protein. Science 273, 613–622. 504. Jemtland, R., Divieti, P., Lee, K., and Segre, G. V. (2003). Hedgehog promotes primary osteoblast differentiation and increases PTHrP mRNA expression and iPTHrP secretion. Bone 32, 611–620. 505. Hsu, S. C., Galceran, J., and Grosschedl, R. (1998). Modulation of transcriptional regulation by LEF-1 in response to Wnt-1 signaling and association with beta-catenin. Mol. Cell Biol. 18, 4807–4818. 506. Kim, S., Koga, T., Isobe, M., Kern, B. E., Yokochi, T., Chin, Y. E., Karsenty, G., Taniguchi, T., and Takayanagi, H. (2003). Stat1 functions as a cytoplasmic attenuator of Runx2 in the transcriptional program of osteoblast differentiation. Genes Dev. 17, 1979–1991. 507. Xiao, L., Naganawa, T., Obugunde, E., Gronowicz, G., Ornitz, D. M., Coffin, J. D., and Hurley, M. M. (2004). Stat1 controls postnatal bone formation by regulating fibroblast growth factor signaling in osteoblasts. J. Biol. Chem. 279, 27743–27752. 508. Takayanagi, H., Kim, S., Koga, T., and Taniguchi, T. (2005). Stat1mediated cytoplasmic attenuation in osteoimmunology. J. Cell Biochem. 94, 232–240. 509. Sunters, A., Thomas, D. P., Yeudall, W. A., and Grigoriadis, A. E. (2004). Accelerated cell cycle progression in osteoblasts overexpressing the c-fos proto-oncogene: induction of cyclin A and enhanced CDK2 activity. J. Biol. Chem. 279, 9882–9891.

258 510. Ichida, F., Nishimura, R., Hata, K., Matsubara, T., Ikeda, F., Hisada, K., Yatani, H., Cao, X., Komori, T., Yamaguchi, A., and Yoneda, T. (2004). Reciprocal roles of MSX2 in regulation of osteoblast and adipocyte differentiation. J. Biol. Chem. 279, 34015–34022. 511. Dodig, M., Tadic, T., Kronenberg, M. S., Dacic, S., Liu, Y. H., Maxson, R., Rowe, D. W., and Lichtler, A. C. (1999). Ectopic Msx2 overexpression inhibits and Msx2 antisense stimulates calvarial osteoblast differentiation. Dev. Biol. 209, 298–307. 512. Yoshizawa, T., Takizawa, F., Iizawa, F., Ishibashi, O., Kawashima, H., Matsuda, A., Endo, N., and Kawashima, H. (2004). Homeobox protein MSX2 acts as a molecular defense mechanism for preventing ossification in ligament fibroblasts. Mol. Cell Biol. 24, 3460–3472. 513. Cheng, S. L., Shao, J. S., Charlton-Kachigian, N., Loewy, A. P., and Towler, D. A. (2003). Msx2 promotes osteogenesis and suppresses adipogenic differentiation of multipotent mesenchymal progenitors. J. Biol. Chem. 278, 45969–45977. 514. Ishii, M., Merrill, A. E., Chan, Y. S., Gitelman, I., Rice, D. P., Sucov, H. M., and Maxson, R. E., Jr. (2003). Msx2 and Twist cooperatively control the development of the neural crest-derived skeletogenic mesenchyme of the murine skull vault. Development 130, 6131–6142. 515. Zamurovic, N., Cappellen, D., Rohner, D., and Susa, M. (2004). Coordinated activation of Notch, Wnt and TGF-beta signaling pathways in BMP-2 induced osteogenesis: Notch target gene Hey1 inhibits mineralization and Runx2 transcriptional activity. J. Biol. Chem. 279, 37704–37715. 516. Lian, J. B., Javed, A., Zaidi, S. K., Lengner, C., Montecino, M., van Wijnen, A. J., Stein, J. L., and Stein, G. S. (2004). Regulatory controls for osteoblast growth and differentiation: role of Runx/Cbfa/AML factors. Crit. Rev. Eukaryot. Gene Expr. 14, 1–41. 517. Hess, J., Hartenstein, B., Teurich, S., Schmidt, D., Schorpp-Kistner, M., and Angel, P. (2003). Defective endochondral ossification in mice with strongly compromised expression of JunB. J. Cell Sci. 116, 4587–4596. 518. Jochum, W., David, J. P., Elliott, C., Wutz, A., Plenk, H., Jr., Matsuo, K., and Wagner, E. F. (2000). Increased bone formation and osteosclerosis in mice overexpressing the transcription factor Fra-1. Nat. Med. 6, 980–984. 519. Kveiborg, M., Sabatakos, G., Chiusaroli, R., Wu, M., Philbrick, W. M., Horne, W. C., and Baron, R. (2004). DeltaFosB induces osteosclerosis and decreases adipogenesis by two independent cell-autonomous mechanisms. Mol. Cell Biol. 24, 2820–2830.

JANE B. LIAN AND GARY S. STEIN 520. Sabatakos, G., Sims, N. A., Chen, J., Aoki, K., Kelz, M. B., Amling, M., Bouali, Y., Mukhopadhyay, K., Ford, K., Nestler, E. J., and Baron, R. (2000). Overexpression of DeltaFosB transcription factor(s) increases bone formation and inhibits adipogenesis. Nat. Med. 6, 985–990. 521. Lee, M. H., Kim, Y. J., Kim, H. J., Park, H. D., Kang, A. R., Kyung, H. M., Sung, J. H., Wozney, J. M., Kim, H. J., and Ryoo, H. M. (2003). BMP-2-induced Runx2 expression is mediated by Dlx5, and TGF-beta 1 opposes the BMP-2-induced osteoblast differentiation by suppression of Dlx5 expression. J. Biol. Chem. 278, 34387–34394. 522. Miyama, K., Yamada, G., Yamamoto, T. S., Takagi, C., Miyado, K., Sakai, M., Ueno, N., and Shibuya, H. (1999). A BMP-inducible gene, dlx5, regulates osteoblast differentiation and mesoderm induction. Dev. Biol. 208, 123–133. 523. Tadic, T., Dodig, M., Erceg, I., Marijanovic, I., Mina, M., Kalajzic, Z., Velonis, D., Kronenberg, M. S., Kosher, R. A., Ferrari, D., and Lichtler, A. C. (2002). Overexpression of Dlx5 in chicken calvarial cells accelerates osteoblastic differentiation. J. Bone Miner. Res. 17, 1008–1014. 524. Tai, G., Polak, J. M., Bishop, A. E., Christodoulou, I., and Buttery, L. D. (2004). Differentiation of osteoblasts from murine embryonic stem cells by overexpression of the transcriptional factor osterix. Tissue Eng. 10, 1456–1466. 525. Yang, X., Matsuda, K., Bialek, P., Jacquot, S., Masuoka, H. C., Schinke, T., Li, L., Brancorsini, S., Sassone-Corsi, P., Townes, T. M., Hanauer, A., and Karsenty, G. (2004). ATF4 is a substrate of RSK2 and an essential regulator of osteoblast biology; implication for CoffinLowry Syndrome. Cell 117, 387–398. 526. Umayahara, Y., Billiard, J., Ji, C., Centrella, M., McCarthy, T. L., and Rotwein, P. (1999). CCAAT/enhancer-binding protein delta is a critical regulator of insulin-like growth factor-I gene transcription in osteoblasts. J. Biol. Chem. 274, 10609–10617. 527. Iyer, V. V., Kadakia, T. B., McCabe, L. R., and Schwartz, R. C. (2004). CCAAT/enhancer-binding protein-beta has a role in osteoblast proliferation and differentiation. Exp. Cell Res. 295, 128–137. 528. Takeshita, S., Namba, N., Zhao, J. J., Jiang, Y., Genant, H. K., Silva, M. J., Brodt, M. D., Helgason, C. D., Kalesnikoff, J., Rauh, M. J., Humphries, R. K., Krystal, G., Teitelbaum, S. L., and Ross, F. P. (2002). SHIP-deficient mice are severely osteoporotic due to increased numbers of hyper-resorptive osteoclasts. Nat. Med. 8, 943–949.

Chapter 15

Signaling in Bone T. John Martin Natalie A. Sims

St. Vincent’s Institute of Medical Research, 9 Princes Street, Fitzroy 3065, Australia University of Melbourne, Department of Melbourne, St. Vincent’s Health, 41 Victoria Pde, Fitzroy 3065, Australia

VIII. RANK Signaling IX. Coupling of Bone Formation to Resorption – Release of Growth Factors from Bone Matrix X. Coupling of Bone Formation to Resorption – Autocrine/Paracrine Regulation by Differentiating Osteoblasts XI. Coupling of Bone Formation to Resorption – Are Osteoclasts a Source of Coupling Activity? References

I. II. III. IV. V.

Abstract Introduction The Control of Osteoclasts Signaling in the Control of Osteoclast Activity Signals from the Osteoblast Lineage that Control Osteoclast Formation VI. Hormone and Cytokine Influences on the Contact-dependent Regulation of Osteoclasts VII. Discovery of the Physiological Signaling Mechanisms in Osteoclast Control

I. ABSTRACT

themselves, growing in the resorption space, can communicate through cell contact and paracrine signaling mechanisms to differentiate. Finally, it is proposed that transiently activated osteoclasts can contribute to the coupling of bone formation to resorption by producing activity that influences preosteoblast participation in bone formation.

The activities of osteoblasts and osteoclasts to form and resorb bone are controlled by circulating hormones and locally generated cytokines and growth factors. The latter are especially important, comprising complex signaling systems in bone that determine how bone is formed and resorbed in the remodeling process. Cells of the osteoblast lineage regulate osteoclast formation from hemopoietic precursors through contact-dependent mechanisms that are controlled by hormones and by the production of locally generated inhibitors. The key effectors are products of the tumor necrosis factor ligand and receptor family. Local signaling that results in bone formation during remodeling takes place in several ways. Growth factors released from resorbed bone matrix can contribute to preosteoblast differentiation and bone formation. The preosteoblasts Dynamics of Bone and Cartilage Metabolism

II. INTRODUCTION Bone formation and resorption proceed throughout life. The processes are more rapid during skeletal growth, at which stage the term modeling is used. Modeling takes place from the beginning of skeletogenesis during fetal life until the end of the second decade when the longitudinal growth of the skeleton is completed. In the modeling process bone is formed at a location different from the sites 259

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260 of resorption, leading to a change in the shape or macroarchitecture of the bone. It is responsible for determining the size and shape of bone, such as the simultaneous widening of long bones and development of the medullary cavity by bone formation at the periosteal surface and resorption at the endosteal surface, respectively. The remodeling process, which continues throughout adult life, is an integral part of the calcium homeostatic system and provides a mechanism for self-repair and adaptation to physical stress. In the adult human skeleton, approximately 5–10% of the existing bone is replaced every year. A characteristic feature of bone remodeling is that the process does not occur uniformly throughout the skeleton. Remodeling of bone occurs in focal or discrete packets known as basic multicellular units (BMUs) of bone turnover. The cellular sequence is always initiated by osteoclastic bone resorption to be followed by osteoblastic new bone formation. This sequence of events is initiated asynchronously throughout the skeleton, at sites that are geographically and chronologically separated from each other. Both bone formation and resorption occur at the same place so that there is no change in the shape of the bone. After a certain amount of bone is removed as a result of osteoclastic resorption and the osteoclasts have moved away from the site, a reversal phase takes place in which a cement line is laid down. Osteoblasts then synthesize matrix, which becomes mineralized. The processes of bone resorption and formation are kept in balance by systemic hormones and cytokines, with the maintenance of a normal, healthy skeletal mass depending on information transfer taking place among osteoblasts, osteoclasts, immune cells, and constituents of the bone matrix. Remodeling thus maintains the mechanical integrity of the skeleton by replacing old bone with new bone [1–5]. The maintenance of adequate trabecular and cortical bone requires that bone formation and resorption should be balanced, such that a high or low level of resorption is usually associated with a similar change in the level of bone formation. The theory that resorption is followed by an equal amount of formation has come to be known as “coupling”. However, during life, the effects of growth and aging, including changes in mechanical stress, mean that this theory of equal bone replacement rarely holds true. During growth there is a positive balance, with the amount of bone replaced at individual BMUs exceeding that lost [3], and with aging there is a negative balance at individual BMUs [6], with gradual attrition of bone. In common metabolic states, such as post-menopausal osteoporosis, while coupling exists and both bone formation and resorption are occurring at a higher level than normal, the amount of bone formed is not equal to that resorbed and bone density is reduced.

T. JOHN MARTIN AND NATALIE A. SIMS

Until the early 1980s it was understood that bone metabolism was regulated by circulating hormones. Parathyroid hormone (PTH) and 1,25(OH)2 vitamin D3, promoted bone resorption, sex steroids had some poorly defined beneficial effects on the skeleton, and it was thought that there must be unknown factors promoting bone formation. The discoveries of subsequent years revealed that, although circulating hormones are important controlling factors, the key influences are locally generated cytokines which influence bone cell function and communication in complex ways, and often are themselves regulated in turn by the hormones. Discoveries of the many intercellular communication and signaling pathways in bone, together with the recent contributions from mouse and human genetics, have contributed enormously to the understanding of bone physiology and pathology, and have provided many new approaches to the development of drugs to prevent and treat bone diseases. Indeed, many cytokines that had originally been discovered by virtue of their actions on the immune or hematopoietic systems have been revealed as fundamental local factors in the control of bone cell function. This is exemplified most directly and simply by the remarkable number of skeletal phenotypes that exist in genetically altered mice, which either under- or overexpress these cytokines or their receptors.

III. THE CONTROL OF OSTEOCLASTS Osteoclasts are the only cells that can resorb bone. Their formation and activity are controlled by signals emanating predominantly from cells of the osteoblast lineage, but also from T and B cells. Further to this, in disease states other cell types have been shown to support osteoclast formation, such as breast cancer cells and synovial fibroblasts in rheumatoid arthritis. The proposal that osteoblasts mediate hormonal stimulation of osteoclast formation and activity [7] led eventually to the identification of the molecules responsible for physiological control of osteoclast formation and activity by osteoblasts. In addition to these communication pathways directing the osteoclast lineage, it has long been proposed that a local “coupling factor” plays a key role in bone remodeling by linking bone resorption to subsequent formation. Although such a coupling factor has never been identified, there have been suggestions that such activity is released from bone matrix during resorption [8, 9]. Advances of the last few years allow consideration of other contributing factors, including the possibility that signaling from the activated osteoclast could play a part in the process, helping to stimulate osteoblast lineage cells to replace the resorbed bone in each BMU. In this way,

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Chapter 15 Signaling in Bone

intercellular communication appears to exist in both directions between osteoblasts and osteoclasts.

IV. SIGNALING IN THE CONTROL OF OSTEOCLAST ACTIVITY The first indications of the importance of intercellular signaling in bone came in the early 1980s, in that, when osteoclasts were isolated from newborn rat or mouse bone, they required the presence of contaminating osteoblastic cells in order to be fully active and resorb bone [10, 11]. The observations that isolated osteoblasts of various origins responded to bone-resorbing hormones and possessed receptors for these factors, in addition to the lack of evidence demonstrating receptors or direct responses to these hormones in osteoclasts, led to the concept that bone-resorbing factors must act first on osteoblasts, most likely bone-lining cells. This was proposed to release factors that influence the bone-resorbing activity of osteoclasts [7, 12]. Furthermore, since osteoclasts are derived from hemopoietic progenitors and not from a local bone cell, Chambers came to the same conclusion by arguing that, since the osteoclast derives from a “wandering” cell, it made sense to have its activity programmed by an authentic bone cell, i.e. the osteoblast [13]. Work over the next few years established the concept of an osteoblast-derived stimulator of osteoclast resorption [10, 14, 15] and by the mid-1980s the field was at the stage of accepting that intercellular signaling may be an important mechanism in bone cell biology. Among the questions raised were whether a single stromal/osteoblastic cell factor existed that is responsible for osteoclast activation, and if so, is it cell-associated or secreted? Would the formation of osteoclasts be at least as important as regulation of their activity? The next real advances came with the development of methods to study osteoclast formation in vitro.

V. SIGNALS FROM THE OSTEOBLAST LINEAGE THAT CONTROL OSTEOCLAST FORMATION Major advances required the development of murine bone marrow cultures, with reproducible assays of osteoclast formation [16–19]. Takahashi observed that more than 90% of the TRAP-positive mononucleated cell clusters and multinucleated cells formed in mouse marrow cultures, in response to bone-resorbing stimuli, were located near colonies of alkaline phosphatase-positive mononucleated

cells (possibly osteoblasts) [18]. This suggested that osteoblastic cells are involved in osteoclast formation, in addition to the evidence produced in the few earlier years of their influence on osteoclast activity. Proof of this came from the experimental system established by Takahashi [16] in which osteoclasts were formed from osteoblast-rich cultures from newborn mouse calvariae grown in co-culture with mouse spleen cells treated with 1,25(OH)2D3. Importantly, when the osteoblastic cells were separated from the spleen cells by a 0.45−µm membrane filter, but shared the same media, no osteoclast formation occurred. Similar results were obtained with bone marrow-derived stromal cell lines including MC3T3-G2/PA6, ST2, and KS-4 [20, 21]. Thus, it was clear that direct cell-to-cell contact of osteoclast precursors with osteoblasts, or their precursors, is necessary for osteoclast differentiation.

VI. HORMONE AND CYTOKINE INFLUENCES ON THE CONTACTDEPENDENT REGULATION OF OSTEOCLASTS With increasing acceptance of a role for cells of the osteoblast lineage in controlling osteoclast formation and activity by a contact-dependent mechanism, it was important to understand how this process was regulated by hormones and cytokines that stimulate osteoclast formation. Prostaglandin (PG)-induced osteoclast formation in mouse bone marrow cultures was mediated by cAMP [22]. Likewise, PTH and PTHrP, acting through their common receptor, promoted osteoclast formation in marrow cultures by a cAMP-dependent mechanism [23, 24], and the effect of interleukin-1 (IL-1) resulted from the generation of PGE2 as an intermediate effector [25]. There remains no convincing evidence for the existence of functional PTH/ PTHrP receptors in osteoclasts, capable of supporting a direct stimulatory effect of PTH on the osteoclast. A second signaling mechanism for regulation of osteoclastogenesis by osteoblasts was provided by the steroid hormone, 1,25(OH)2D3 which had very similar effects on osteoclast formation in marrow cultures and in co-cultures of osteoblasts with hemopoietic cells [17]. 1,25(OH)2D3 signals by combining with its receptor and translocating to the nucleus to influence transcriptional events. Finally, a membrane-bound receptor complex involving a 130 kDa glycoprotein (gp130) [26] provides for osteoclast formation under the influence of the group of cytokines that use this signaling mechanism. In co-cultures of mouse stromal cells with hemopoietic cells, simultaneous

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treatment with IL-6 and its soluble receptor (sIL-6R) induced osteoclast formation, but when added separately they were ineffective [26]. The other cytokines in this group, IL-11, leukemia inhibitory factor (LIF), and oncostatin M (OSM), all of which use gp130 as a common transducer, also stimulated osteoclast formation [26]. When cells from IL-6R-overexpressing transgenic mice were used in crossover co-cultures with hemopoietic cells from wildtype mice, the expression of IL-6R by osteoblastic cells was shown to be indispensable for the induction of osteoclasts [27]. This clearly demonstrated that stimulation of osteoclast formation by IL-6 required action of the cytokine on the osteoblast, despite the fact that osteoclasts possessed its receptor. A central role of the gp130-coupled cytokines in osteoclast development was further suggested by the observations that PTH, 1, 25(OH)2D3 , PGE2 , IL-1, and TNFα all promoted IL-11 production by osteoblastic cells [28]. Furthermore, addition of neutralizing anti-gp130 to co-cultures fully blocked stimulation of osteoclast formation in response to IL-1, and partly blocked the responses to PTH, 1,25(OH)2D3 , and PGE2. Thus the concept of stromal/osteoblastic regulation of osteoclastogenesis through local signaling mechanisms was firmly established, along with its regulation by a number of circulating and local factors. Despite the fact that they fell into three main classes with respect to their initial signaling mechanisms (Fig. 1), it seemed that a common pathway for these agents was the membrane stromal factor called variously “stromal osteoclast-forming activity” (SOFA) [29] or “osteoclast differentiation factor” (ODF) [19]. It was assumed that these agents must converge in their actions at some stage before finally generating the crucial membrane factor [30, 31].

VII. DISCOVERY OF THE PHYSIOLOGICAL SIGNALING MECHANISMS IN OSTEOCLAST CONTROL Of the many multifunctional cytokines that had some role in osteoclast formation, none provided an explanation for the molecular regulation of osteoclast formation and activity that was evident from the foregoing studies. In a form of murine osteopetrosis resulting from a mutation in the coding region of the M-CSF gene in the op/op mouse [32, 33], M-CSF was found to play a role in both proliferation and differentiation of osteoclast progenitors [34]. On the other hand, M-CSF inhibited the bone-resorbing activity of isolated osteoclasts [35], and osteoclasts were found to be rich in M-CSF receptors [36]. Bone resorption in organ culture was reduced by M-CSF, GM-CSF, and IL-3 [37], and all three cytokines inhibited osteoclast generation in mouse bone marrow cultures. The conclusion from these and other observations was that none of these hemopoietic growth factors fulfilled criteria expected of one that is specific for osteoclast formation, and certainly not the predicted ODF/SOFA. The hypothesis was compelling, but, by the mid-1990s there had been no significant advances towards identifying these control mechanisms. Resolution of the question came in 1997 with the discovery by two groups independently that osteoclast formation is controlled physiologically by regulated interactions among members of the TNF ligand and receptor families. The discovery of osteoprotegerin (OPG), a soluble member of the TNF receptor superfamily, revealed it as a powerful inhibitor of osteoclast formation [38, 39].

Figure 1 Three distinct signaling pathways in cells of the osteoblast lineage result in osteoclast formation, through a mechanism depending on contact between the osteoblastic cells and the hemopoietic precursors of osteoclasts. The membrane protein responsible for promoting osteoclast differentiation was discovered to be RANKL, binding to its receptor, RANK. The process is inhibited by the decoy receptor, OPG (not shown).

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This provided the means of identifying and cloning the elusive ODF/SOFA, which proved to be a TNF ligand family member that came to be called receptor activator of nuclear factor κB ligand (RANKL), as the common factor mediating osteoclast formation in response to all known stimuli [40, 41]. As a membrane protein, RANKL fulfilled the predictions of earlier work, that osteoclast differentiation required contact-dependent activation of hemopoietic precursors. The communication with the hemopoietic lineage results from RANKL binding to its receptor on the osteoclast lineage, RANK, thereby initiating signaling essential for osteoclast differentiation. Figure 1 illustrates the regulated production of RANKL that is essential for osteoclast formation and maintenance of activity. The boneresorbing cytokines and hormones, with disparate signaling mechanisms, converge in promoting RANKL production. The decoy receptor, OPG, has an essential physiological role as a paracrine regulator of osteoclast formation, produced by the osteoblasts and binding RANKL to limit its activation of osteoclast formation through its receptor, RANK. All of these discoveries have been validated by studies in genetically altered mice, establishing clearly the essential physiological role of these TNF ligand and receptor family members in controlling osteoclast formation and activity. First, transgenic overexpression of OPG in mice results in generation of mice with osteopetrosis because of failure to form osteoclasts [38]. Genetic ablation of OPG, on the other hand, leads to severe high turnover osteoporosis [42]. Removal of the stimulatory pathway by genetic ablation of RANKL also results in osteopetrosis because RANKL is necessary for normal osteoclast formation [43]. Finally, genetic ablation of the hemopoietic cell receptor, RANK, also leads to osteopetrosis [44]. Because this signaling pathway is functional also in immune cells, RANK-null mice have severe abnormalities in that system, reflecting an intriguing link with the immune system, which was evident very early in the discovery process. The role of OPG as a paracrine effector has been illustrated by in vitro experiments that lead to the conclusion that constitutive production of this decoy receptor is necessary to limit the osteoclast formation resulting from RANKL stimulation [45]. Two other groups were successful in identifying and cloning RANKL, but these groups were interested in its role in the immune response [46, 47]. Wong et al. [46] identified and characterized a TNF-related activationinduced cytokine (TRANCE) during a search for apoptosisregulatory genes in murine T-cell hybridomas, finding it to be predominantly expressed on T cells and in lymphoid organs and controlled by the T-cell receptor through a calcineurin-regulated pathway. The putative receptor for

TRANCE was detected on mature dendritic cells [46, 48]. In studying the processing and presentation of antigens by dendritic cells to T cells, Anderson et al. [47] characterized receptor activator of NF-κB (RANK), a new member of the TNF receptor family derived from dendritic cells, and its ligand RANKL, which they recognized to be identical to TRANCE [46]. Furthermore, production of soluble RANKL by activated T cells directly stimulated osteoclast formation in vitro and in vivo [49], with important pathophysiological implications. It is apparent that lymphocytes exert important regulatory controls on osteoclast function. Physiological control of bone resorption is thus dependent on the essential regulatory function of RANKL, not only in promoting osteoclast formation, but also their survival and activity [50], as predicted from the earlier demonstration of osteoclast activation through contact with osteoblastic cells [10, 51]. By treating with RANKL and M-CSF it became possible to prepare osteoclasts in relatively large numbers without the participation of stromal/osteoblastic precursors, including the preparation of human osteoclasts from peripheral blood [52, 53]. As predicted, the important osteotropic hormones and cytokines, including 1,25(OH)2D3, IL-11, PTH, and PGE2, stimulated osteoblast RANKL production, triggering osteoclast development, and the same treatments reduced production of OPG. The physiological function of OPG as a paracrine regulator, suggested by the phenotypes of OPG-null and OPG-transgenic mice, was demonstrated using in vitro organ and cell culture methods also [45]. The functions of RANKL extend to pathological states of increased bone resorption, where increased local RANKL production contributes to bone destruction in disease states such as breast cancer metastases to bone [54] in rheumatoid arthritis [49], multiple myeloma [55], and osteoporosis [56].

VIII. RANK SIGNALING The activation of RANK by its ligand initiates signaling events in cells of the monocyte–macrophage series that result in the expression of genes required for osteoclast differentiation. The key initial step is binding of the intracellular domain of RANK to TNFR-associated cytoplasmic factors (TRAFs). The C-terminal region within residues 544 to 616 is required for binding to TRAF 1, 2, 3, 5, and 6 [57, 58], and there is an additional separate binding site for TRAF 6 at amino acids 340–421 [58]. RANK signaling was found to activate both NF-κB and JNK activity, and deletion of the 72 C-terminal residues of RANK resulted

264 in a major reduction in NF-κB and JNK activity [57, 58]. Further ablation of the TRAF 6-binding region ablated all residual NF-κB and JNK activity [58]. Mutation of the TRAF 6-binding site in RANK completely inhibited NF-κB activation, while JNK activation was inhibited to a lesser extent. It was concluded that binding of RANK to TRAF 6 is pivotal for NF-κB activation, and the TRAF 2-binding site was subsequently shown to be required for JNK activation [57]. Earlier work had shown that inactivating mutations of both the p50 and p52 components of NF-κB in mice resulted in osteopetrosis because of failed osteoclast development [59]. In keeping with the importance of NF-κB signaling in osteoclast formation, mice rendered null for the TRAF 6 gene have osteopetrosis [60, 61], consistent with a critical role for RANK/TRAF 6 in the activation of osteoclastic bone resorption. A novel aspect of signaling in osteoclast development is the ability of RANKL to limit its own osteoclastogenic effect by promoting interferon-β (IFN-β) production by monocytic osteoclast precursors [62]. This inhibits osteoclast formation by preventing RANKL-induced expression of c-fos. The latter was known as an essential transcription factor in osteoclast differentiation, since c-fos-null mice were osteopetrotic because of failed osteoclast formation [63]. Participation by IFN-β in this internal control process provides another mechanism of local signaling that could regulate bone remodeling. The extent of osteoclastic resorption within individual BMUs needs to be limited. If it were to continue unchecked, resorption would be excessive. RANKL-induced IFN-β provides an appealing mechanism, contained within the osteoclast itself, by which osteoclast formation could be controlled at critical sites in normal bone remodeling. Further elucidation of the RANK signaling pathway came with the identification that RANKL treatment considerably enhanced the expression and nuclear location of nuclear activator of activated T cells (NFAT)-2 (NFATc1), a transcription factor regulated by calcineurin and calcium [64]. It was shown by several methods that NFATc1 activation is necessary and sufficient for osteoclast differentiation. NFATc1 up-regulation did not occur in either c-fos or TRAF 6-null mice, suggesting that both pathways are necessary for up-regulation of NFATc1 by RANKL. A further, novel aspect of RANKL signaling emerged in this work with the demonstration that RANKL-induced oscillations in cytoplasmic calcium in osteoclast precursors were necessary for its effect. Both calcium chelators and calcineurin inhibitors suppressed NFATc1 expression and nuclear translocation. These discoveries provided the first real link between the observations of failed osteoclast development in mice deficient in RANK signaling and c-fos-null mice.

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IX. COUPLING OF BONE FORMATION TO RESORPTION – RELEASE OF GROWTH FACTORS FROM BONE MATRIX While much is known about the communication processes among cells of bone that determine osteoclast formation and activity, less is understood of the signaling processes that govern osteoblast differentiation and bone formation. In particular it is unknown how the extent of bone formation in response to resorption is controlled, and how the two processes are so closely linked. Interventions that increase bone resorption are usually accompanied by increased bone formation. A recent example of this comes from observations in mice overexpressing TGFβ in osteoblasts, that had increased bone resorption and formation, with a net bone loss [65]. By crossing these mice with other genetically manipulated mice these authors showed that the increased formation was dependent upon osteoclastic resorption and was not a direct effect of TGF. The very nature of the remodeling process, occurring as it does in different parts of the skeleton at different times, highlights the importance of locally generated and regulated factors in the process. When the cycle is initiated, say by PTH or by mechanical strain which would generate cytokines or prostanoids [66], the thin layer of nonmineralized matrix under these cells is initially digested by collagenase to expose the mineralized matrix which osteoclasts can resorb [67]. PTH stimulates collagenase production and secretion in osteoblastic cells [68]; in mice whose collagenase is unable to digest type I collagen, the bone resorbing action of PTH is severely attenuated [69]. According to this model of initiation of resorption, osteoclasts or their late precursors near to final maturation and activation must be available close to the sites where they are required. Although this is likely, there is no direct in vivo evidence for it. Toward the end of resorption, mononuclear cells are seen at the bottom of resorption pits, where they remove demineralized collagen. The mononuclear cells ascribed this function have sometimes been fibroblasts, but more commonly macrophages. Recent work, applying light and electron microscopy to a number of in vitro models, describes how bone-lining cells, through the mediation of matrix metalloproteinases, carry out the task of cleaning resorption pits after osteoclasts have completed their task [70]. In order to do this the cells must use differentiated properties that they acquire and retain as part of the osteoblast lineage. These include especially the capacity to respond appropriately to the cytokines and growth factors that most likely govern these steps. A favored hypothesis for some years is that this “coupling” is regulated by bone formation factors released

Chapter 15 Signaling in Bone

from the bone matrix during bone resorption [9, 10]. Indeed, a large number of substances which are mitogenic to osteoblasts or which stimulate bone formation in vivo can be extracted from bone matrix [71]. These include insulin-like growth factor (IGF) I and II [72], acidic and basic fibroblast growth factor (FGF) [73], transforming growth factor ß (TGFß) 1 and 2 [74], and TGFß heterodimers [75], bone morphogenetic proteins (BMPs) 2, 3, 4, 6, and 7 [76–78], platelet-derived growth factor (PDGF) [79], and probably others. Several questions should be considered regarding the role of these substances in the coupling of bone formation to bone resorption: (1) which cells produce them and under what circumstances?; (2) do they stimulate bone formation in vivo?; (3) can they be released from the matrix in active form and in controlled amounts during bone resorption?; (4) is there evidence for an increase in the abundance of these substances at sites of bone remodeling?; and (5) are there regulated mechanisms by which they are activated? IGF-I, IGF-II, bFGF, TGFβ, and PDGF are produced by rat osteoblastic cells. IGF-I and IGF-II production is enhanced by stimulators of bone formation, such as PGE2 and PTH [80, 81]. Elevated levels of IGF-I mRNA were found in bone from estrogen-deficient rats, where bone turnover is increased [82, 83]. During bone growth in rats, there is a close association between osteogenesis and IGF-I expression [83]. However, following marrow ablation which causes a substantial increase in bone formation, the rise in IGF-I mRNA was seen after the appearance of differentiated osteoblasts, suggesting that it did not initiate bone formation in that system [84]. In human bone, the major form of IGF is IGF-II, which is also produced by human bone cells in culture [9, 10]. Bone is one of the most abundant sources of TGFβ [85], and this growth factor is produced by all osteoblastic cells examined. The BMPs are members of the TGFβ superfamily. BMP-2 and BMP-4 are produced in adult bovine pre-odontoblasts [86] and human fetal teeth [87]. BMP-7 (OP-1) was localized in human embryos in hypertrophic chondrocytes, osteoblasts, and periosteum as well as other tissues [77], while BMP-3 was found in human embryonic lung and kidney in addition to perichondrium, periosteum, and osteoblasts [78]. Both bFGF [88] and PDGF [79] were shown to be produced by bone cells or bone explants in culture. These factors could thus be involved in bone remodeling but the time and site for their synthesis and secretion in vivo has not yet been determined. Prostaglandin E, primarily E2, is another bone cell-produced cytokine, which is up-regulated by mechanical strain in vitro [89] and stimulates both bone resorption and formation [90]. Many growth factors stimulate bone formation in vivo. IGF-I, injected into humans or rats, increases both bone

265 resorption and bone formation [91], yet reports on its relative effects on trabecular and cortical bone are inconclusive [92]. When injected together with the IGF-binding protein IGFBP-3 into rats, it was reported to increase bone volume [93]. BMPs injected into bone stimulate bone formation locally and produce a positive bone balance. TGFβ, from the same family of proteins, has a similar effect. When injected next to the periosteum or endosteum, local bone formation is substantially augmented in rats and other species [94, 95]. At the same time, endocortical bone resorption is increased; thus, like IGF-I, TGFβ seems to stimulate both resorption and formation, however, the local balance is clearly positive. bFGF, injected both locally and systemically, was also reported to increase bone formation [96]. PGE1 and PGE2 have long been known to be potent stimulators of bone formation when given either locally or systemically, both to humans or experimental animals [97, 98]. These substances could thus contribute to the bone formation observed in remodeling, if secreted or released in active form at the appropriate site and time. It was proposed that TGFß, which is produced as an inactive precursor in bone and bone cells is present in the matrix and can be activated by acidification or proteolytic cleavage by resorbing osteoclasts [99]. Consistent with this, TGFß activity was recovered from conditioned media of in vitro resorbing osteoclasts [100]. It remains to be shown if the other growth factors also survive the proteolytic cleavage of the acidic hydrolases present in resorption lacunae. Other questions raised by this model of coupling, via growth factor release from the matrix, relate to the time course and the distance between the resorption and formation processes and whether activation can be controlled with sufficient precision in this way. Osteoclastic bone resorption proceeds for about 2 weeks before formation follows and continues for 3–4 months. The osteoblast precursors, which should respond to the “coupling factors” are many microns away from where active osteoclast resorption is in progress. The osteoblastic lineage cells produce TGFβ in latent form and the IGFs as complexes bound to a family of specific, high-affinity binding proteins (IGFBPs), which regulate their bioavailability [101]. TGFβ may be released from latent complexes at appropriate sites in bone by plasmin generated locally by plasminogen activators, in a manner controlled temporally and spatially by hormones and cytokines [102]. A similar local control could free IGF-1 from association with its inhibitory binding protein [103]. Although there is no obvious skeletal phenotype in mice with inactivated genes for plasminogen activators, in vivo investigation of such possibilities would require treatment of such animals with anabolic agents such as PTH.

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X. COUPLING OF BONE FORMATION TO RESORPTION – AUTOCRINE/PARACRINE REGULATION BY DIFFERENTIATING OSTEOBLASTS The above theory of coupling requires dissolution of growth factors from matrix and their activation by acidification and/or proteases. An extension of this that is not necessarily exclusive arises from the work of Boyde and colleagues [104], in which they showed in vitro that rat calvarial cells grown on bone slices with mechanically excavated crevices and grooves made bone in those defects, filling them exactly to a flat surface. The findings suggested that it is the topography of the bone that determines the timing, siting, and extent of new bone formation, and that in vivo this would take place in the resorbed spaces vacated by osteoclasts. Both the proposed growth factor involvement and the work of Gray et al. [104] imply that, once the formation process is established, the participating cells themselves are able to sense spatial limits, and most likely do so by chemical communication that takes place between the developing osteoblasts. The likely mediators of these signaling processes are the same growth factors and cytokines that are proposed to be of matrix origin.

XI. COUPLING OF BONE FORMATION TO RESORPTION – ARE OSTEOCLASTS A SOURCE OF COUPLING ACTIVITY? Observations in genetically manipulated mice suggest that the osteoclast itself could also be the source of an activity that contributes to the fine control of the coupling process. Generation of coupling activity was suggested by increased bone formation in OPG−/− mice [105], which are severely osteoporotic because of excessive osteoclast formation. In bone sections from mice in this high bone-turnover state sites of active bone resorption very commonly had active osteoblasts located nearby, suggesting that the coupling activity in this high-turnover state was more likely derived from osteoclasts themselves. Some indication of an osteoclast role comes also from human genetics. In individuals with the osteopetrotic syndrome, ADOII, due to inactivating mutations in the chloride-7 channel (ClC-7), bone resorption is deficient because of the failure of the osteoclast acidification process. Bone formation in these patients is nevertheless normal, rather than diminished as might be expected because of the greatly impaired resorption [106]. Furthermore, in mice deficient in either c-src [107], ClC-7 [108], or tyrosine phosphatase epsilon [109], bone resorption is inhibited

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without inhibition of formation. In these three knockout mouse lines osteoclast resorption is greatly reduced by the mutation, although osteoclast numbers are not reduced. Indeed osteoclast numbers are actually increased because of reduced osteoclast apoptosis. A possibility is that these osteoclasts, although unable to resorb bone, are nevertheless capable of generating a factor (or factors) required for bone formation. On the other hand, mice lacking c-fos, which are unable to generate osteoclasts, have reduced bone formation as well as resorption [63]. The cytokines that signal through gp130 play an important role in intercellular communication processes in bone, with evidence indicating that they can be involved in regulation of both bone resorption and formation [28, 110]. We addressed this by studying mice in which each of the two gp130-dependent signaling pathways was specifically attenuated, and found that inactivation of the SHP2/ras/ MAPK signaling pathway (gp130Y757F/Y757F mice) yielded mice with greater osteoclast numbers and bone resorption, as well as greater bone formation than wild-type mice. This increased bone remodeling resulted in less bone because the increase in resorption was relatively greater than that in formation. In other words the coupling process was imprecise in a way that resembles the result of estrogen withdrawal, as in ovariectomy. gp130Y757F/Y757F mice crossed with IL-6-null mice had similarly high osteoclast numbers and increased bone resorption, however these mice showed no corresponding increase in bone formation and thus had extremely low bone mass. Thus resorption alone is insufficient to promote the coupled bone formation, but the active osteoclasts are the likely source. Furthermore this indicated that stimulation of bone formation coupled to the high level of bone resorption in gp130Y757F/Y757F mice is an IL-6-dependent process, though it does not necessarily show that it is mediated by IL-6 itself [111]. Evidence of quite a different nature points further to a role for the activated osteoclast in the coupling process. Mice rendered null for calcitonin (CT−/− mice) have increased bone formation [112], as do those deficient in the CT receptor (CTR+/− mice) [113]. These unexpected findings can be explained by the hypothesized role of active osteoclasts. The best documented action of calcitonin is its acute inhibition of osteoclast function following injection [114]. If transient osteoclast activation is necessary for bone formation, then removal of calcitonin would result in osteoclasts remaining active in a way that allows them to continue contributing to bone formation, thus giving rise to the phenotype expressed in CT−/− and in CTR+/− mice. The real physiological function of CT has never been clearly understood, but these new observations, together with what is proposed, could reveal it as a short-term regulator of osteoclast activity which

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Figure 2 Three main pathways contributing to the process of coupling of bone formation to resorption. 1. Osteoclasts resorb matrix, releasing stored growth factors that are able to promote osteoblast precursor division or maturation [9]. 2. Preosteoblasts in the resorption space divide, communicate with each other through gap junctions and paracrine signals to program differentiation and sense spatial requirements [104]. 3. Osteoclasts, activated by RANKL generated in osteoblasts, release coupling activity that is required for the osteoblast response [123].

through its action on the osteoclast also regulates bone formation. Finally, the finding that effective inhibition of resorption markedly attenuates, or even abolishes, the anabolic effect of PTH, also points to a role for the osteoclast. The anabolic effect of PTH is lost in c-fos−/− mice that are osteopetrotic because of failure of osteoclast formation [115]. Treatment of patients with osteoporosis concomitantly with PTH and a bisphosphonate resulted in significant early blunting of the anabolic response to PTH [116, 117], with the obvious implication that combining the anabolic PTH with antiresorptive bisphosphonate would be contraindicated [118, 119]. If some aspect of resorption is required for the anabolic effect of PTH, how is that connection made? Treatment of rats with a single subcutaneous injection of PTH results in a transient increase in mRNA for RANKL and a decrease in that for OPG, with maximum effect at 1 hour, and returning to control within 2 hours, leading us to suggest that a subtle or transient increase in osteoclast formation or activation might be needed to prepare the bone surface for new matrix deposition [120, 121]. Finally and most importantly, Holtrop et al. [122] showed that intravenous injection of PTH in young rats resulted in transient activation of osteoclasts in vivo, evident within 30 minutes, and followed only some hours later at high PTH doses by increased osteoclast number. Thus it is clear that rapid activation of osteoclasts in response to a single injection of PTH can occur, and it is likely that this is the consequence of a transient increase in RANKL production by cells at sites available for activation of BMUs. Our interpretation of the foregoing and other data is that what is needed for full expression of the anabolic action of PTH, in addition to its direct effects on committed

preosteoblasts, is a transient effect on the osteoclast, achieved by promoting activation, rather than formation, of osteoclasts [123]. The precise way in which the osteoclast is involved in the anabolic process needs to be clearly understood because of the implications for sequential or combined use of therapeutic resorption inhibitors and anabolic agents, and for the development of new anabolic agents. Figure 2 illustrates this schematically, with the hypothesis that there can be at least three main pathways involved in the coupling of bone formation to resorption. The first is the release of any of several growth factors from bone matrix as a result of resorption. The second is the communication among osteoblast precursors, through gap junctions and growth factors, that equips them to sense the space that needs to be filled after resorption. The third is the postulated production of coupling activity from activated osteoclasts, a regulated process that could depend on the controlled generation of RANKL and transient osteoclast activation.

Acknowledgement Work from the authors’ laboratories supported by the National Health & Medical Research Council of Australia.

References 1. Frost, H. M. (1964). Dynamics of bone remodeling. In “Bone Biodynamics” (H. M. Frost, ed.), pp. 315–333. Little, Brown & Co., Boston, MA. 2. Harris, W. H., and Heaney, R. P. (1969). Skeletal renewal and metabolic bone disease. N. Engl. J. Med. 280, 193–202.

268 3. Parfitt, A. M. (1982). The coupling of bone formation to bone resorption: a critical analysis of the concept and of its relevance to the pathogenesis of osteoporosis. Metab. Bone Dis. Relat. Res. 4, 1–6. 4. Eriksen, E. F. (1986). Normal and pathological remodeling of human trabecular bone: three dimensional reconstruction of the remodeling sequence in normals and in metabolic bone disease. Endocr. Rev. 7, 379–408. 5. Martin, T. J., and Rodan, G. A. (2001). Coupling of bone resorption and formation during bone remodeling. In “Osteoporosis” (J., Kelsey, ed.), 2nd ed., pp. 361–372. Academic Press, San Diego, CA. 6. Lips, P., Courpron, P., and Meunier, P. J. (1978). Mean wall thickness of trabecular bone packets in the human iliac crest: changes with age. Calcif. Tissue Res. 26, 13–17. 7. Rodan, G. A., and Martin, T. J. (1981). Role of osteoblasts in hormonal control of bone resorption – a hypothesis. Calcif. Tissue Int. 33, 349–351. 8. Howard, G. A., Bottemiller, B. L., Turner, R. T., Rader, J. I., and Baylink, D. J. (1981). Parathyroid hormone stimulates bone formation and resorption in organ culture: evidence for a coupling mechanism. Proc. Natl Acad. Sci. USA 78, 3204–3208. 9. Mohan, S., and Baylink, D. J. (1991). The role of IGF-II in the coupling of bone formation to resorption. In “Modern Concepts of Insulin-like Growth Factors” (E. M. Spencer, ed.), pp. 19–174. Elsevier, New York. 10. Chambers, T. J., and Magnus, C. J. (1982). Calcitonin alters behaviour of isolated osteoclasts. J. Pathol. 136, 27–39. 11. Chambers, T. J. (1985). The pathobiology of the osteoclast. J. Clin. Pathol. 38, 241–252. 12. Martin, T. J., Partridge, N. C., Greaves, M., Atkins, D., and Ibbotson, K. J. (1979). Prostaglandin effects on bone and role in cancer hypercalcemia. In “Molecular Endocrinology” (M. Szelke, ed.), pp. 251–264. Elsevier, Amsterdam. 13. Chambers, T. J. (1980). The cellular basis of bone resorption. Clin. Orthop. 283–293. 14. McSheehy, P. M., and Chambers, T. J. (1986). Osteoblast-like cells in the presence of parathyroid hormone release soluble factor that stimulates osteoclastic bone resorption. Endocrinology 119, 1654–1659. 15. McSheehy, P. M., and Chambers, T. J. (1986). Osteoblastic cells mediate osteoclastic responsiveness to parathyroid hormone. Endocrinology 118, 824–828. 16. Takahashi, N., Akatsu, T., Udagawa, N., Sasaki, T., Yamaguchi, A., Moseley, J. M., Martin, T. J., and Suda, T. (1988). Osteoblastic cells are involved in osteoclast formation. Endocrinology 123, 2600–2602. 17. Takahashi, N., Akatsu, T., Sasaki, T., Nicholson, G. C., Moseley, J. M., Martin, T. J., and Suda, T. (1988). Induction of calcitonin receptors by 1 alpha, 25-dihydroxyvitamin D3 in osteoclast-like multinucleated cells formed from mouse bone marrow cells. Endocrinology 123, 1504–1510. 18. Takahashi, N., Yamana, H., Yoshiki, S., Roodman, G. D., Mundy, G. R., Jones, S. J., Boyde, A., and Suda, T. (1988). Osteoclast-like cell formation and its regulation by osteotropic hormones in mouse bone marrow cultures. Endocrinology 122, 1373–1382. 19. Suda, T., Takahashi, N., and Martin, T. J. (1992). Modulation of osteoclast differentiation. Endocr. Rev. 13, 66–80. 20. Udagawa, N., Takahashi, N., Akatsu, T., Sasaki, T., Yamaguchi, A., Kodama, H., Martin, T. J., and Suda, T. (1989). The bone marrow-derived stromal cell lines MC3T3-G2/PA6 and ST2 support osteoclast-like cell differentiation in cocultures with mouse spleen cells. Endocrinology 125, 1805–1813. 21. Yamashita, T., Asano, K., Takahashi, N., Akatsu, T., Udagawa, N., Sasaki, T., Martin, T. J., and Suda, T. (1990). Cloning of an osteoblastic cell line involved in the formation of osteoclast-like cells. J. Cell. Physiol. 145, 587–595. 22. Akatsu, T., Takahashi, N., Debari, K., Morita, I., Murota, S., Nagata, N., Takatani, O., and Suda, T. (1989). Prostaglandins promote osteoclastlike

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23.

24.

25.

26.

27.

28.

29.

30.

31.

32.

33.

34.

35.

36.

37.

38.

cell formation by a mechanism involving cyclic adenosine 3′, 5′monophosphate in mouse bone marrow cell cultures. J. Bone. Miner. Res. 4, 29–35. Akatsu, T., Takahashi, N., Udagawa, N., Sato, K., Nagata, N., Moseley, J. M., Martin, T. J., and Suda, T. (1989). Parathyroid hormone (PTH)-related protein is a potent stimulator of osteoclast-like multinucleated cell formation to the same extent as PTH in mouse marrow cultures. Endocrinology 125, 20–27. Evely, R. S., Bonomo, A., Schneider, H. G., Moseley, J. M., Gallagher, J., and Martin, T. J. (1991). Structural requirements for the action of parathyroid hormone-related protein (PTHrP) on bone resorption by isolated osteoclasts. J. Bone Miner. Res. 6, 85–93. Akatsu, T., Takahashi, N., Udagawa, N., Imamura, K., Yamaguchi, A., Sato, K., Nagata, N., and Suda, T. (1991). Role of prostaglandins in interleukin-1-induced bone resorption in mice in vitro. J. Bone Miner. Res. 6, 183–189. Tamura, T., Udagawa, N., Takahashi, N., Miyaura, C., Tanaka, S., Yamada, Y., Koishihara, Y., Ohsugi, Y., Kumaki, K., Taga, T., Kishimoto, T., and Suda, T. (1993). Soluble interleukin-6 receptor triggers osteoclast formation by interleukin 6. Proc. Natl Acad. Sci. USA 90, 11924–11928. Udagawa, N., Takahashi, N., Katagiri, T., Tamura, T., Wada, S., Findlay, D. M., Martin, T. J., Hirota, H., Taga, T., Kishimoto, T., and Suda, T. (1995). Interleukin (IL)-6 induction of osteoclast differentiation depends on IL-6 receptors expressed on osteoblastic cells but not on osteoclast progenitors. J. Exp. Med. 182, 1461–1468. Romas, E., Udagawa, N., Zhou, H., Tamura, T., Saito, M., Taga, T., Hilton, D. J., Suda, T., Ng, K. W., and Martin, T. J. (1996). The role of gp130-mediated signals in osteoclast development: regulation of interleukin 11 production by osteoblasts and distribution of its receptor in bone marrow cultures. J. Exp. Med. 183, 2581–2591. Chambers, T. J., Owens, J. M., Hattersley, G., Jat, P. S., and Noble, M. D. (1993). Generation of osteoclast-inductive and osteoclastogenic cell lines from the H-2KbtsA58 transgenic mouse. Proc. Natl Acad. Sci. USA 90, 5578–5582. Martin, T. J., and Ng, K. W. (1994). Mechanisms by which cells of the osteoblast lineage control osteoclast formation and activity. J. Cell Biochem. 56, 357–366. Suda, T., Udagawa, N., Nakamura, I., Miyaura, C., and Takahashi, N. (1995). Modulation of osteoclast differentiation by local factors. Bone 17, 87S-91S. Felix, R., Cecchini, M. G., and Fleisch, H. (1990). Macrophage colony stimulating factor restores in vivo bone resorption in the op/op osteopetrotic mouse. Endocrinology 127, 2592–2594. Yoshida, H., Hayashi, S., Kunisada, T., Ogawa, M., Nishikawa, S., Okamura, H., Sudo, T., and Shultz, L. D. (1990). The murine mutation osteopetrosis is in the coding region of the macrophage colony stimulating factor gene. Nature 345, 442–444. Takahashi, N., Udagawa, N., Akatsu, T., Tanaka, H., Shionome, M., and Suda, T. (1991). Role of colony-stimulating factors in osteoclast development. J. Bone Miner. Res. 6, 977–985. Hattersley, G., Dorey, E., Horton, M. A., and Chambers, T. J. (1988). Human macrophage colony-stimulating factor inhibits bone resorption by osteoclasts disaggregated from rat bone. J. Cell Physiol. 137, 199–203. Hofstetter, W., Wetterwald, A., Cecchini, M. C., Felix, R., Fleisch, H., and Mueller, C. (1992). Detection of transcripts for the receptor for macrophage colony-stimulating factor, c-fms, in murine osteoclasts. Proc. Natl Acad. Sci. USA 89, 9637–9641. Lorenzo, J. A., Sousa, S. L., Fonseca, J. M., Hock, J. M., and Medlock, E. S. (1987). Colony-stimulating factors regulate the development of multinucleated osteoclasts from recently replicated cells in vitro. J. Clin. Invest. 80, 160–164. Simonet, W. S., Lacey, D. L., Dunstan, C. R., Kelley, M., Chang, M. S., Luthy, R., Nguyen, H. Q., Wooden, S., Bennett, L., Boone, T., Shimamoto, G., DeRose, M., Elliott, R., Colombero, A., Tan, H. L.,

269

Chapter 15 Signaling in Bone

39.

40.

41.

42.

43.

44.

45.

46.

47.

48.

49.

50.

Trail, G., Sullivan, J., Davy, E., Bucay, N., Renshaw-Gegg, L., Hughes, T. M., Hill, D., Pattison, W., Campbell, P., Sander, S., Van, G., Tarpley, J., Derby, P., Lee, R., and Boyle, W. J. (1997). Osteoprotegerin: a novel secreted protein involved in the regulation of bone density. Cell 89, 309–319. Tsuda, E., Goto, M., Mochizuki, S., Yano, K., Kobayashi, F., Morinaga, T., and Higashio, K. (1997). Isolation of a novel cytokine from human fibroblasts that specifically inhibits osteoclastogenesis. Biochem. Biophys. Res. Commun. 234, 137–142. Lacey, D. L., Timms, E., Tan, H. L., Kelley, M. J., Dunstan, C. R., Burgess, T., Elliott, R., Colombero, A., Elliott, G., Scully, S., Hsu, H., Sullivan, J., Hawkins, N., Davy, E., Capparelli, C., Eli, A., Qian, Y. X., Kaufman, S., Sarosi, I., Shalhoub, V., Senaldi, G., Guo, J., Delaney, J., and Boyle, W. J. (1998). Osteoprotegerin ligand is a cytokine that regulates osteoclast differentiation and activation. Cell 93, 165–176. Yasuda, H., Shima, N., Nakagawa, N., Yamaguchi, K., Kinosaki, M., Mochizuki, S., Tomoyasu, A., Yano, K., Goto, M., Murakami, A., Tsuda, E., Morinaga, T., Higashio, K., Udagawa, N., Takahashi, N., and Suda, T. (1998). Osteoclast differentiation factor is a ligand for osteoprotegerin/osteoclastogenesis-inhibitory factor and is identical to TRANCE/RANKL. Proc. Natl Acad. Sci. USA 95, 3597–3602. Bucay, N., Sarosi, I., Dunstan, C. R., Morony, S., Tarpley, J., Capparelli, C., Scully, S., Tan, H. L., Xu, W., Lacey, D. L., Boyle, W. J., and Simonet, W. S. (1998). Osteoprotegerin-deficient mice develop early onset osteoporosis and arterial calcification. Genes Dev. 12, 1260–1268. Kong, Y. Y., Feige, U., Sarosi, I., Bolon, B., Tafuri, A., Morony, S., Capparelli, C., Li, J., Elliott, R., McCabe, S., Wong, T., Campagnuolo, G., Moran, E., Bogoch, E. R., Van, G., Nguyen, L. T., Ohashi, P. S., Lacey, D. L., Fish, E., Boyle, W. J., and Penninger, J. M. (1999). Activated T cells regulate bone loss and joint destruction in adjuvant arthritis through osteoprotegerin ligand. Nature 402, 304–309. Dougall, W. C., Glaccum, M., Charrier, K., Rohrbach, K., Brasel, K., De Smedt, T., Daro, E., Smith, J., Tometsko, M. E., Maliszewski, C. R., Armstrong, A., Shen, V., Bain, S., Cosman, D., Anderson, D., Morrissey, P. J., Peschon, J. J., and Schuh, J. (1999). RANK is essential for osteoclast and lymph node development. Genes Dev 13, 2412–2424. Udagawa, N., Takahashi, N., Yasuda, H., Mizuno, A., Itoh, K., Ueno, Y., Shinki, T., Gillespie, M. T., Martin, T. J., Higashio, K., and Suda, T. (2000). Osteoprotegerin produced by osteoblasts is an important regulator in osteoclast development and function. Endocrinology 141, 3478–3484. Wong, B. R., Rho, J., Arron, J., Robinson, E., Orlinick, J., Chao, M., Kalachikov, S., Cayani, E., Bartlett, F. S., 3rd, Frankel, W. N., Lee, S. Y., and Choi, Y. (1997). TRANCE is a novel ligand of the tumor necrosis factor receptor family that activates c-Jun N-terminal kinase in T cells. J. Biol. Chem. 272, 25190–25194. Anderson, D. M., Maraskovsky, E., Billingsley, W. L., Dougall, W. C., Tometsko, M. E., Roux, E. R., Teepe, M. C., DuBose, R. F., Cosman, D., and Galibert, L. (1997). A homologue of the TNF receptor and its ligand enhance T-cell growth and dendritic-cell function. Nature 390, 175–179. Wong, B. R., Josien, R., Lee, S. Y., Sauter, B., Li, H. L., Steinman, R. M., and Choi, Y. (1997). TRANCE (tumor necrosis factor [TNF]-related activation-induced cytokine), a new TNF family member predominantly expressed in T cells, is a dendritic cell-specific survival factor. J. Exp. Med. 186, 2075–2080. Horwood, N. J., Elliott, J., Martin, T. J., and Gillespie, M. T. (1998). Osteotropic agents regulate the expression of osteoclast differentiation factor and osteoprotegerin in osteoblastic stromal cells. Endocrinology 139, 4743–4746. Jimi, E., Akiyama, S., Tsurukai, T., Okahashi, N., Kobayashi, K., Udagawa, N., Nishihara, T., Takahashi, N., and Suda, T. (1999). Osteoclast differentiation factor acts as a multifunctional regulator in

51.

52.

53.

54.

55.

56.

57.

58.

59.

60.

61.

62.

63.

64.

murine osteoclast differentiation and function. J. Immunol. 163, 434–442. Jimi, E., Nakamura, I., Amano, H., Taguchi, Y., Tsurukai, T., Tamura, M., Takahashi, N., and Suda, T. (1996). Osteoclast function is activated by osteoblastic cells through a mechanism involving cell-to-cell contact. Endocrinology 137, 2187–2190. Quinn, J. M., Elliott, J., Gillespie, M. T., and Martin, T. J. (1998). A combination of osteoclast differentiation factor and macrophagecolony stimulating factor is sufficient for both human and mouse osteoclast formation in vitro. Endocrinology 139, 4424–4427. Matsuzaki, K., Udagawa, N., Takahashi, N., Yamaguchi, K., Yasuda, H., Shima, N., Morinaga, T., Toyama, Y., Yabe, Y., Higashio, K., and Suda, T. (1998). Osteoclast differentiation factor (ODF) induces osteoclast-like cell formation in human peripheral blood mononuclear cell cultures. Biochem. Biophys. Res. Commun. 246, 199–204. Thomas, R. J., Guise, T. A., Yin, J. J., Elliott, J., Horwood, N. J., Martin, T. J., and Gillespie, M. T. (1999). Breast cancer cells interact with osteoblasts to support osteoclast formation. Endocrinology 140, 4451–4458. Lai, F. P., Cole-Sinclair, M., Cheng, W. J., Quinn, J. M., Gillespie, M. T., Sentry, J. W., and Schneider, H. G. (2004). Myeloma cells can directly contribute to the pool of RANKL in bone bypassing the classic stromal and osteoblast pathway of osteoclast stimulation. Br. J. Haematol. 126, 192–201. Eghbali-Fatourechi, G., Khosla, S., Sanyal, A., Boyle, W. J., Lacey, D. L., and Riggs, B. L. (2003). Role of RANK ligand in mediating increased bone resorption in early postmenopausal women. J. Clin. Invest. 111, 1221–1230. Darnay, B. G., Ni, J., Moore, P. A., and Aggarwal, B. B. (1999). Activation of NF-kappaB by RANK requires tumor necrosis factor receptor-associated factor (TRAF) 6 and NF-kappaB-inducing kinase. Identification of a novel TRAF6 interaction motif. J. Biol. Chem. 274, 7724–7731. Galibert, L., Tometsko, M. E., Anderson, D. M., Cosman, D., and Dougall, W. C. (1998). The involvement of multiple tumor necrosis factor receptor (TNFR)-associated factors in the signaling mechanisms of receptor activator of NF-kappaB, a member of the TNFR superfamily. J. Biol. Chem. 273, 34120–34127. Franzoso, G., Carlson, L., Xing, L., Poljak, L., Shores, E. W., Brown, K. D., Leonardi, A., Tran, T., Boyce, B. F., and Siebenlist, U. (1997). Requirement for NF-kappaB in osteoclast and B-cell development. Genes Dev. 11, 3482–3496. Naito, A., Azuma, S., Tanaka, S., Miyazaki, T., Takaki, S., Takatsu, K., Nakao, K., Nakamura, K., Katsuki, M., Yamamoto, T., and Inoue, J. (1999). Severe osteopetrosis, defective interleukin-1 signalling and lymph node organogenesis in TRAF6-deficient mice. Genes Cells 4, 353–362. Lomaga, M. A., Yeh, W. C., Sarosi, I., Duncan, G. S., Furlonger, C., Ho, A., Morony, S., Capparelli, C., Van, G., Kaufman, S., van der Heiden, A., Itie, A., Wakeham, A., Khoo, W., Sasaki, T., Cao, Z., Penninger, J. M., Paige, C. J., Lacey, D. L., Dunstan, C. R., Boyle, W. J., Goeddel, D. V., and Mak, T. W. (1999). TRAF6 deficiency results in osteopetrosis and defective interleukin-1, CD40, and LPS signaling. Genes Dev 13, 1015–1024. Takayanagi, H., Kim, S., Matsuo, K., Suzuki, H., Suzuki, T., Sato, K., Yokochi, T., Oda, H., Nakamura, K., Ida, N., Wagner, E. F., and Taniguchi, T. (2002). RANKL maintains bone homeostasis through c-Fos-dependent induction of interferon-beta. Nature 416, 744–749. Grigoriadis, A. E., Wang, Z. Q., Cecchini, M. G., Hofstetter, W., Felix, R., Fleisch, H. A., and Wagner, E. F. (1994). c-Fos: a key regulator of osteoclast-macrophage lineage determination and bone remodeling. Science 266, 443–448. Takayanagi, H., Kim, S., Koga, T., Nishina, H., Isshiki, M., Yoshida, H., Saiura, A., Isobe, M., Yokochi, T., Inoue, J., Wagner, E. F., Mak, T. W., Kodama, T., and Taniguchi, T. (2002). Induction and activation of the

270

65.

66.

67.

68.

69.

70.

71.

72.

73.

74.

75.

76.

77.

78.

79.

80.

81.

transcription factor NFATc1 (NFAT2) integrate RANKL signaling in terminal differentiation of osteoclasts. Dev. Cell 3, 889–901. Erlebacher, A., Filvaroff, E. H., Ye, J. Q., and Derynck, R. (1998). Osteoblastic responses to TGF-beta during bone remodeling. Mol. Biol. Cell 9, 1903–1918. Rodan, G. A. (1991). Mechanical loading, estrogen deficiency, and the coupling of bone formation to bone resorption. J. Bone Miner. Res. 6, 527–530. Chambers, T. J., and Fuller, K. (1985). Bone cells predispose bone surfaces to resorption by exposure of mineral to osteoclastic contact. J. Cell Sci. 76, 155–165. Partridge, N. C., Jeffrey, J. J., Ehlich, L. S., Teitelbaum, S. L., Fliszar, C., Welgus, H. G., and Kahn, A. J. (1987). Hormonal regulation of the production of collagenase and a collagenase inhibitor activity by rat osteogenic sarcoma cells. Endocrinology 120, 1956–1962. Chiusaroli, R., Maier, A., Knight, M. C., Byrne, M., Calvi, L. M., Baron, R., Krane, S. M., and Schipani, E. (2003). Collagenase cleavage of type I collagen is essential for both basal and parathyroid hormone (PTH)/PTH-related peptide receptor-induced osteoclast activation and has differential effects on discrete bone compartments. Endocrinology 144, 4106–4116. Everts, V., Delaisse, J. M., Korper, W., Jansen, D. C., TigchelaarGutter, W., Saftig, P., and Beertsen, W. (2002). The bone lining cell: its role in cleaning Howship’s lacunae and initiating bone formation. J. Bone Miner. Res. 17, 77–90. Hauschka, P. V., Mavrakos, A. E., Iafrati, M. D., Doleman, S. E., and Klagsbrun, M. (1986). Growth factors in bone matrix. Isolation of multiple types by affinity chromatography on heparin-Sepharose. J. Biol. Chem. 261, 12665–12674. Schmid, C., and Ernst, M. (1992). Insulin-like growth factors. In “Cytokines and Bone Metabolism”, pp. 229–265. CRC Press, Boca Raton, FL. Rodan, S. B., and Rodan, G. A. (1992). Fibroblast growth factor and platelet-derived growth factor. In “Cytokines and Bone Metabolism”, pp. 115–145. CRC Press, Boca Raton, FL. Centrella, M., McCarthy, T. L., and Canalis, E. (1991). Transforming growth factor-beta and remodeling of bone. J. Bone Joint Surg. Am. 73, 1418–1428. Ogawa, Y., Schmidt, D. K., Dasch, J. R., Chang, R. J., and Glaser, C. B. (1992). Purification and characterization of transforming growth factor-beta 2.3 and -beta 1.2 heterodimers from bovine bone. J. Biol. Chem. 267, 2325–2328. Harris, S. E., Sabatini, M., Harris, M. A., Feng, J. Q., Wozney, J., and Mundy, G. R. (1994). Expression of bone morphogenetic protein messenger RNA in prolonged cultures of fetal rat calvarial cells. J. Bone Miner. Res. 9, 389–394. Vukicevic, S., Latin, V., Chen, P., Batorsky, R., Reddi, A. H., and Sampath, T. K. (1994). Localization of osteogenic protein-1 (bone morphogenetic protein-7) during human embryonic development: high affinity binding to basement membranes. Biochem. Biophys. Res. Commun. 198, 693–700. Vukicevic, S., Paralkar, V. M., Cunningham, N. S., Gutkind, J. S., and Reddi, A. H. (1990). Autoradiographic localization of osteogenin binding sites in cartilage and bone during rat embryonic development. Dev. Biol. 140, 209–214. Rydziel, S., Ladd, C., McCarthy, T. L., Centrella, M., and Canalis, E. (1992). Determination and expression of platelet-derived growth factor-AA in bone cell cultures. Endocrinology 130, 1916–1922. McCarthy, T. L., Centrella, M., Raisz, L. G., and Canalis, E. (1991). Prostaglandin E2 stimulates insulin-like growth factor I synthesis in osteoblast-enriched cultures from fetal rat bone. Endocrinology 128, 2895–2900. McCarthy, T. L., Centrella, M., and Canalis, E. (1989). Parathyroid hormone enhances the transcript and polypeptide levels of insulin-like

T. JOHN MARTIN AND NATALIE A. SIMS

82.

83.

84.

85.

86.

87.

88.

89.

90.

91.

92.

93.

94.

95.

96.

97.

98.

99.

growth factor I in osteoblast-enriched cultures from fetal rat bone. Endocrinology 124, 1247–1253. Turner, R. T., Backup, P., Sherman, P. J., Hill, E., Evans, G. L., and Spelsberg, T. C. (1992). Mechanism of action of estrogen on intramembranous bone formation: regulation of osteoblast differentiation and activity. Endocrinology 131, 883–889. Shinar, D. M., Endo, N., Halperin, D., Rodan, G. A., and Weinreb, M. (1993). Differential expression of insulin-like growth factor-I (IGF-I) and IGF-II messenger ribonucleic acid in growing rat bone. Endocrinology 132, 1158–1167. Suva, L. J., Seedor, J. G., Endo, N., Quartuccio, H. A., Thompson, D. D., Bab, I., and Rodan, G. A. (1993). Pattern of gene expression following rat tibial marrow ablation. J. Bone Miner. Res. 8, 379–388. Centrella, M., Horowitz, M. C., Wozney, J. M., and McCarthy, T. L. (1994). Transforming growth factor-beta gene family members and bone. Endocr. Rev. 15, 27–39. Nakashima, M., Nagasawa, H., Yamada, Y., and Reddi, A. H. (1994). Regulatory role of transforming growth factor-beta, bone morphogenetic protein-2, and protein-4 on gene expression of extracellular matrix proteins and differentiation of dental pulp cells. Dev. Biol. 162, 18–28. Heikinheimo, K. (1994). Stage-specific expression of decapentaplegicVg-related genes 2, 4, and 6 (bone morphogenetic proteins 2, 4, and 6) during human tooth morphogenesis. J. Dent. Res. 73, 590–597. Globus, R. K., Plouet, J., and Gospodarowicz, D. (1989). Cultured bovine bone cells synthesize basic fibroblast growth factor and store it in their extracellular matrix. Endocrinology 124, 1539–1547. Somjen, D., Binderman, I., Berger, E., and Harell, A. (1980). Bone remodeling induced by physical stress is prostaglandin E2 mediated. Biochim. Biophys. Acta. 627, 91–100. Raisz, L. G., and Martin, T. J. (1984). Prostaglandins in bone and mineral metabolism. In “Bone and Mineral Research” (W. A. Pecked, ed.), pp. 286–310. Elsevier, Amsterdam. Spencer, E. M., Liu, C. C., Si, E. C., and Howard, G. A. (1991). In vivo actions of insulin-like growth factor-I (IGF-I) on bone formation and resorption in rats. Bone 12, 21–26. Tobias, J. H., Chow, J. W., and Chambers, T. J. (1992). Opposite effects of insulin-like growth factor-I on the formation of trabecular and cortical bone in adult female rats. Endocrinology 131, 2387–2392. Bagi, C. M., Brommage, R., Deleon, L., Adams, S., Rosen, D., and Sommer, A. (1994). Benefit of systemically administered rhIGF-I and rhIGF-I/IGFBP-3 on cancellous bone in ovariectomized rats. J. Bone Miner. Res. 9, 1301–1312. Critchlow, M. A., Bland, Y. S., and Ashhurst, D. E. (1994). The effects of age on the response of rabbit periosteal osteoprogenitor cells to exogenous transforming growth factor-beta 2. J. Cell Sci. 107 (Pt 2), 499–516. Joyce, M. E., Roberts, A. B., Sporn, M. B., and Bolander, M. E. (1990). Transforming growth factor-beta and the initiation of chondrogenesis and osteogenesis in the rat femur. J. Cell Biol. 110, 2195–2207. Kawaguchi, H., Kurokawa, T., Hanada, K., Hiyama, Y., Tamura, M., Ogata, E., and Matsumoto, T. (1994). Stimulation of fracture repair by recombinant human basic fibroblast growth factor in normal and streptozotocin-diabetic rats. Endocrinology 135, 774–781. Marks, S. C., Jr., and Miller, S. (1988). Local infusion of prostaglandin E1 stimulates mandibular bone formation in vivo. J. Oral Pathol. 17, 500–505. Lund, J. E. (1982). The toxicology of PGE1 and PGE2. In “Prostaglandins in Clinical Medicine”, pp. 93–109. Yearbook Medical Publishers, New York. Bonewald, L. F., Wakefield, L., Oreffo, R. O., Escobedo, A., Twardzik, D. R., and Mundy, G. R. (1991). Latent forms of transforming

271

Chapter 15 Signaling in Bone

100.

101.

102.

103.

104.

105.

106.

107.

108.

109.

110. 111.

growth factor-beta (TGF beta) derived from bone cultures: identification of a naturally occurring 100-kDa complex with similarity to recombinant latent TGF beta. Mol. Endocrinol. 5, 741–751. Oreffo, R. O., Mundy, G. R., Seyedin, S. M., and Bonewald, L. F. (1989). Activation of the bone-derived latent TGF beta complex by isolated osteoclasts. Biochem. Biophys. Res. Commun. 158, 817–823. Underwood, L. E., Thissen, J. P., Lemozy, S., Ketelslegers, J. M., and Clemmons, D. R. (1994). Hormonal and nutritional regulation of IGF-I and its binding proteins. Horm. Res. 42, 145–151. Martin, T. J., Allan, E. H., and Fukumoto, S. (1993). The plasminogen activator and inhibitor system in bone remodeling. Growth Regul. 3, 209–214. Campbell, P. G., Novak, J. F., Yanosick, T. B., and McMaster, J. H. (1992). Involvement of the plasmin system in dissociation of the insulin-like growth factor-binding protein complex. Endocrinology 130, 1401–1412. Gray, C., Boyde, A., and Jones, S. J. (1996). Topographically induced bone formation in vitro: implications for bone implants and bone grafts. Bone 18, 115–123. Nakamura, M., Udagawa, N., Matsuura, S., Mogi, M., Nakamura, H., Horiuchi, H., Saito, N., Hiraoka, B. Y., Kobayashi, Y., Takaoka, K., Ozawa, H., Miyazawa, H., and Takahashi, N. (2003). Osteoprotegerin regulates bone formation through a coupling mechanism with bone resorption. Endocrinology 144, 5441–5449. Cleiren, E., Benichou, O., Van Hul, E., Gram, J., Bollerslev, J., Singer, F. R., Beaverson, K., Aledo, A., Whyte, M. P., Yoneyama, T., deVernejoul, M. C., and Van Hul, W. (2001). Albers-Schonberg disease (autosomal dominant osteopetrosis, type II) results from mutations in the ClCN7 chloride channel gene. Hum. Mol. Genet. 10, 2861–2867. Marzia, M., Sims, N. A., Voit, S., Migliaccio, S., Taranta, A., Bernardini, S., Faraggiana, T., Yoneda, T., Mundy, G. R., Boyce, B. F., Baron, R., and Teti, A. (2000). Decreased c-Src expression enhances osteoblast differentiation and bone formation. J. Cell Biol. 151, 311–320. Karsdal, M. A., Henriksen, K., Sorensen, M. G., Gram, J., Schaller, S., Dziegiel, M. H., Heegard, A. M., Christophersen, P., Martin, T. J., Christiansen, C., and Bollerslev, J. (2005). Acidification of the osteoclast resorption compartment provides insight into the coupling of bone formation to bone resorption. Am. J. Pathol. 166, 467–476. Chiusaroli, R., Knobler, H., Luxenburg, C., Sanjay, A., Granot-Attas, S., Tiran, Z., Miyazaki, T., Harmelin, A., Baron, R., and Elson, A. (2004). Tyrosine phosphatase epsilon is a positive regulator of osteoclast function in vitro and in vitro. Mol. Biol. Cell 15, 234–244. Heymann, D., and Rousselle, A. V. (2000). gp130 Cytokine family and bone cells. Cytokine 12, 1455–1468. Sims, N. A., Jenkins, B. J., Quinn, J. M., Nakamura, A., Glatt, M., Gillespie, M. T., Ernst, M., and Martin, T. J. (2004). Glycoprotein 130

112.

113.

114. 115.

116.

117.

118. 119.

120.

121.

122.

123.

regulates bone turnover and bone size by distinct downstream signaling pathways. J. Clin. Invest. 113, 379–389. Hoff, A. O., Catala-Lehnen, P., Thomas, P. M., Priemel, M., Rueger, J. M., Nasonkin, I., Bradley, A., Hughes, M. R., Ordonez, N., Cote, G. J., Amling, M., and Gagel, R. F. (2002). Increased bone mass is an unexpected phenotype associated with deletion of the calcitonin gene. J. Clin. Invest. 110, 1849–1857. Dacquin, R., Davey, R. A., Laplace, C., Levasseur, R., Morris, H. A., Goldring, S. R., Gebre-Medhin, S., Galson, D. L., Zajac, J. D., and Karsenty, G. (2004). Amylin inhibits bone resorption while the calcitonin receptor controls bone formation in vitro. J. Cell Biol. 164, 509–514. Sexton, P. M., Findlay, D. M., and Martin, T. J. (1999). Calcitonin. Curr. Med. Chem. 6, 1067–1093. Demiralp, B., Chen, H. L., Koh, A. J., Keller, E. T., and McCauley, L. K. (2002). Anabolic actions of parathyroid hormone during bone growth are dependent on c-fos. Endocrinology 143, 4038–4047. Black, D. M., Greenspan, S. L., Ensrud, K. E., Palermo, L., McGowan, J. A., Lang, T. F., Garnero, P., Bouxsein, M. L., Bilezikian, J. P., and Rosen, C. J. (2003). The effects of parathyroid hormone and alendronate alone or in combination in postmenopausal osteoporosis. N. Engl. J. Med. 349, 1207–1215. Finkelstein, J. S., Hayes, A., Hunzelman, J. L., Wyland, J. J., Lee, H., and Neer, R. M. (2003). The effects of parathyroid hormone, alendronate, or both in men with osteoporosis. N. Engl. J. Med. 349, 1216–1226. Khosla, S. (2003). Parathyroid hormone plus alendronate—a combination that does not add up. N. Engl. J. Med. 349, 1277–1279. Martin, T. J. (2004). Does bone resorption inhibition affect the anabolic response to parathyroid hormone? Trends Endocrinol. Metab. 15, 49–50. Onyia, J. E., Bidwell, J., Herring, J., Hulman, J., and Hock, J. M. (1995). In vitro, human parathyroid hormone fragment (hPTH 1–34) transiently stimulates immediate early response gene expression, but not proliferation, in trabecular bone cells of young rats. Bone 17, 479–484. Ma, Y. L., Cain, R. L., Halladay, D. L., Yang, X., Zeng, Q., Miles, R. R., Chandrasekhar, S., Martin, T. J., and Onyia, J. E. (2001). Catabolic effects of continuous human PTH (1–38) in vitro is associated with sustained stimulation of RANKL and inhibition of osteoprotegerin and gene-associated bone formation. Endocrinology 142, 4047– 4054. Holtrop, M. E., King, G. J., Cox, K. A., and Reit, B. (1979). Time-related changes in the ultrastructure of osteoclasts after injection of parathyroid hormone in young rats. Calcif. Tissue Int. 27, 129–135. Martin, T. J., and Sims, N. A. (2005). On the coupling of bone formation to resorption. Trends Molecular Med. 11, 76–81.

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Chapter 16

Parathyroid Hormone: Structure, Function and Dynamic Actions Lorraine A. Fitzpatrick John P. Bilezikian

I. II. III. IV. V. VI. VII.

Global Development, Amgen, Thousand Oaks, CA Departments of Medicine and Pharmacology, College of Physicians and Surgeons, Columbia University, New York, NY

VIII. Structure of the PTH/PTHrP (PTH1R) Receptor IX. Activation of the Cyclic Adenosine Monophosphate Second-Messenger System by Parathyroid Hormone X. Identification of a Second PTH Receptor XI. Physiological Actions of PTH XII. Cell-to-Cell Communication: Osteoblasts and Osteoclasts XIII. Preferential Actions of PTH at Selected Skeletal Sites References

Introduction Structure of the PTH Gene Chromosome Location Control of Gene Expression Biosynthesis of Parathyroid Hormone Metabolism of Parathyroid Hormone Receptor Interactions of Parathyroid Hormone and Parathyroid Hormone-Related Protein

I. INTRODUCTION

1,25-dihydroxy vitamin D and enhances the intestinal absorption of calcium. These actions of PTH are mediated through a G protein-coupled receptor system in the cells of target tissues. A rise in extracellular calcium inhibits further secretion of parathyroid hormone.

Parathyroid hormone (PTH) is an 84 amino acid hormone that regulates calcium homeostasis via its actions on target tissues. PTH maintains serum calcium concentrations within a narrow physiological range by direct actions on bone and kidney tissue and an indirect action, via 1,25-dihydroxy vitamin D, on the intestinal tract. PTH release and gene expression, in turn, are regulated by serum calcium concentrations. Hypocalcemia stimulates the release of PTH from the parathyroid gland. By reabsorption of calcium in the distal convoluted tubule of the kidney or by osteoclast-mediated bone resorption, serum calcium concentrations increase. PTH also stimulates renal 1-α-hydroxylase activity in the kidney which increases Dynamics of Bone and Cartilage Metabolism

II. STRUCTURE OF THE PTH GENE The gene that encodes PTH is representative of typical eukaryotic genes with consensus sequences for initiation of RNA synthesis, RNA splicing and polyadenylation. Restriction enzyme analysis of the human PTH gene has indicated polymorphism in the cleavage products in different individuals that are useful for genetic analysis [1]. 273

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Complementary DNAs encoding for human, rat, chicken, bovine and dog PTH have been cloned. These genes have several structural features in common: all contain two introns or intervening sequences and three exons (Fig. 1) [2]. The introns vary in size among species. The first intron is large; the second intron is smaller (about 100 base pairs) in human, bovine, and rat genes. The first intron encodes the 5′ noncoding sequences; the second exon encodes most of the “prepro” sequence of prepro parathyroid hormone. The mature PTH sequence and the 3′ noncoding region are contained within the third exon. RNA is transcribed from the sequences in both the introns and exons. Subsequently, RNA sequences derived from the introns are spliced. The mature PTH mRNA is then translated into prepro PTH. Considerable homology among the mammalian PTH genes is seen between human and bovine proteins (85%) and between human and rat proteins (75%). Identity is less in the 3′ noncoding region. In contrast, human and bovine PTH genes contain two functional TATA transcription

Figure 1

start sites where the rat and chicken genes have only one. Both initiation sites of transcription are utilized in the bovine and human genes. Tissue specificity of PTH gene expression is determined by coding regions upstream of the structural gene. Analysis of these regulatory sequences has been difficult due to the lack of a well-defined cell line for gene expression. A cAMP-response element has been identified and other sequences that bind 1,25(OH)2D3 receptors have been recognized [3]. PTH gene expression is regulated transcriptionally by the same mechanism, in part, by calcium and phosphate [4, 5].

III. CHROMOSOME LOCATION The human PTH gene has been localized to the short arm of chromosome 11 and is close to the calcitonin gene [6, 7]. Additional localization studies have indicated that the human PTH gene is located at band 11p15 [8].

Primary structure of human preparathyroid hormone. The arrows indicate sites of specific cleavages which occur in the sequence of biosynthesis and peripheral metabolism. The biologically active sequence is noted. Reprinted from Choren and Rosenblatt [2a], with permission.

Chapter 16 Parathyroid Hormone: Structure, Function and Dynamic Actions

Restriction fragment length polymorphisms indicate that the human PTH gene is linked to genes encoding calcitonin, catalase, insulin, H-ras, and β-globin [1]. The development of mice homozygous for a null PTH allele assisted in assessing the role of PTH in the modulation of skeletal development in the fetus. Mice carrying the disrupted PTH gene (PTH−/− mice) were derived by homologous recombination in embryonic stem cells [9]. Parathyroid glands were greatly enlarged and PTH expression was absent in these mice, compared to wild-type littermates. In the PTH−/− mice, the skull was abnormal and mineralization of skull bones was enhanced. The vertebral column had smaller vertebral bodies and mineralized metacarpal and metatarsal bones were shorter. Lengths of the tibae were normal and the hypertrophic cartilage zone was enlarged with a higher ratio of the hypertrophic zone to proliferating zone. However, proliferation of chondrocytes was not altered, and there was enhancement of the deposition of type X collagen in the hypertrophic zone. Cortical thickness of long bones was increased, but the trabecular components were diminished. Reduced osteoblast numbers were present in the primary spongiosa, with increased apoptosis. PTH appeared to be essential for normal cartilage remodeling, and the reduced osteoblasts resulted in reduced trabecular bone volume. When PTH−/− mice were placed on a low-calcium diet, renal 1-α-hydyroxylase expression increased in spite of the lack of PTH. There was a rise in 1,25(OH)2D3 levels, marked osteoclastogenesis, and profound bone resorption. These latter studies indicate that in the complete absence of PTH, calcium stores can be mobilized through the actions of 1,25(OH)2D3 to guard against extreme hypocalcemia [10]. Clinical studies have attempted to link gene polymorphisms in the PTH gene to specific human disease states. No linkages were found in bone phenotypes in Chinese subjects [11], or in sporadic idiopathic hypoparathyroidism [12]. PTH gene polymorphism accounted for about 7–9% of the total variances of bone dimensional variables in a group of 91 healthy Caucasian women. Bone diameter, cortical thickness, and cross-sectional area at standard sites of the metacarpals, radius, and femur were measured with radiogrammetry. Higher metacarpal diameter and cross-sectional cortical area and a slower decrease in radial cortical area with age were associated with the absence of the BstBI restriction site of the PTH gene [13]. In secondary hyperparathyroidism associated with renal hemodialysis, PTH genotypes were determined by PCR and by restriction fragment length polymorphisms (RFLPs) of BstBI and DraII. There were no differences in these genotypes between hemodialysis patients and healthy controls. There was a significant difference in the serum intact PTH levels between Bb/bb and BB genotypes

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(P < 0.02) [14]. In another study, PTH genotypes were determined by PCR and BstBI or DRAII RFLPs were determined in patients with primary hyperparathyroidism. The BstBI polymorphism of the PTH gene was closely linked to bone mineral density, but not the development of primary hyperparathyroidism. However, serum calcium was higher in those with the BB genotype, compared to the BB/bb genotypes, suggesting that the severity of primary hyperparathyroidism may be linked to this genotype [15]. A novel mutation in the signal peptide of the prepro PTH gene was associated with autosomal-recessive familialisolated hypoparathyroidism in a child of consanguineous parents who presented with neonatal hypocalcemia seizures. Replacement of thymine with a cytosine was found in the first nucleotide of position 23 in the 25-amino acid signal peptide, resulting in the replacement of the normal Ser (TCG) with a Pro (CCG). The parents were heterozygous for the allele [16].

IV. CONTROL OF GENE EXPRESSION A. The Role of Extracellular Calcium Minute-to-minute regulation of serum calcium levels is dependent on the regulation of secretion of PTH by calcium. Long-term homeostasis is dependent on several additional regulatory mechanisms. Expression of the PTH gene occurs almost exclusively in the parathyroid chief cells. Early studies indicated that increased levels of extracellular calcium resulted in a reversible fall in PTH mRNA in cultured parathyroid cells [17]. Suppression was most pronounced after 72 hours of incubation. The rate of PTH gene transcription was also reduced in the presence of high concentrations of calcium. In vivo studies in intact rats revealed rapid changes in PTH mRNA in response to changes in serum calcium concentrations [18]. The largest effects were noted in response to lowered levels of calcium (2.6–2.1 mmol/liter). Increased levels of serum calcium, whether induced by infusion of calcium or by transplantation of Walker carcinosarcoma 256 cells, had little effect on PTH mRNA levels. One major difference in the in vivo studies in comparison to the in vitro work is in the time course of response to calcium. In vivo, high calcium levels lead to a rapid fall in PTH mRNA within a few hours; in vitro, a longer time frame is required to detect changes in PTH mRNA in the models tested. Another major difference is that in vivo, hypercalcemia does not alter PTH mRNA levels, suggesting that at the molecular level the parathyroid gland is responsive to the signal of hypocalcemia.

276 The parathyroid cell senses changes in extracellular calcium through a calcium-sensing receptor (CaR). This receptor is a membrane-bound, seven transmembrane protein that has three major domains: an extracellular 612-amino acid ligand-binding segment, a hydrophobic 250-amino acid membrane-spanning section, and a cytosolic COOH-terminal tail of ~250 amino acids [19]. Activation of the CaR by extracellular calcium ions engages the mitogen-activating protein kinase C cascade through both G protein-linked phospholipase C and a member of the Src tyrosine kinase family [20, 21]. The pathway eventually leads to activation of phospholipase A2 and the generation of arachidonic acid [20]. Thus, PTH secretion is inhibited via the CaR when it is activated by elevated concentrations of ionized calcium. Prolonged hypercalcemia eventually leads to reduced parathyroid cell proliferation, an effect that is also likely to be mediated by the CaR [22]. Conversely, low serum levels of ionized calcium reduce CaR activity and PTH secretion increases. Many studies to evaluate the genetic mechanism of the action of calcium and the search for DNA sequences responsible for the transcriptional effects of calcium, have been hampered by the lack of a well-differentiated parathyroid cell line that produces PTH in response to changes in extracellular calcium. Several short sequences located several thousand base pairs upstream from the start site of PTH gene transcription may be involved in the regulation of PTH by calcium. A negative calcium regulatory element (nCaRE) has been identified in the PTH gene. It is also present in the atrial natriuretic peptide gene [23]. Further studies by the same investigators revealed the identity of a redox factor protein [1] that activates several transcription factors and binds nCaRE. The level of ref1 mRNA and protein were elevated by an increase in extracellular calcium concentration. Unfortunately, these studies had to be performed in a nonparathyroid cell line so the role of the protein in the regulation of PTH expression remains to be determined. The in vivo effects of calcium also may be due to posttranscriptional mechanisms. Preliminary evidence indicates that regions in the 3′ UTR of PTH mRNA binds specifically to cytosolic proteins and may be involved in the post-transcriptional increase in PTH gene expression induced by decreased calcium concentration [24]. Bovine parathyroid cells were incubated in the presence of low extracellular calcium (0.4 mM) and resulted in a post-transcriptional increase in the membrane-bound polysomal content of PTH mRNA [25]. Using a cell-free assay system, Vadher et al. [26] presented preliminary evidence that the 3′-untranslated region is important in the translational regulation of PTH synthesis by cytosolic

Lorraine A. Fitzpatrick and John P. Bilezikian

regulatory proteins. These in vitro studies add support for a post-translational role of calcium on PTH gene expression. Taken together, these studies suggest that posttranslational regulation of PTH mRNA may involve calcium-sensitive proteins binding to the 3′ UTR region. Additional work indicates that 26 nucleotide is the minimal sequence for protein binding in the PTH mRNA 3′ UTR. By using chimeric growth hormone PTH 63 nucleotide transcripts, the investigators were able to determine that the protein- binding region of the PTH mRNA 3′ UTR is necessary and sufficient to confer responsiveness to calcium and phosphate. This element determined PTH mRNA stability and levels [27].

B. The role of 1,25 dihydroxyvitamin D In vivo and in vitro studies support the significant role of 1,25 dihydroxyvitamin D in the regulation of PTH gene expression via a well-defined feedback loop between PTH and 1,25(OH)2D3. The renal synthesis of 1-α hydroxylase is increased by PTH; in turn, 1,25(OH)2D3 increases serum calcium concentrations by enhancing intestinal absorption of calcium. 1,25(OH)2D3 reduces transcription of the PTH gene. In bovine parathyroid cells in primary culture, 1,25(OH)2D3 exposure results in reduced levels of PTH mRNA and PTH gene transcription rates [28, 29]. In vitro studies in rats confirm the regulatory role of 1,25(OH)2D3 in PTH gene expression. Rats that are injected with 1,25(OH)2D3 at levels that do not alter serum calcium levels show decreased transcription of the PTH gene and reduced levels of PTH mRNA [30]. Transfection assays and DNA-binding studies have attempted to identify specifically the sequences responsible for modulation of the PTH gene transcription by 1,25(OH)2D3. Utilizing a rat pituitary cell line (GH4), Okazaki et al. linked 684 base pairs of the 5′-flanking region of the human PTH gene to a reporter gene. Transfection of the rat pituitary cell line resulted in responsiveness of gene expression to 1,25(OH)2D3 [31]. DNA binding studies identified DNA sequences upstream of the PTH gene that bind to 1,25(OH)2D3 receptors in vitro. A specific 26 base pair sequence located 125 base pairs upstream from the transcription start site on the PTH gene binds to 1,25(OH)2D3 receptors. Confirmation of this activity was performed by transfection of a pituitary cell line with the sequence linked to a reporter gene, with the result that 1,25(OH)2D3 suppressed the reporter gene [3]. In contrast, the human PTH DNA sequence does not mediate repression of transcription in the osteoblast cell line, ROS 17/2.8, in spite of the fact that vitamin D response elements (VDRE) are active in these cells. One difference is that

Chapter 16 Parathyroid Hormone: Structure, Function and Dynamic Actions

human PTH DNA sequences contain a single copy of a hexameric motif homologous to those repeated in the up-regulatory vitamin D response elements. When nuclear extracts from bovine parathyroid cells are incubated with the vitamin D receptor, the receptor binds to the downregulatory human PTH DNA sequence independent of the presence of the retinoid X receptor (RXR). In nuclear extracts from GH4CI pituitary cells, two vitamin D-containing complexes are present, one of which contains RXR. In nuclear extracts derived from the ROS 17/2.8 osteoblasts, a single VDR-dependent complex containing RXR is present. The negative 1,25(OH)2D3 response element contains one copy of a motif that is duplicated in the mouse osteopontin gene, a gene that is up-regulated by 1,25(OH)2D3. When the osteocalcin vitamin D response element is utilized as a probe, only VDR–RXR-containing complexes are generated from the nuclear extracts of all three cell types. These experiments indicate that transcriptional repression in response to 1,25(OH)2D3 differs from the up-regulatory response elements in sequence composition and in the ability to bind VDR independent of RXR [32]. Calreticulin is a calcium-binding protein present in the lumen of the endoplasmic reticulum of the cell. Using a model of chronic hypocalcemia, with increased PTH gene expression, calreticulin protein was increased in the nuclear fraction of parathyroids. In vitro, calreticulin blocked DNA binding of the VDR-RXRβ heterodimers to the cPTH VDRE. The studies suggest that calreticulin may prevent the transcriptional effect of 1,25 (OH)2D3 on the PTH promoter [33].

C. Phosphate as a Modulator of PTH Gene Expression PTH has a potent effect on the tubular reabsorption of phosphate by the kidney. This observation has been utilized as a clinical test to evaluate target organ responsiveness to parathyroid hormone. In severe renal failure, hyperphosphatemia results in secondary hyperparathyroidism. Clinically, the effects of the associated hypocalcemia and decrease in serum levels of 1,25(OH)2D3 have complicated our understanding of the role of phosphate on PTH secretion. Slatopolsky and Bricker [34] showed that in experimental renal failure, restriction of dietary phosphate prevented secondary hyperparathyroidism. Clinical studies confirmed the utility of restricting oral phosphate intake. These initial in vitro and in vivo studies suggested that phosphate directly alters that production of 1,25(OH)2D3. Recently, a series of elegant, carefully designed studies have shown that the effect of phosphate on PTH is independent

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of serum calcium and 1,25(OH)2D3 levels. Uremic dogs placed on diets deficient in both calcium and phosphate have lower levels of serum phosphate and calcium but no change in the concentration of 1,25(OH)2D3 [35]. In hypophosphatemic rats, phosphate regulates the PTH gene independent of the effects of phosphate on calcium and 1,25(OH)2D3 [36]. These studies indicate that phosphate regulates PTH in a hypophosphatemic, normocalcemic model in the presence of normal levels of 1,25(OH)2D3. Nuclear transcript run-on assays show that the decrease in PTH mRNA due to low phosphate was post-transcriptional in contrast to the transcriptional effects of 1,25(OH)2D3 on the PTH gene. Recently, protein–mRNA binding studies in hypophosphatemic rats reveal decreased binding of PTH cytosolic proteins. Confirmation of the functional role was provided with an in vitro degradation assay. The functional life of intact transcripts was reduced from 40 minutes in parathyroid proteins obtained from control rats to 5 minutes in the proteins from hypophosphatemic animals. In contrast, parathyroid proteins from hypocalcemic rats had increased binding and intact transcripts were present for a longer period of time (180 minutes). The study also indicates that degradation of PTH mRNA is dependent on sequences in the 3′ UTR region [37]. In human parathyroid tissue, it appears that phosphate directly regulates PTH secretion and PTH mRNA levels [38]. An additional mechanism by which 1,25(OH)2D3 may regulate PTH gene expression is through alterations in the 1,25(OH)2D3 receptor. Modulation of the concentration of the 1,25(OH)2D3 receptor may alter the effect of 1,25(OH)2D3 at its target sites. The administration of 1,25(OH)2D3 to rats results in an increase in 1,25(OH)2D3 receptors in parathyroid tissue with a concomitant decrease in PTH mRNA [39]. In another study, weanling rats fed a calcium-deficient diet became markedly hypocalcemic with high serum levels of 1,25(OH)2D3. There was no change in 1,25(OH)2D3 receptor mRNA levels in spite of the high levels of 1,25(OH)2D3; furthermore, PTH mRNA levels were increased. The low serum calcium levels may prevent the increase in parathyroid 1,25(OH)2D3 receptor levels and suppression of PTH mRNA levels. These findings have been confirmed in an avian model [40] and in vitamin D-deficient rats [41]. The use of calcitriol analogs that are biologically active but result in less hypercalcemia also have helped to separate the effects of calcium from the actions of vitamin D on PTH gene expression. Detailed dose–response curves suggest that 1,25(OH)2D3 is the most effective of the vitamin D forms to inhibit PTH levels in vivo. The ability of 1,25(OH)2D3 to decrease PTH gene expression is a significant therapeutic management tool for patients at

278 risk of developing secondary hyperparathyroidism. The downside to the use of this agent is the possibility of raising the serum calcium level. The negative regulation of the PTH gene in response to 1,25(OH)2D3 is mediated by a 684 base pair region upstream of the transcription start site [42]. Another transcriptional regulator, Nuclear factor Y (NF-Y), is an abundant transcription factor and regulates the expression of about 30% of all eukaryotic genes. NF-Y binds to the extended VDRE region of the human PTH gene [43], resulting in enhanced transcriptional activity. Further enhancement is due to additional binding sites for NF-Y, establishing this factor as an important regulator of the expression of the PTH gene and a necessary ingredient for full transcriptional activity of the promoter. Disruption of the synergism between the NF-Y molecules bound to the PTH gene promoter region and modulation of the transcriptional activity of NF-Y by VDR and 1,25(OH)2D3 results in repression of the gene.

D. Other Regulators of PTH Gene Expression Other circulating factors besides calcium, 1,25(OH)2D3 and phosphate may modulate PTH gene transcription. A cAMP response element on the PTH gene suggests that hormones that stimulate adenylyl cyclase activity may increase PTH gene transcription. Another potential regulator is glucocorticoids that increase mRNA levels in dispersed, hyperplastic human parathyroid cells [44] and attenuate the decrease in PTH mRNA in response to 1,25(OH)2D3 in dispersed bovine parathyroid cells [45]. Other intracellular signaling mechanisms are capable of PTH gene regulation. Protein kinases A and C regulate PTH mRNA in vitro in dispersed bovine parathyroid cells [46, 47]. Inhibition of levels of protein kinase C by the addition of staurosporine or a phorbol ester results in decreased levels of PTH mRNA, stimulators of protein kinase A increased PTH mRNA levels. The physiological relevance of the role of intracellular second messengers in the minute-to-minute control of calcium homeostasis remains to be delineated. Estrogen is thought to play a role in the regulation of PTH gene expression due to the proposed roles of PTH in the pathophysiology of postmenopausal osteoporosis. In primary cultures of bovine parathyroid cells, estrogen increases the secretion of PTH. Rat parathyroid glands contain estrogen receptors for estrogen and in ovariectomized rats, estradiol administration increases PTH mRNA [48]. Progesterone has also been demonstrated to enhance secretion of PTH and synthesis of PTH mRNA in cell culture. The addition of 19-nor progestin R5020 to ovariectomized rats results in a two-fold increase in PTH

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mRNA [49]. In the rat, PTH mRNA levels are increased in the proestrus and estrus phase, suggesting alteration of PTH gene expression in response to the rat estrus cycle.

V. BIOSYNTHESIS OF PARATHYROID HORMONE Mammalian PTH is an 84-amino acid polypeptide. The primary amino acid sequence is highly conserved among mammalian species, with strongest homology in the N-terminus. The greatest variation is in the mid-molecule between residues 39 and 52, and the C-terminus contains several stretches of homology [50]. Parathyroid hormone is synthesized in the parathyroid cell as a 115-amino acid single-chain polypeptide termed “preproparathyroid hormone”. The “pre” sequence is the “leader” or “signal” sequence and is identical to signal sequences associated with other polypeptide hormones. The “preproPTH” sequences are known for human, rat, bovine, pig and chicken. All have in common the 25-residue “pre” and 6-residue “pro” sequences. The “pre” sequence contains a hydrophobic stretch of amino acids preceded by a positively charged residue. The signal sequence is cleaved from the amino terminus during the process of synthesis. The signal sequence binds to a signal recognition particle (SRP), an intracellular RNA–protein complex that recognizes the signal sequence. The SRP binds to a receptor on the rough endoplasmic reticulum and directs preproPTH to a protein-lined channel, thereby “docking” the protein. A signal peptidase on the inner surface of the endoplasmic reticulum cleaves the signal sequence. The resultant peptide, “proparathyroid hormone” is left in the cisternae of the endoplasmic reticulum. The “pro” region is a highly positively charged hexapeptide and its cleavage is necessary to initiate biological activity of the hormone [51]. Packaged into vesicles, ProPTH progresses to the Golgi apparatus where the amino-terminal “pro” sequences are removed by the proprotein convertase furin [52]. PTH is concentrated into dense core secretory vesicles that fuse with the plasma membrane to release the contents in response to a fall in extracellular calcium concentration.

VI. METABOLISM OF PARATHYROID HORMONE Proteolysis of newly synthesized PTH is influenced by extracellular calcium [53, 54]. Most of the hormone secreted under conditions of hypercalcemia is already fragmented whereas, in the case of hypocalcemia, the

Chapter 16 Parathyroid Hormone: Structure, Function and Dynamic Actions

secreted molecule tends to be intact and active. Variable extracellular calcium levels, which are similar to mechanisms in proteolytic cleavage of the stored hormone, provides a dynamic regulatory mechanism. Intact PTH is rapidly cleared from the circulation with a half-life of approximately 2 minutes [55]. The liver plays the dominant role in PTH metabolism, removing 60–70% of the hormone. The kidney removes 20–30% by glomerular filtration. Uptake by other cells such as bone cells accounts for a minor component. Clearance by the liver is under the direction of the high-capacity, nonsaturable Kupffer cells where PTH undergoes extensive proteolysis. Some of the C-terminal fragments are released into the bloodstream to be removed ultimately by the kidneys [56]. PTH is not only rapidly cleared from the circulation, but it is also rapidly cleaved by endoproteases resulting in a series of carboxy-terminal fragments that are in the circulation. Carboxy-terminal fragments make up 50–90% of the total circulating PTH immunoactivity, in spite of the fact that, quantitatively, only 10–20% of secreted intact PTH is converted to circulating carboxy-terminal fragments. This is because clearance of these fragments is limited by glomerular filtration mechanisms in the kidney. In renal insufficiency, large amounts of carboxy-terminal fragments are found in the circulation.

VII. RECEPTOR INTERACTIONS OF PARATHYROID HORMONE AND PARATHYROID HORMONE-RELATED PROTEIN The discovery that PTH and PTHrP share a common receptor has generated additional information about the structure–function relationship of these two peptides. The gene for PTHrP is located on chromosome 12 but it is thought that PTH and PTHrP share a common ancestral gene [57]. There is strong homology in the first half of the active amino-terminal core of PTH and PTHrP in that eight of the first 13 amino acid residues are identical. Beyond these first 13 residues, the remainder of the first 34 amino acids differ in sequence, but topographically may fold in ways that are very similar to each other. In the nonhomologous 14–34 portions, substantial threedimensional structure is shared between the two peptides, helping to account for their ability to bind to a common receptor [58–60]. PTHrP is synthesized in adult and fetal tissue and mediates several paracrine/autocrine functions. The physicochemical properties of PTH receptors have been partially characterized with affinity cross-linking PTH ligands and solubilization of the receptor [61].

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Biological properties of PTH and PTHrP have further defined the important residues in the ligand and allowed creation of analogs that can modify the activity of the receptor. PTH(1-34) and PTHrP(1-36) bind to the same receptor with equal affinity [62]. The human PTH receptor binds both PTH and PTHrP but binds to PTH preferentially [63]. Diversity exists in the receptor mRNAs of different lengths that encode the PTH receptor in several tissues and organs, suggesting that certain receptors may specifically bind specific configurations of the PTH molecule.

VIII. STRUCTURE OF THE PTH/PTHrP (PTH1R) RECEPTOR cDNAs encoding the PTH/PTHrP (also designated PTH1R) receptors predict that they would have topographical features in common with other G protein-coupled receptors. The PTH/PTHrP receptors have an aminoterminal extracellular sequence, three extracellular loops, three intracellular loops, and a carboxy-terminal cytoplasmic region and are approximately 590 amino acids in length. The hydrophilic regions are linked by seven relatively hydrophobic membrane-spanning helices (Fig. 2). PTH1Rs from different species are highly homologous; rat and mouse PTH1Rs are 99% identical at the amino acid level and human and rat PTH1Rs are 91% identical. The greatest amino acid differences are limited to discrete areas in the first extracellular loop and to the carboxyterminal sequences. Renal and bone receptors are identical within each species suggesting that the same receptor transcript is present in these two traditional target tissues [63]. The PTH1R is activated equivalently by intact and N-terminal PTH and PTHrP peptides and capable of coupling to several different G-proteins. This ligand–receptor interaction results in activation of multiple signaling pathways concurrently including cAMP, PLC, cytoplasmic calcium, and PKC. The intracellular tail of the receptor couples to a pertussis toxin-sensitive Gi protein that inhibits adenylyl cyclase [64]. The membrane-spanning helices of PTH1Rs are strongly conserved across species and serve a vital role in the signal transduction across the membrane. Studies that utilized mutant or chimeric receptors have generated information about the importance of several domains in the PTH/ PTHrP receptors responsible for ligand-binding, G-protein binding and receptor activation. The PTH1R is capable of stimulating multiple second-messenger pathways. Stably transfected LLC-PK1 porcine cells expressing rat or opposum PTH1Rs exhibited high-affinity binding of PTH, dose-dependent activation of cyclic AMP, and release

280 Primary sequence and typological structure of the human PTH2R. The seven putative transmembrane-spanning regions are indicated based on alignment with other members of the secretin/PTH receptor subfamily. Conserved residues with the opossum kidney PTH1R are indicated by the black circles. Reprinted from Turner et al. [54], with permission.

Figure 2

Chapter 16 Parathyroid Hormone: Structure, Function and Dynamic Actions

of intracellular calcium were evident. The EC50 of the cyclic AMP response was 20–50-fold lower than the intracellular calcium response. PTH also altered cell proliferation that was mimicked by the addition of forskolin, phorbol esters, calcium ionophores, and cAMP analogs. The effect on phosphate transport was only seen when a phorbol ester was added [65]. These data suggest that activation of multiple intracellular pathways is possible with a single ligand. Scanning mutagenetic techniques of the intracellular regions of the receptor have provided evidence for the roles of various intracellular regions critical for receptor activation, receptor–ligand binding and activation of various second messengers. For example, activation of phospholipase C requires one of the intracellular loops. Mutation of residues in the N-terminal region of the third intracellular loop of the opossum PTH1R and expression in COS-7 cells results in normal binding of ligand but impaired adenylyl cyclase and phospholipase C activation. These data suggest that the N-terminal region of the third intracellular loop plays a critical role in the coupling of Gs- and Gq-mediated second-messenger systems [66]. Further studies of the interactions between PTH and PTH1R have taken advantage of mutagenetic approaches. The interaction of PTH(1-34) with PTH1R involves highaffinity binding of the C terminus of the ligand with portions of the receptor’s extracellular N-terminal domain and the extracellular loops in the transmembrane domain. The N-terminus of the ligand interacts with the transmembrane domains to catalyze the G-protein activation(s) required for signal transduction [50]. The critical role in receptor activation of the juxtamembrane (“J”) domain is highlighted by the fact that several mutations in this area result in constitutive (i.e., ligand-independent) receptor activation (Jansen’s metaphyseal chondrodysplasia) [67]. One of the most interesting and provocative studies regarding structure–function relationships between receptors and ligands is evaluation of the cavity formed by the receptor into which the ligands fit. Specific amino acid residues are responsible for optimal ligand binding and signal transduction. The polypeptides secretin and parathyroid hormone were used as models of two ligands with no sequence homology and no cross reactivity with the other’s receptor. Mutation of a single amino acid in the second transmembrane domain of the secretin receptor to the corresponding amino acid in the PTH receptor resulted in a receptor that could bind to and transmit the intracellular signals affected by PTH. The reciprocal mutation in the PTH receptor conferred responsiveness to secretin in a similar manner. Thus, transmembrane residues on receptors recognize a specific ligand and restrict the access and activation of inappropriate ligands [68].

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The PTH1R undergoes internalization after phosphorylation. β-arrestins may terminate receptor–G protein coupling and promote receptor endocytosis, although there may be differences in the recycling of the receptor depending on the cell type [50]. PTH1R internalization can be dissociated from receptor activation and, in some cells, N-truncated PTH peptide such as PTH (7-34) and PTH (7–84) may promote PTH1R endocytosis via a β-arrestin independent mechanism [69].

IX. ACTIVATION OF THE CYCLIC ADENOSINE MONOPHOSPHATE SECOND-MESSENGER SYSTEM BY PARATHYROID HORMONE Parathyroid hormone was one of the first hormones to be shown to utilize the cAMP second-messenger system. In the kidney, cAMP is involved in PTH-mediated effects to reduce calcium excretion, enhance tubular reabsorption of phosphate and to stimulate 1,25(OH)2D3 formation. Cyclic AMP is also believed to be the mediator of many of the actions of PTH on bone [70, 71]. Clinical evidence indicates the important role of cAMP in the mediation of the effects of PTH. Patients suffering from a hereditary syndrome of PTH resistance, pseudohypoparathyroidism type 1, do not respond to exogenously administered PTH with an increase in urinary cAMP [72]. The full-length 84-amino acid polypeptide is not required for activation of adenylyl cyclase. PTH(1-34) is as potent as the intact molecule. Carboxy-terminal fragments are inactive and progressive loss in adenylyl cyclase-stimulating activity occurs with the stepwise deletion of amino acids from the amino-terminal end of PTH(1-34) [73]. PTH(1-25) is also inactive, suggesting that the 25–34 positions are important for the binding of PTH to its receptor. Stepwise removal of the amino acids from the aminoterminal end of the molecule rapidly reduces the adenylyl cyclase-stimulating properties such that PTH(2–34) is a very weak stimulator. Further truncation leads to analogs that are competitive inhibitory, such as PTH(7-34). Activation of cAMP stimulates protein kinase A [74]. The osteoblast PTH receptor is also coupled to Gαq, leading to the activation of phospholipase C (PLC) and hydrolysis of phosphatidylinositol bisphosphonate (PIP2) to inositol triphosphate (IP3) and diacylglycerol (DAG). IP3 binds to specific receptors and mediates the release of calcium from intracellular stores [75, 76]. DAG is a critical cofactor for the activation of PKC isoenzymes [77]. Recent studies have indicated that PTH stimulates hydrolysis of phosphatidylcholine through activation of phospholipase D

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in osteoblast-like cells [78]. Phospholipase D-mediated hydrolysis of phosphatidylcholine produces sustained generation of DAG and more prolonged activation of PKC. Thus, PTH stimulation of phospholipase D is an additional pathway that activates PKC isoenzymes in the parathyroid cell. PKC isoenzymes can regulate PTH-stimulated interleukin-6 (IL-6) promoter activity in osteoblastic cells, suggesting that PKC has an important role in bone remodeling due to the IL-6-mediated bone resorption that can occur [75, 76, 79].

X. IDENTIFICATION OF A SECOND PTH RECEPTOR The classical PTH1R recognizes the amino-terminal portion of PTH and the homologous N-terminus region of PTHrP. Activation of PTH1R is indistinguishable if the ligand is PTH(1-34), PTHrP(1-36) or intact PTH(1-84). The second receptor, known as PTH2R, differs substantially from PTH1R in that PTH2R responds only to PTH as a ligand. Homologs of this receptor have been identified in rat and in zebrafish. It is expressed widely in many tissues, particularly the central nervous system. The true ligand for the PTH2R is probably not PTH but rather, in the central nervous system, a PTH2R-selective activator termed tuberoinfundibular peptide (TIP). This 39-amino acid peptide shows limited sequence homology to PTH and is a selective agonist of PTH2Rs. Selectivity of the receptors for PTH and PTH/PTHrP has been explored by constructing chimeras or PTH1R and PTH2R in which the extracellular amino-terminal domains were exchanged. In cells expressing PTH2R with the PTH1R amino terminus, the cAMP response to PTH or PTHrP is identical. Mutations to the PTH1R sequence were made in each of the seven transmembrane spanning domains. Mutations in the transmembrane domains 3 and 7 resulted in receptors unable to respond to PTHrP [80]. In the ligand itself, position 23 (Trp in PTH and Phe in PTHrP) determines binding selectivity and position 5 (Ile in PTH and His in PTHrP) is responsible for signaling selectivity. To determine the site in PTH2R that discriminates between the two residues at site 5, PTH2R and PTH1R chimeras were constructed, expressed in COS-7 cells and tested for their response to cAMP. Two single residues in the membranespanning/loop region of the receptor determine signaling selectively, Ile244 at the extracellular end of the transmembrane helix 3 and Try318 at the carboxy-terminal portion of extracellular loop 2 [81]. It appears that the C-terminal portion of TIP39 can bind well to PTH1R. A third PTH receptor, PTH3R, has been identified in zebrafish [82]. This receptor is more closely related,

by amino acid sequence, to mammalian PTH1R than to PTH2R. It is uncertain whether this receptor shows preferential affinity for PTH or to PTHrP. Moreover, since a mammalian variant has not yet been identified, its role in human subjects is uncertain. Large portions of the PTH C-terminus are preserved across species, suggesting the possibility of additional independent biological function(s) for this region of the PTH molecule. This putative, distinct PTH receptor with specificity for the carboxyl-terminal region of PTH was suggested by the response of osteoblasts to the C-terminal portion of PTH(53-84) [83, 84]. Several investigators have identified a C-terminal specific radiolabeled ligand that binds to parathyroid cells, osteosarcoma cells, and to clonal osteocytic cells derived from the PTH1R-null mice [85, 86]. Further work has indicated that the important binding determinants are in the region hPTH(24-27) [50]. In another report, the removal of the C-terminal Gln84 residue inhibited the binding and activity of skeletal C-terminal PTHRs [83]. Recently, a unique clonal osteocytic cell line isolated from the null PTH1R mouse was found to express abundant CPTHRs. Various recombinant and synthetic CPTH peptides were used to map ligand determinants of the CPTHR binding and bioavailability [87]. Eight specific, highly conserved amino acid residues within the PTH sequence were identified to play a key role in the affinity of PTH for CPTHR. These amino acids are displayed over three critical domains, and when all are present, result in maximal binding affinity.

XI. PHYSIOLOGICAL ACTIONS OF PTH A. Catabolic Actions of PTH at the Cellular Level The complexity of the skeleton with its many components has made evaluation of the cellular effects of PTH a challenge to elucidate. Both in vivo and in vitro approaches have been utilized. With in vitro studies, the usual physiological milieu is lacking and potentially important intercommunication among various cells as well as cytokines/ growth factor interactions in response to PTH may be missed. With in vivo experiments, such interactions among cell types can be appreciated but it is difficult to assign a particular role, even when using a specific cell type. Thus, both kinds of studies are needed for optimal insight into the mechanism of PTH action in bone. Since PTH can be catabolic to bone, it is reasonable to focus on the osteoclast. However, the osteoclast is relatively inaccessible for direct studies. Our understanding of the catabolic actions of PTH is further diminished by the fact that osteoclasts do not respond directly to PTH. They do

Chapter 16 Parathyroid Hormone: Structure, Function and Dynamic Actions

not contain receptors for PTH, and hence, all interactions appear to be the result of direct effects on the osteoblast [90]. When osteoblasts are stimulated by PTH, it is believed that the subsequent generation of factor or factors result in stimulation of osteoclast function and numbers (for review see references [91, 93]). The initial rapid response to PTH involves the cells lining the endosteal surface of bone [93]. This rapid response (less than 1 hour) that results in the release of calcium into the circulation, is an important regulatory mechanism for the quick correction of hypocalcemia. This initial phase of PTH activity is associated with increased metabolic activity of the osteoclast and does not require new protein synthesis. A second phase of calcium mobilization that is dependent on protein synthesis occurs approximately 24 hours later. This second phase results in an increase in the number and activity of osteoclasts. Current concepts attribute such indirect actions on the osteoclast to direct effects on the osteoblast: incubation of osteoclasts with osteoblast-like cells results in osteoclast activation. Osteoblasts and adjacent bone marrow stromal cells contain PTH1 receptors. The increased number and activity of osteoclasts [94] may also be mediated through cells of the osteoblast lineage such as lining cells [90]. Several clonal, conditionally immortalized PTH-responsive, bone marrow stromal cell lines derived from mouse marrow, were shown to support formation of tartrate-resistant acid phosphatase-positive multinucleated cells in response to PTH. These multinucleated cells contained calcitonin receptors and formed resorption lacunae on dentine slices, consistent with the osteoclast phenotype [95]. The precise mechanism by which PTH stimulates the osteoclast to activate bone resorption remains unknown. Many different enzymes or other factors are released by PTH. They include collagenase, lysosomal hydroxylase, acid phosphatases, carbonic anhydrase, H+, K+-ATPases, Na+–Ca++ exchange systems, cathepsin B, and cysteine proteases. An acidic environment, essential for the protonation of the alkaline salt hydroxyapatite, is formed by the osteoclast when it seals by podosomes the area between the osteoclast membrane and the mineralized surface. Several enzymes are present at this site such as H+, K+-ATPase. Omeprozole blocks this proton pump resulting in inhibition of PTH-mediated bone resorption [96]. Carbonic anhydrase, also activated by PTH, generates hydrogen ion for H+, K+-ATPase activity. PTH-mediated bone resorption is attenuated by inhibition of carbonic anhydrase [97]. Amiloride and its analog, 3′4′-dichlorobenzamil (DCB) affects calcium transport systems such as ATPdependent calcium pumps. These compounds inhibit PTHinduced bone resorption in neonatal mouse calvaria [98]. The mechanism of bone resorption remains controversial,

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however, as other studies suggest that PTH is not required for the buffering of protons in neonatal mouse calvaria [99]. The hypothetical construct suggests that PTH induces catabolic activation in the osteoblast that in turn influences the osteoclast. PTH induces the production of plasmin metalloproteinases such as tissue-type plasminogen activator (tPA) that may initiate bone resorption via breakdown of the extracellular matrix. In neonatal rat osteoblast cell cultures, bovine PTH(1-34) increases tPA by approximately 6–8-fold. After induction of osteoblast differentiation with 1,25-dihydroxyvitamin D3, PTH had no effect on tPA [100]. Other catabolic actions of PTH in the osteoblast include stimulation of hyaluronase synthesis [101] and induction of collagenase-3 gene transcription [102].

B. Anabolic Actions of PTH at the Cellular Level The multiple anabolic actions of PTH on the skeleton are due directly to its effects on osteoblasts that contain receptors for PTH. PTH induces the retraction of preosteoblasts from the bone surface through a calcium-dependent, proteolytic modification of the cytoskeleton. Regulation of cell attachment via regulation of E-cadherins [103] might be a mechanism by which PTH leads to an increase in osteoblast number. PTH may also be mitogenic for bone cells in vivo [104]. The role of PTH in the differentiation of osteoblasts was evaluated in a bone morphogeneic protein (BMP)dependent system using a mesenchymal progenitor cell (C3H10T1/2). Constitutive expression of the PTH/PTHrP receptor resulted in stimulation of osteogenic development. PTH(1-34) stimulated the early and suppressed the late stages of osteogenic development. The effect appeared to be mediated by a cAMP signaling cascade. Other PTH peptides such as PTH(28-48) or PTH(53-84) did not result in significant responses. Parathyroid hormone enhances collagenase synthesis, inhibits type I collagen synthesis, and reduces alkaline phosphatase activity in the osteoblast [105–107]. Further studies have better defined the roles of carboxyl-terminal fragments of PTH versus amino-terminal fragments in these effects. In the osteoblast-like cell line UMR-106, PTH(1-84) and PTH(1-34) inhibited cell proliferation and stimulated alkaline phosphatase activity; C-terminal fragments had no effect. Expression of type I procollagen was stimulated by PTH(35-84), PTH(53-84), and PTH(69-84), inhibited by PTH(1-34), and unaffected by PTH(1-84). These results suggest that the carboxy-terminus region of PTH may contain information in its sequence to influence bone formation [108].

284 The effects of PTH on 24-hydroxylase in osteoblasts are opposite to those in the kidney. In combination with 1,25-dihydroxyvitamin D3, PTH enhances mRNA levels of the 24-hydroxylase cytochrome P450 components. This synergism is in marked contrast to regulation of 24-hydroxylase in the kidney where PTH and 1,25dihydroxyvitamin D have antagonistic effects [109]. Parathyroid hormone alters the nuclear matrix in osteoblasts. The rat type I collagen alpha 1(I) polypeptide chain (COL1A1) promoter confirmation is linked to cell structure via the nuclear matrix. Parathyroid hormone increased the binding of a soluble nuclear protein (NMP4) and decreased COL1A1 mRNA in osteosarcoma cells [110]. The same laboratory demonstrated that PTH can alter osteoblast gene expression via changes in the organization of the nucleus structural proteins [111]. The mechanism by which PTH exerts an anabolic effect is unknown. Nuclear matrix proteins such as NuMA, topoisomerase II-α and -β mediate nuclear organization. NuMA may maintain the structural integrity of the interphase nucleus and form discrete transcriptional domains. Further work on the changes in nuclear matrix in response to PTH has been described. PTH treatment down-regulated the number of topoisomerase II-α-immunopositive cells, and correlated with a decrease in S-phase cells in bone tissues and cultured osteoblast-like cells. The authors suggest that anabolic doses of PTH attenuate the proliferative capacity of osteogenic cells by targeting topoisomerase II-α expression [112]. Other investigators have suggested that an amphiregulin may mediate in part an anabolic effect on osteoblasts. Using microassays to target gene expression profile changes in PTH-treated UMR-105-01 osteoblast osteosarcoma cell line, amphiregulin (AR), a member of the epidermal growth factor family, was identified. AR is bifunctional because it inhibits the growth of several human tumor cells, and stimulates the proliferation of other normal cells such as fibroblasts. PTH highly up-regulates expression of amphiregulin, and is a potent growth factor for preosteoblasts. However, AR is bifunctional in bone, as it inhibits further differentiation of these cells. AR-null mice have less tibial trabecular bone than wild-type mice, suggesting a role of AR in PTH-mediated bone formation and resorption [113].

XII. CELL-TO-CELL COMMUNICATION: OSTEOBLASTS AND OSTEOCLASTS Cell-to-cell communication is enhanced by parathyroid hormone. PTH enhances the formation of gap junctions

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in calvarial osteoblasts [114]. The enhancement of gap junctions, primarily composed of connexin 43 (Cx43), is due to an increased rate of Cx43 gene transcription by PTH. These results, shown for UMR 106 cells, vary when other osteoblast-like cells are studied. The differences may be due to the stage of differentiation of the particular cell model studied. PTH changes the level of Cx43 gene expression in proliferating and maturing osteoblasts but has little effect on nondividing, differentiated osteoblasts [115, 116]. The factors responsible for communication between cells of the osteoblast lineage and those that are osteoclast in nature are the subject of much interest and speculation. Utilizing a human osteosarcoma cell line, SaOS-2, Elias et al. [117] demonstrated that PTH stimulates production of IL-11 protein and mRNA. They proposed this cytokine as the link between the osteoblast and osteoclast. Other investigators have evaluated the effect of IGF-1 in mediating cell signaling between these two bone cells. In newborn rat calvaria, PTH stimulated the production of IGF-1 and IGFBP-3 at the message and peptide levels [118]. Other studies have implicated PTH-induced production of prostaglandins by the osteoblast to induce bone resorption by the osteoclast. In primary cultures of human osteoblastlike cells, PTH has been shown to induce cyclooxygenase-2 gene expression resulting in prostaglandin E2 production [119]. At the molecular level, the influence of PTH on c-fos gene expression in UMR-106 cells was explored [120]. Both PTH(1-34) and PTHrP(1-34) induced c-fos gene expression. Antisense oligonucleotides inhibited PTH-mediated cell proliferation in the osteoblast-like cells and also inhibited PTH-enhanced osteoclast-like cell formation. Those results suggest PTH might regulate bone formation and bone resorption by effects on c-fos gene expression.

A. Nonclassical Actions of PTH PTH promotes the intestinal absorption of calcium though the regulation of renal 1-α-hydroxylation of vitamin D to 1,25(OH)2D3. In pathological or pharmacologic studies, PTH may regulate calcium metabolism via direct stimulation of intestinal calcium absorption [121]. Other nonclassical actions of PTH include the stimulation of gluconeogenesis and alanine uptake in animals [122], a natriuretic and calciuric effect in thyroparathyroidectomized dogs [123], induction of chronotropic effects in rat cardiomyocytes (375), stimulation of intracellular calcium response in pancreatic islets, cardiomyocytes,

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hepatocytes, and proximal renal tubular cells. Most of these actions are consistent with activation of PTH1R.

XIII. PREFERENTIAL ACTIONS OF PTH AT SELECTED SKELETAL SITES A. Animal Models Many studies in animals and in humans have attempted to understand the basis of a selective effect of PTH on cortical bone for catabolic events and on cancellous bone for anabolic events. The effect of PTH on indices of bone formation such as bone mineral density are significantly increased after only 10 days and biomechanical properties improved after 15 days in a rat model [124]. In another study, Wistar rats were administered parathyroid hormone after ovariectomy. The entire skeleton showed an increase in bone mineral content during the period of administration of PTH and no effect of withdrawal was noted. The metaphysis was highly sensitive to PTH and the lumbar vertebrae and diaphysis had moderate changes in bone mass in response to PTH, while the skull and caudal vertebral bodies were not responsive to PTH. After withdrawal of PTH, BMD decreased markedly at the sites that were initially responsive to PTH, readministration of PTH resulted in accretion of BMD at PTHsensitive sites [125]. Weekly administration of PTH to ovariectomized rats effectively stimulated bone formation in both the cancellous and cortical compartments suggesting that administration may not have to be as frequent as originally planned in the initial studies in humans [126]. An ovariectomized rat model was utilized to test the effect of analogs of human PTH(1-34) which differ from the native sequence in that their receptor-activating properties could promote bone formation. Single substitutions for serine in the 3-position result in peptides that are partial agonists in the kidney. Cancellous bone volume was significantly lower in the vehicle-treated group of animals and none of the compounds altered cancellous bone volume. All three peptides produced marked dose-related increases in bone formation rates but the two analogs were less potent than hPTH(1-34). All three peptides produced doserelated increases in osteocalcin. Bone resorption markers had variable effects: pyridinoline cross-link excretion was altered by treatment with hPTH(1-34), but the analog [His]hPTH(1-34) caused a dose-dependent decrease in this parameter of bone resorption [127]. Evidence exists for the role of PTH in the responsiveness of the skeleton to mechanical stress [128]. PTH increased the mechanical strength of the femur diaphysis in

aged rats. In normal rats, mechanical stimulation of the caudal vertebra induced an osteogenic response. In the absence of PTH, this response was attenuated and restored by a single injection of PTH before loading. C-fos expression in osteocytes was detectable only in rats receiving both PTH and mechanical stimulation [129]. One of the most informative models regarding the effect of PTH on the skeleton is a mouse model in which the genes encoding the PTHR1 peptide have been ablated by homologous recombination. These mice have skeletal dysplasia due to accelerated endochondral bone formation and die at birth or in utero. Targeted expression of constitutively active PTH1 receptors resulted in delayed mineralization, decelerated conversion of proliferative chondrocytes into hypertrophic cells, and delay of vascular invasion [130]. Targeted disruption of either PTHr1 [131] or PTHrP [132] in mice and defective PTHrP/PTHR1 signaling in man [133, 134] leads to a lethal form of skeletal dysplasia manifest by accelerated differentiation and decreased proliferation of growth plate chondrocytes. Transgenic mice models are used increasingly to predict skeletal response to PTH. The efficacy of intermittent PTH treatment on bone varies among the tested strains of mice with differences in peak bone mass and structure. To assess the responses of various skeletal sites with high or low cancellous bone mass, mature C57BL/6 mice were ovariectomized or sham operated and treatment with PTH(1-34) or vehicle [135]. After 4–11 weeks of ovariectomy, there was a 44% loss of cancellous bone in the proximal tibia and a 25% loss of cancellous bone in the vertebra, with impaired trabeculae architecture and high bone turnover. In the intact animals, PTH increased cancellous bone volume to a greater extent in the vertebral body compared to the proximal tibia. In the ovariectomized mice, PTH increased cancellous bone volume to a greater extent in the vertebral body, a site with moderate cancellous bone loss, compared to the proximal tibia. Treatment added a small amount of cortical bone to the tibia, but did not significantly increase cortical width of the vertebral body. These comparisons suggest that intermittent PTH is more effective at sites where there is an adequate number of trabecular present at the beginning of the treatment. The authors also proposed that there was an interaction of PTH anabolic action and mechanical loading, since the tibia had a greater increase in cortical bone.

B. Human Studies Several informative studies that address the effect of PTH on bone markers in human subjects were performed

286 in patients with primary or secondary hyperparathyroidism. For a complete description of studies in patients with primary hyperparathyroidism, refer to Chapter 46. In a group of patients with secondary hyperparathyroidism and hypovitaminosis D, treatment with either cholecalciferol or calcitriol resulted in a lowering of N-telopeptide excretion [136]. In a small group of patients with prostate cancer, the excessive bone formation that can occur in metastatic disease results in hypocalcemia and subsequent stimulation of PTH in response to the calcium demand. In these patients, serum bone alkaline phosphatase and urinary levels of bone collagen are markedly elevated. Infusion of the bisphosphonate, pamidronate, significantly decreased serum levels of calcium, phosphorus, bone alkaline phosphatase and urinary markers of bone resorption [137]. In patients with primary hyperparathyroidism, or in normal volunteers receiving PTH infusions (1-38), serum ICTP (C-terminal of type I collagen) was elevated and serum PICP (the carboxy-terminal propeptide of type I collagen) was decreased [138]. Osteoporotic and normal postmenopausal women were studied before and after calcium deprivation to assess responsiveness to endogenous PTH. Baseline values of bone turnover, which included osteocalcin and urinary deoxypyridinoline and pyridinoline cross-link excretion, were higher in a group of postmenopausal osteoporotic women in spite of decreased serum PTH levels as compared to normal postmenopausal women. Calcium deprivation resulted in similar changes in serum levels of calcium, PTH, 1,25(OH)2D3, and pyridinoline and deoxypyridinoline cross-link excretion. Serum osteocalcin increased and serum procollagen carboxy-terminal propeptide decreased in normal women who were calcium-deprived but these bone turnover markers were not altered in the postmenopausal osteoporotic group. These data suggest that there is no difference in the skeletal responsiveness to PTH in patients with osteoporosis [139]. To determine the role of PTH in age-related and nocturnal increases in bone resorption, calcium infusion was utilized to suppress endogenous PTH secretion in young and elderly normal women. Serum PTH and urinary cross-linked N-telopeptide of type I collagen (NTx) were circadian in pattern. Peak levels of PTH occurred in the mid-afternoon and at night, and a rise in urinary NTx was found at night. At baseline, levels of urinary NTx were higher in the elderly compared to young women. Calcium infusion reduced the PTH peaks but did not alter the nocturnal increase in urinary NTx excretion. The authors suggest that PTH is responsible for the age-related increase in bone resorption but does not mediate the circadian pattern of bone resorption [140]. In contrast, administration of recombinant PTH(1-84) to healthy postmenopausal

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women resulted in no change in urinary deoxypryridinoline, although urinary cAMP excretion was appropriately elevated in response to PTH administration [141]. Racial differences exist in the incidence of hip and vertebral fractures: African-American women have a lower incidence of fracture at these sites compared to Caucasian women. One possible mechanism is a difference in the sensitivity of the skeleton to PTH. Cosman and colleagues infused PTH(1-34) in healthy premenopausal black and white women and measured indices of bone turnover. Baseline levels of 1,25(OH)2D3 were significantly lower in black women. There were trends toward higher PTH and lower urinary calcium and pyridinoline levels in this group. No differences in serum calcium or endogenous levels of PTH were noted among the groups during PTH infusion. Black women had lower levels of urinary calcium during the PTH infusion and no differences were detected in bone formation markers among black and white women. Marked differences were noted in markers of bone resorption. Cross-linked N-telopeptides, cross-linked C-telopeptides and free pyridinoline cross-links were much more elevated in white subjects as compared to blacks. These findings suggest that black women have decreased sensitivity and may be an explanation for the relative preservation of skeletal tissue [142]. In postmenopausal women receiving estrogen replacement therapy, daily administration of PTH(1-34) (400 units/ day) over a 3-year period resulted in a continuous increase in bone mineral density compared to no increase in the estrogen-treatment-only control group. The increase in vertebral BMD was 13% while the increase at the hip was less at 2.7%; no loss of bone was noted at any skeletal site tested. Serum osteocalcin measurements were increased 55% during the first 6 months of treatment while a marker of bone resorption, excretion of crosslinked N-telopeptide, increased only by 20%. These changes were interpreted to be due to an uncoupling of the bone formation–resorption cycle and the differences fell toward baseline during the study progression. These data suggest that PTH has potent anabolic effects on the human skeleton [143]. The effects of PTH as an anabolic agent on bone markers is given complete coverage in Chapter 46. In a randomized, double-blind, placebo-controlled trial of idiopathic osteoporosis in men, the effect of administration of parathyroid hormone was evaluated on markers at bone turnover. Bone formation markers that included osteocalcin, carboxyterminal propeptide of type I collagen, and bone-specific alkaline phosphatase, increased in the PTH-treated group. The carboxy-terminal propeptide of type I collagen peaked at 67% above baseline at 6 months and bone-specific alkaline phosphatase peaked at 169% above baseline at 9 months. Osteocalcin was highest (230%) at 1 year.

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PTH administration also dramatically altered bone resorption markers. Pyridinoline was 131% above baseline at 9 months and urinary N-telopeptide was 375% above baseline at 1 year. The relationship between pyridinoline and markers of bone formation were highly correlated in the PTH-treated group but not in the control group. Elevations in bone formation were consistent with a 13.5% increase in bone mass at the lumbar spine in the PTH-treated group at 18 months. Pyridinoline levels at baseline and osteocalcin at 3 months were the best predictors of response to treatment. Parathyroid hormone is the potent stimulator of bone turnover even in men with idiopathic low turnover osteoporosis [144]. Several randomized, placebo-controlled, multicenter studies have evaluated the role of PTH(1-34), teriparatide, as a therapeutic intervention for postmenopausal osteoporosis. In 1637 postmenopausal women with prior vertebral fractures, daily subcutaneous injection of placebo, 20 or 40 µg of teriparatide were administered [145]. The relative risks of fracture compared to the placebo group were 0.35 (95% CI 0.22–0.55) and 0.31 (95% CI 0.19–0.50) in the 20 and 40 µg groups, respectively. New nonvertebral fractures occurred in 6% of women in the placebo group and 3% of each treatment group (RR 0.47 and 0.46, respectively). The 20-µg and 40-µg doses of teriparatide increased BMD by 9 and 13% in the lumbar spine and by 3 and 6% in the femoral neck compared to placebo (20-µg and 40-µg, respectively). In this same trial, the relationship between prior fractures and the risk of new fractures was evaluated [146]. Among the placebo patients with mild, moderate, or severe prevalent vertebral fractures, 10%, 13%, and 28%, respectively, developed vertebral fractures and 4%, 8%, and 23% developed moderate or severe vertebral fractures (both P < 0.001). In the teriparatide-treated group, there were no significant increases in vertebral fracture risk in these subgroups. Similar results were noted regarding nonvertebral fractures. To understand the mechanism by which PTH affects cortical or trabecular bone, a study that evaluated the combination of teriparatide with alendronate or either alone provided additional insights [147]. Using DXA as a measurement, the findings with teriparatide were similar to other trials with increases in BMD at the lumbar spine and total hip, but decreases at the distal one-third radius. Quantitative CT was used to measure volumetric bone mineral density of trabecular bone at the spine and hip and volumetric bone mineral density and geometric variables in cortical bone at the hip. The integral volumetric density (cortical plus trabecular bone) at the spine was similar to the pattern seen in the areal density of the spine. The volumetric density of the trabecular bone at the spine increased markedly, however the increase in the teriparatide

group was approximately twice as great as that found in the alendronate plus teriparatide group. The volumetric density of the trabecular bone at the hip increased with teriparatide administration. In contrast, the volumetric density of cortical bone at the total hip decreased significantly in the parathyroid hormone group.

References 1. Miric, A., and Levine, M. A. (1992). Analysis of the preproPTH gene by denaturing gradient gel electrophoresis in familial isolated hypoparathyroidism. J. Clin. Endocrinol. Metab. 74, 509–516. 2. Kronenberg, H. M., Igarashi, T., Freeman, M. W., Okazaki, T., Brand, S. J., Wiren, K. M., and Potts, J. T., Jr. (1986). Structure and expression of the human parathyroid hormone gene. Recent. Prog. Horm. Res. 42, 641–663. 3. Demay, M. B., Kiernan, M. S., DeLuca, H. F., and Kronenberg, H. M. (1992). Sequences in the human parathyroid hormone gene that bind the 1,25-dihydroxyvitamin D3 receptor and mediate transcriptional repression in response to 1,25-dihydroxyvitamin D3. Proc. Natl Acad. Sci. USA 89, 8097–8101. 4. Kilav, R., Silver, J., and Naveh-Many, T. (1995). Parathyroid hormone gene expression in hypophosphatemic rats. J. Clin. Invest. 96, 327–333. 5. Moallem, E., Silver, J., Kilav, R., and Naveh-Many, T. (1998). RNA protein binding and post-transcriptional regulation of PTH gene expression by calcium and phosphate. J. Biol. Chem. 273, 5253–5259. 6. Antonarakis, S. E., Phillips, J. A., Mallonee, R. L., Kazazian, H. H. J., Fearon, E. R., Waber, P. G., Kronenberg, H. M., Ullrich, A., and Meyers, D. A. (1983). Beta-globin locus is linked to the parathyroid hormone (PTH) locus and lies between the insulin and PTH loci in man. Proc. Natl Acad. Sci. USA 80, 6615–6619. 7. Mayer, H., Breyel, E., Bostock, C., and Schmidtke, J. (1983). Assignment of the human parathyroid hormone gene to chromosome 11. Hum. Genet. 64, 283–285. 8. Zabel, B. U., Kronenberg, H. M., Bell, G. I., and Shows, T. B. (1985). Chromosome mapping of genes on the short arm of human chromosome 11: parathyroid hormone gene is at 11p15 together with the genes for insulin, c-Harvey-ras 1, and b-hemoglobin. Cytogenet. Cell. Genet. 39, 200–205. 9. Miao, D., He, B., Karaplis, A. C., and Goltzman, D. (2002). Parathryoid hormone is essential for normal fetal bone formation. J. Clinical Invest. 109, 1173–1182. 10. Miao, D., He, B., Lanske B., et al. (2004). Skeletal abnormalities in PTH-null mice are influenced by dietary calcium. Endocrinology 145, 2046–2053. 11. Zhou, X-G., Liu, Y-Z., Li, and et al. (2003). Parathyroid hormone gene with bone phenotypes in Chinese. Biochem. Biophys. Res. Comm. 307, 666–671. 12. Goswami, R., Mohappatra, T., Gupta, N., and et al. (2004). Parathyroid hormone gene polymorphism and sporadic idiopathic hypoparathyroidism. J. Clin. Endocrinol. Metab. 89, 4840–4845. 13. Gong, G., Johnson, M. L., Barger-Lux, M. J., and Heaney, R. P. (1999). Association of bone dimensions with a parathyroid hormone gene polymorphism in women. Osteoporosis Int. 9, 307–311. 14. Gohda, T., Shou, I., Fukui, M., and et al. (2002). Parathyroid hormone gene polymorphism and secondary hyperparathyroidism in hemodialysis patients. J. Kidney Dis. 39, 1255–12560. 15. Kanzawa, M., Sugimoto, T., Kobayashi, T., Kobayashi, A., and Chihara, K. (1999). Parathyroid hormone gene polymorphisms in primary hyperparathyroidism. Clin. Endocrinol. 50, 583–588.

288 16. Sunthornthepvarakul, T., Churesigaew, S., and Ngowngarmratana, S. (1999). A novel mutation of the signal peptide of the preproparathyroid hormone gene associated with autosomal recessive familial isolated hypoparathyroidism. J. Clin. Endocrinol. Metab. 84, 3792–3796. 17. Russell, J., Lettieri, D., and Sherwood, L. M. (1983). Direct regulation of calcium of cytoplasmic messenger ribonucleic acid coding for preproparathyroid hormone in isolated bovine parathyroid cells. J. Clin. Invest. 72, 1851–1855. 18. Naveh-Many, T., Friedlander, M. M., Mayer, H., and Silver, J. (1989). Calcium regulates parathyroid hormone messenger ribonucleic acid (mRNA), but not calcitonin mRNA in vivo in the rat. Dominant role of 1,25-dihydroxyvitamin D. Endocrinology 125, 275–280. 19. Brown, E. M. (2005). Editorial: mutant extracellular calcium-sensing receptors and severity of disease. J. Clin. Endocrinol. Metab. 90, 1246–1248. 20. Kifor, O., Diaz, R., Butters, R., and Brown, E. M. (1997). The Ca2+sensing receptor (CaR) activates phospholipases C, A2, and D in bovine parathyroid and CaR-transfected, human embryonic kidney (HEK293) cells. J. Bone Miner. Res. 12, 715–725. 21. Kifor, O., MacLeod, R. J., Diaz, R., Bai, M., Yamaguchi, T., Yao, T., Kifor, I., and Brown, E. M. (2001). Regulation of MAP kinase by calcium-sensing receptor in bovine parathyroid and CaR-transfected HEK293 cells. Am. J. Physiol. Renal. Physiol. 280, F291–F302. 22. Parfitt, A. M. (2001). Parathyroid growth. In “The Parathyroids Basic and Clinical Concepts” (J. P. Bilezikian, R. Marcus, and M. A. Levine, eds), 2nd ed., pp. 293–329. Academic Press, San Diego, CA. 23. Okazaki, T., Zajac, J. D., Igarashi, T., Ogata, E., and Kronenberg, H. M. (1991). Negative regulatory elements in the human parathyroid hormone gene. J. Biol. Chem. 266, 21903–21910. 24. Moallem, E., Kilav, R., Silver, J., and Naveh-Many, T. (1998). RNAprotein binding and post-transcriptional regulation of parathyroid hormone gene expression by calcium and phosphatase. J. Biol. Chem. 273, 5253–5259. 25. Hawa, N. S., O’Riordan, J. L., and Farrow, S. M. (1993). Posttranscriptional regulation of bovine parathyroid hormone synthesis. J. Mol. Endocrinol. 10, 43–49. 26. Vadher, S., Hawa, N. S., O’Riordan, J. L., and Farrow, S. M. (1996). Translational regulation of parathyroid hormone gene expression and RNA: protein interactions. J. Bone Miner. Res. 11, 746–753. 27. Kilav, R., Bell, O., Le, S-Y., Silver, J., and Naveh-Many, T. (2004). The parathyroid hormone mRNA3′-untranslated region AU-rich element is an unstructured functional element. J. Biol. Chem. 279, 2109–2116. 28. Silver, J., Russell, J., and Sherwood, L. M. (1985). Regulation by vitamin D metabolites of messenger ribonucleic acid for preproparathyroid hormone in isolated bovine parathryoid cells. Proc. Natl. Acad. Sci. USA 82, 4270–4273. 29. Russell, J., Lettieri, D., and Sherwood, L. M. (1986). Suppression by 1,25(OH)2D3 of transcription of the pre-proparathyroid hormone gene. Endocrinology 119, 2864–2866. 30. Silver, J., Naveh-Many, T., Mayer, H., Schmelzer, H. J., and Popovtzer, M. M. (1986). Regulation by vitamin D metabolites of parathyroid hormone gene transcription in vivo in the rat. J. Clin. Invest. 78, 1296–1301. 31. Okazaki, T., Igarashi, T., and Kronenberg, H. M. (1988). 5′-flanking region of the parathyroid hormone gene mediates negative regulation by 1,25-(OH)2 vitamin D3. J. Biol. Chem. 263, 2203–2208. 32. Mackey, S. L., Heymont, J. L., Kronenberg, H. M., Demay, M. B. (1996). Vitamin D receptor binding to the negative human parathyroid hormone vitamin D response element does not require the retinoid x receptor. Mol. Endocrinol. 10, 298–305. 33. Sela-Brown, A., Russell, J., Koszewski, N. J., and et al. (1998). Calreticulin inhibits vitamin D’s action on the PTH gene in vitro and may prevent vitamin D’s effect in vivo in hypocalcemic rats. Mol. Endocrinol. 12, 1193–1200.

Lorraine A. Fitzpatrick and John P. Bilezikian 34. Slatopolsky, E., and Bricker, N. S. (1973). The role of phosphorus restriction in the prevention of secondary hyperparathyroidism in chronic renal disease. Kidney Int. 4, 141–145. 35. Lopez-Hilker, S., Dusso, A. S., Rapp, N. S., Martin, K. J., and Slatopolsky, E. (1990). Phosphorus restriction reverses hyperparathyroidism in uremia independent of changes in calcium and calcitriol. Am. J. Physiol. 259, F432–F437. 36. Kilav, R., Silver, J., Biber, J., Murer, H., and Naveh-Many, T. (1995). Coordinate regulation of the rat renal parathyroid hormone receptor mRNA and the Na-Pi cotransporter mRNA and protein. Am. J. Physiol. 268, F1017–F1022. 37. Moallem, E., Kilav, R., Silver, J., and Naveh-Many, T. (1998). RNAProtein binding and post-transcriptional regulation of parathyroid hormone gene expression by calcium and phosphate. J. Biol. Chem. 273, 5253–5259. 38. Almaden, Y., Hernandez, A., Torregrosa, V., Campistol, J., Torres, A., and Rodriguez, M. S. (1995). High phosphorous directly stimulates PTH secretion by human parathyroid tissue. J. Am. Soc. Nephrol. 6, 957. 39. Naveh-Many, T., Marx, R., Keshet, E., Pike, J. W., and Silver, J. (1990). Regulation of 1,25-dihydroxyvitamin D3 receptor gene expression by 1,25-dihydroxyvitamin D3 in the parathyroid in vivo. J. Clin. Invest. 86, 1968–1975. 40. Russell, J., Bar, A., Sherwood, L. M., and Hurwitz, S. (1993). Interaction between calcium and 1,25-dihydroxyvitamin D3 in the regulation of preproparathyroid hormone and vitamin D receptor messenger ribonucleic acid in avian parathyroids. Endocrinology 132, 2639–2644. 41. Brown, A. J., Zhong, M., Finch, J., Ritter, C., and Slatopolsky, E. (1995). The roles of calcium and 1,25-dihydroxyvitamin D3 in the regulation of vitamin D receptor expression by rat parathyroid glands. Endocrinology 136, 1419–1425. 42. Okazaki, T., Igarashi, T., and Kronenberg, H. M. (1988). 5′-Flanking region of the parathyroid hormone gene mediates negative regulation by 1,25-(OH)2 vitamin D3. J. Biol. Chem. 263, 2203–2208. 43. Jääskeläainen, T., Huhtakangas, J., and Mäenpää, P. H. (2005). Negative regulation of human parathyroid hormone gene promoter by vitamin D3 through nuclear factor Y. Biochem. Biophys. Res. Commun. 328, 831–837. 44. Peraldi, M. N., Rondeau, E., Jousset, V., et al. (1990). Dexamethasone increases preproparathyroid hormone messenger RNA in human hyperplastic parathyroid cell in vitro. Eur. J. Clin. Invest. 20, 392–397. 45. Karmali, R., Farrow, S., Hewison, M., Barker, S., and O’Riordan, J. L. H. (1989). Effects of 1,25-dihydroxyvitamin D3 and cortisol on bovine and human parathyroid cells. J. Endocrinol. 123, 137–142. 46. Moallem, E., Silver, J., and Naveh-Many, T. (1995). Regulation of parathyroid hormone messenger RNA levels by protein kinase A and C in bovine parathyroid cells. J. Bone Miner. Res. 10, 447–452. 47. Clarke, B. L., Hassager, C., and Fitzpatrick, L. A. (1993). Regulation of parathyroid hormone release by protein kinase-C is dependent on extracellular calcium in bovine parathyroid cells. Endocrinology 132, 1168–1175. 48. Naveh-Many, T., Almogi, G., Livni, N., and Silver, J. (1992). Estrogen receptors and biologic response in rat parathyroid tissue and C cells. J. Clin. Invest. 90, 2434–2438. 49. Epstein, E., Silver, J., Almogi, G., Livni, N., and Naveh-Many, T. (1996). Parathryoid hormone mRNA levels are increased by progestins and vary during the rat estrous cycle. Am. J. Physiol. 33, E158–E163. 50. Murray, T. M., Rao, L. G., Divieti, P., and Bringhurst, F. R. (2005). Parathyroid hormone secretion and action: evidence for discrete receptors for the carboxyl-terminal region and related biological actions of carboxyl-terminal ligands. Endocrine Rev. 26, 78–113. 51. Kronenberg, H. M., Bringhurst, F. R., Segre, G. V., and Potts, J. T., Jr. (1994). Parathyroid hormone biosynthesis and metabolism. In “The Parathyroids Basic and Clinical Concepts” (J. P. Bilezikian, R. Marcus, and M. A. Levine, eds.), pp. 125–137. Raven Press, Ltd., New York.

Chapter 16 Parathyroid Hormone: Structure, Function and Dynamic Actions 52. Lazure, C., Gauthier, D., Jean, F., Boudreault, A., Seidah, N. G., Bennett, H. P., and Hendy, G. N. (1998). In vitro cleavage of internally quenched fluorogenic human proparathyroid hormone and proparathyroid-related peptide substrates by furin. Generation of a potent inhibitor. J. Biol. Chem. 273, 8572–8580. 53. Habener, J. F., Kemper, B., and Potts, J. T., Jr. (1975). Calcium-dependent intracellular degradation of parathyroid hormone: a possible mechanism for the regulation of hormone stores. Endocrinology 97, 431–441. 54. Chu, L. L. H., MacGregor, R. R., Anast, C. S., et al. (1973). Studies on the biosynthesis of rat parathyroid hormone and proparathyroid hormone: adaptation of the parathyroid gland to dietary restriction of calcium. Endocrinology 93, 915–924. 55. Bringhurst, F. R., Stern, A. M., Yotts, M., Mizrahi, N., Segre, G. V., and Potts, J. T., Jr. (1988). Peripheral metabolism of PTH: fate of biologically active amino terminus in vivo. Am. J. Physiol. 255, E886–E893. 56. Bringhurst, F. R., Segre, G. V., Lampman, G. W., and Potts, J. T., Jr. (1982). Metabolism of parathyroid hormone by Kupffer cells: analysis by reverse-phase high-performance liquid chromatography. Biochemistry 21, 4252–4258. 57. Yasuda, T., Banville, D., Hendy, G. N., and Goltzman, D. (1989). Characterization of the human parathyroid hormone-like peptide gene: functional and evolutionary aspects. J. Biol. Chem. 264, 7720–7725. 58. Caulfield, M. P., McKee, R. L., Goldman, M. E., et al. (1990). The bovine renal parathyroid hormone (PTH) receptor has equal affinity for two different amino acid sequences: the receptor binding domains of PTH and PTH-related protein are located within the 14–34 region. Endocrinology 127, 83–87. 59. Strewler, G. J., Stern, P. H., Jacobs, J. W., et al. (1987). Parathyroid hormone-like protein from human renal carcinoma cells. Structural and functional homology with parathyroid hormone. J. Clin. Invest. 80, 1803–1807. 60. Suva, L. J., Winslow, G. A., Wettenhall, R. E., et al. (1987). A parathyroid hormone-related protein implicated in malignant hypercalcemia, cloning and expression. Science 237, 893–896. 61. Karpf, D. B., Arnaud, C. D., Bambino, T., et al. (1988). Structure properties of the renal parathyroid hormone receptor: hydrodynamic analysis and protease sensitivity. Endocrinology 123, 2611–2619. 62. Jüppner, H., Abou-Samra, A. B., Freeman, M., et al. (1991). A G Proteinlinked receptor for parathyroid hormone and parathyroid hormonerelated peptide. Science 254, 1024–1026. 63. Schipani, E., Karga, H., Karaplis, A. C., et al. (1993). Identical cDNAs encode a human renal and bone parathyroid hormone/parathyroid hormone-related peptide receptor. Endocrinology 132, 2157–2165. 64. Iida-Klein, A., Guo, J., Xie, L. Y., and et al. (1995). Trunication of the carboxyl-terminal region of the rat parathyroid hormone (PTH)/ PTH-related peptide receptor enhances PTH stimulation of the adenylyl cyclase but not phospholipase C. J. Biol. Chem. 270, 8458–8465. 65. Bringhurst, F. R., Juppner, H., Guo, J., Urena, P., Potts, J. T., Jr., Kronenberg, H. M., Abou-Samra, A. B., and Segre, G. V. (1993). Cloned, stably expressed parathyroid hormone (PTH)/PTH-related peptide receptors activate multiple messenger signals and biological responses in LLC-PK1 kidney cells. Endocrinology 132, 2090–2098. 66. Huang, Z., Chen, Y., Pratt, S., Chen, T. H., Bambino, T., Nissenson, R. A., and Shoback, D. M. (1996). The N-terminal region of the third intracellular loop of the parathyroid hormone (PTH)/PTH-related peptide receptor is critical for coupling to cAMP and inositol phosphate/Ca2+ signal transduction pathways. J. Biol. Chem. 271, 33382–33389. 67. Schipani, E., Langman, C. B., Parfitt, A. M., and et al. (1996). Constitutively activated receptors for parathyroid hormone and parathyroid hormone-related peptide in Jansen’s metaphyseal chondrodysplasia. New Engl. J. Med. 335, 708–714. 68. Turner, P. R., Bambino, T., and Nissenson, R. A. (1996). A putative selectivity filter in the G-protein-coupled receptors for parathyroid hormone and secretion. J. Biol. Chem. 271, 9205–9208.

289

69. Sneddon, W. B., Syme, C. A., Bisello, and et al. (2003). Activationindependent parathyroid hormone receptor internalization is regulated by NHERF1 (EBP50). J. Biol. Chem. 278, 43787–43796. 70. Majesca, R. J., Rodan, S. B., and Rodan, G. A. (1980). Parathyroid hormone-responsive clonal cell lines from rat osteosarcoma. Endocrinology 107, 1494–1503. 71. Chase, L. R., Fedak, S. A., and Aurbach, G. D. (1969). Activation of skeletal adenyl cyclase by parathyroid hormone in vitro. Endocrinology 84, 761–768. 72. Chase, L. R., Melson, G. L., and Aurbach, G. D. (1969). Pseudohypoparathyroidism: defective excretion of 3′, 5′-AMP in response to parathyroid hormone. J. Clin. Invest. 48, 1832–1844. 73. Tregear, G. W., Van Rietschoten, J., Greene, E., et al. (1973). Bovine parathyroid hormone: Minimum chain length of synthetic peptide required for biological activity. Endocrinology 93, 1349–1353. 74. Partridge, N. C., Alcorn, D., Michelangeli, V. P., Kemp, B. E., Ryan G. B., and Martin, T. J. (1981). Functional properties of hormonally responsive cultured normal and malignant rat osteoblastic cells. Endocrinology 108, 213–219. 75. Rasmussen, H. (1986). The calcium messenger system (first of two parts). N. Engl. J. Med. 314, 1094–1101. 76. Rasmussen, H. (1986). The calcium messenger system (second of two parts). N. Engl. J. Med. 314, 1164–1170. 77. Ohno, S., Akita, Y., Hata, O., Osada, S., Kubo, K., Konno, Y., and et al. (1991). Structural and functional diversities of a family of signal transducing protein kinases, protein kinase C family, two distinct classes of PKC, conventional cPKC and novel nPKC. Advan. Enzyme Reg. 31, 287–303. 78. Singh, A. T. K., Bhattacharyya, R. S., Radeff, J. M., and Stern, P. H. (2003). Regulation of parathyroid hormone-stimulated phospholipase D in UMR-106 cells by calcium, MAP kinase, and small G Proteins. J. Bone Miner. Res. 18, 1453–1460. 79. Radeff, J. M., Singh, A. T. K., Stern, P. H. (2004). Role of protein kinase A, phospholipase C and phospholipase D in parathyroid hormone receptor regulation of protein kinase Cα and interleukin-6 in UMR-106 osteoblastic cells. Cell Signalling 16, 105–114. 80. Turner, P. R., Mefford, S., Bambino, T., and Nissenson, R. A. (1998). Transmembrane residues together with the amino terminus limit the response of the parathyroid hormone (PTH) 2 receptor to PTH-related peptide. J. Biol. Chem. 273, 3830–3837. 81. Bergwitz, C., Jusseaume, S. A., Luck, M. D., Juppner, H., and Gardella, T. J. (1997) Residues in the membrane-spanning and extracellular loop regions of the parathyroid hormone (PTH) –2 receptor determine signaling selectivity for PTH and PTH-related peptide. J. Biol. Chem. 272, 28861–28868. 82. Gensure, R. C., Ponugoti, B., Gunes, Y., and et al. (2004). Identification and characterization of two parathyroid hormone-like molecules in zebrafish. Endocrinology 145, 1634–1639. 83. Takasu, H., Baba, H., Inomata, N., and et al. (1996). The 69–84 amino acid region of the parathyroid hormone molecule is essential for the interaction of the hormone with the binding sites with carboxyl-terminal specificity. Endocrinology 137, 5537–5543. 84. Murray, T. M., Rao, L. G., and Muzaffar, S. A. (1991). Dexamethasonetreated ROS17/2.8 rat osteosarcoma cells are responsive to human carboxylterminal parathyroid hormone peptide hPTH (53–84): stimulation of alkaline phosphatase. Calcif. Tiss. Int. 49, 120–123. 85. Inomata, N., Akiyama, M., Kubota, N., and Jüppner, H. (1995). Characterization of a novel parathyroid hormone (PTH) receptor with specificity for the carboxyl-terminal region of PTH-(1-84). Endocrinology 136, 4732–4740. 86. Divieti, P., Inomata, N., Chapin, K., Singh, R., Jüppner, H., and Bringhurst, F. R. (2001). Receptors for the carboxyl-terminal region of PTH (1-84) are highly expressed in osteocytic cells. Endocrinology 142, 916–925.

290 87. Divieti, P., Geller, A. I., Sulima, G., Jüppner, H., and Bringhurst, F. R. (2005). Receptors specific for the carboxyl-terminal region of parathyroid hormone on bone-derived cells: determinants of ligand binding and bioactivity. Endocrinology 146, 1863–1870. 88. Eichinger, A., Fiaschi-Taesch, N., Massfelder, T., Fritsch, S., Barthelmebs, M., and Helwig, J. J. (2002). Transcript expression of the turberoinfundibular peptide (TIP)39/PTH2 receptor system and non-PTH1 receptor-mediated tonic effects of TIP39 and other PTH2 receptor ligands in renal vessels. Endocrinology 143, 30336–3043. 89. John, M. R., Arai, M., Rubin, D. A., Jonsson, K. B., and Jüppner, H. (2002). Identification and characterization of the murine and human gene encoding the tuberoinfundibular peptide of 39 residues. Endocrinology 143, 1047–1057. 90. McSheehy, P. M. J., and Chambers, T. J. (1986) Osteoblastic cells mediate osteoclastic responsiveness to parathyroid hormone. Endocrinology 118, 824–828. 91. Martin, T. J. and Udagawa, N. (1998). Hormonal regulation of osteoclast function. Trends Endocrinol. Metab. 9, 6–12. 92. Roodman, G. D. (1998). Osteoclast differentiation and activity. Biochem. Soc. Trans. 26, 7–13. 93. Talmage, R.V., et al. (1978). The demand for bone calcium in maintenance of plasma calcium concentration. In “Mechanisms of Localized Bone Loss” (J. E. Horton, T. M. Tarplay, and W. F. Davis, eds.), pp. 73–91. Information Retrieval, Washington, DC. 94. Rodan, G. A., and Martin, T. J. (1981). Role of osteoblasts in hormonal control of bone resorption – a hypothesis. Calcif. Tiss. Int. 33, 349. 95. Liu, B. Y., Guo, J., Lanske, B., Divieti, P., Kronenberg, H. M., and Bringhurst, F. R. (1998). Conditionally immortalized murine bone marrow stromal cells mediate parathyroid hormone-dependent osteoclastogenesis in vitro. Endocrinology 139, 1952–1964. 96. Tuukkanen, J., and Vaananen, H. K. (1986). Omeprazole, a specific inhibitor of H+-K+-ATPase, inhibits bone resorption in vitro. Calcif. Tiss. Int. 38, 123–125. 97. Hall, G. E., and Kenny, A. D. (1986). Bone resorption induced by parathyroid hormone and dibutyryl cyclic AMP: role of carbonic anhydrase. J. Pharmacol. Exp. Ther. 223, 778–782. 98. Krieger, N. S., and Kim, S. G. (1988). Dichlorobenzamil inhibits stimulated bone resorption in vitro. Endocrinology 122, 415–420. 99. Bushinsky, D. A. (1987). Effects of parathyroid hormone on net proton flux from neonatal mouse calvariae. Am. J. Physiol. 252(4, part 2), F585–F589. 100. Catherwood, B. D., Titus, L., Evans, C. O., Rubin, J., Boden, S. D., and Nanes, M. S. (1994). Increased expression of tissue plasminogen activator messenger ribonucleic acid is an immediate response to parathyroid hormone in neonatal rat osteoblasts. Endocrinology 134, 1429–1436. 101. Midura, R. J., Evanko, S. P., and Hascall, V. C. (1994). Parathyroid hormone stimulates hyaluronan synthesis in an osteoblast-like cell line. J. Biol. Chem. 269, 13200–13206. 102. Selvamurugan, N., Chou, W. Y., Pearman, A. T., Pulumati, M. R., and Partridge, N. C. (1998). Parathyroid hormone regulates the rat collagenase-3 promoter in osteoblastic cells through the cooperative interaction of the activator protein-1 site and the runt domain binding sequence. J. Biol. Chem. 273, 10647–10657. 103. Babich, M., and Foti, L. R. (1994). E-cadherins identified in osteoblastic cells: effects of parathyroid hormone and extracellular calcium on localization. Life Sci. 54, 201–208. 104. Schlüter, K. D., Hellstern, H., Wingender, E., and Mayer, H. (1989). The central part of parathyroid hormone stimulates thymidine incorporation of chondrocytes. J. Biol. Chem. 264, 11087–11092. 105. Hall, A. K., and Dickson, I. R. (1985). The effects of parathyroid hormone on osteoblast-like cells from embryonic chick calcaria. Acta Endocrinol. 108, 217–233.

Lorraine A. Fitzpatrick and John P. Bilezikian 106. Heath, J. K., Atkinson, S. J., Meikle, M. C., and Reynolds, J. J. (1984). Mouse osteoblasts synthesize collagenase in response to bone resorbing agents. Biochim. Biophys. Acta. 802, 151–154. 107. Simon, L. S., Slovik, S. M., Neer, R. M., and Krane, S. M. (1988). Changes in serum levels of type I and III procollagen extension peptides during infusion of human parathyroid hormone fragment (1-34). J. Bone Miner. Res. 3, 241–246. 108. Nasu, M., Sugimoto, T., Kaji, H., et al. (1998). Carboxyl-terminal parathyroid hormone fragments stimulate type-1 procollagen and insulin-like growth factor-binding protein-5 mRNA expression in osteoblastic UMR-106 cells. Endocr. J. 45, 229–234. 109. Armbrecht, J. H., and Hodam, T. L. (1994). Parathyroid hormone and 1,25-dihydroxyvitamin D synergistically induce the 1,25-dihydroxyvitamin D-24-hydroxylase in rat UMR-106 osteoblast-like cells. Biochem. Biophys. Res. Commun. 205, 674–679. 110. Alvarez, M., Thunyakitpisal, P., Morrison, P., Onyia, J., Hock, J., and Bidwell, J. P. (1998). PTH-responsive osteoblast nuclear matrix architectural transcription factor binds to the rat type I collagen promoter. J. Cell Biochem. 69, 336–352. 111. Torrungruang, K., Feister, H., Swartz, D., et al. (1998). Parathyroid hormone regulates the expression of the nuclear mitotic apparatus protein in the osteoblast-like cells. Bone 22, 317–324. 112. Feister, H. A., Onyia, J. E., Miles, R. R., and et al. (2000). The expression of the nuclear matrix proteins NuMA, topoisomerase II-α and -β in bone and osseous cell culture: regulation by parathyroid hormone. Bone 26, 227–234. 113. Qin, L., Tamasis, J., Raggatt, L., and et al. (2005). Amphiregulin is a novel growth factor involved in normal bone development and in the cellular response to parathyroid hormone stimulation. J. Biol. Chem. 280, 3974–3981. 114. Massas, R., and Benayahu, D. (1998). Parathyroid hormone effect on cell-to-cell communication in stromal and osteoblastic cells. J. Cell. Biochem. 69, 81–86. 115. Schiller, P. C., Roos, B. A., and Howard, G. A. (1997). Parathyroid hormone up-regulation of connexin 43 gene expression in osteoblasts depends on cell phenotype. J. Bone Miner. Res. 12, 2005–2013. 116. Civitelli, R., Ziambaras, K., Warlow, P. M., Lecanda, R., Nelson, T., Harley, J., Atal, N., Beyer, E. C., and Steinberg, T. H. (1998). Regulation of connexin43 expression and function by prostaglandin E2 (PGE2) and parathyroid hormone (PTH) in osteoblastic cells. J. Cell Biochem. 68, 8–21. 117. Elias, J. A., Tang, W., and Horowitz, M. C. (1995). Cytokine and hormonal stimulation of human osteosarcoma interleukin-11 production. Endocrinology 136, 489–498. 118. Schmid, C., Schlapfer, I., Peter, M., Boni-Schnetzler, M., and Schwander, J. (1994). Growth hormone and parathyroid hormone stimulate IGFBP-3 in rat osteoblasts. Am. J. Physiol. 267, E226–E233. 119. Conover, C. A. (1995). Insulin-like growth factor binding protein proteolysis in bone cell models. Prog. Growth Factor Res. 6, 301–309. 120. Kano, J., Sugimoto, T., Kanatani, M., Kuroki, Y., Tsukamoto, T., Fukase, M., and Chinara, K. (1994). Second messenger signaling of c-fos gene induction by parathyroid hormeon (PTH) and PTH-related peptide in osteoblastic osteosarcoma cells: its role in osteoblast proliferation and osteoclast-like cell formation. J. Cell. Physiol. 161, 358–366. 121. Nemere, I., and Larsson, D. (2002). Does PTH have a direct effect on intestine? J. Cell. Biochem. 86, 29–34. 122. Hruska, K. A., Blondin, J., Bass, R., et al. (1979). Effect of intact parathyroid hormone on hepatic glucose release in the dog. J. Clin. Invest. 64, 1016–1023. 123. Puschett, J. B., McGowan, J., Fragola, J., Chen, T. C., et al. (1984). Difference between 1-84 parathyroid hormone and the 1-34 fragment on renal tubular calcium transport in the dog. Miner. Electrolyte Metab. 10, 271–274.

Chapter 16 Parathyroid Hormone: Structure, Function and Dynamic Actions 124. Toromanoff, A., Ammann, P., and Riond, J. L. (1998). Early effects of short-term parathyroid hormone administration on bone mass, mineral content, and strength in female rats. Bone 22, 217–223. 125. Kishi, T., Hagino, H., Kishimoto, H., et al. (1998). Bone responses at various skeletal sites to human parathyroid hormone in ovariectomized rats: effects of long-term administration, withdrawal, and readministration. Bone 22, 515–522. 126. Okimoto, N., Tsurukami, H., Okazaki, Y., et al. (1998). Effects of a weekly injection of human parathyroid hormone (1-34) and withdrawal on bone mass, strength, and turnover in mature ovariectomized rats. Bone 22, 523–531. 127. Lane, N. E., Kimmel, D. B., Nilsson, M. H., Cohen, F. E., Newton, S., Nissenson, R. A., and Strewler, G. J. (1996). Bone-selective analogs of human PTH(1-34) increase bone formation in an ovariectomized rat model. J. Bone Miner. Res. 11, 614–625. 128. Ejersted, C., Oxlund, H., Eriksen, E. F., et al. (1998). Withdrawal of parathyroid hormone treatment causes rapid resorption of newly formed vertebral cancellous and endocortical bone in old rats. Bone 23, 43–52. 129. Chow, J. W. M., Fox, S., Jagger, C. J., et al. (1998). Role for parathyroid hormone in mechanical responsiveness of rat bone. Am. J. Physiol. Endocrin. Metab. 37, E146–E154. 130. Schipani, E., Lanske, B., Hunzelman, J., et al. (1997). Targeted expression of constitutively active receptors for parathyroid hormone and parathyroid hormone-related peptide delays endochondral bone formation and rescues mice that lack parathyroid hormone-related peptide. Proc. Natl. Acad. Sci. USA 94, 13689–13694. 131. Karaplis, A. C., Luz, A., Glowacki, J., and et al. (1994). Lethal skeletal dysplasia from targeted disruption of the parathyroid hormone-related peptide gene. Genes Development 8, 277–289. 132. Lanske, B., Karaplis, A. C., Lee, K., and et al. (1996). PTH/PTHrP receptor in early development and indian hedgehog-regulated bone growth. Science 273, 663–666. 133. Jobert, A-S., Zhang, P., Couvineau, A., and et al. (1998). Absence of functional receptors for parathyroid hormone and parathyroid hormone-related peptide in blomstrand chondrodysplasia. J. Clin. Invest. 102, 34–40. 134. Karaplis, A. C., He, B., Nguyen, M. T. A., and et al. (1998). Inactivating mutation in the human parathyroid hormone receptor type 1 gene in blomstrand chondrodysplasia. Endocrinology 139, 5255–5258. 135. Zhou, H., Iida-Klein, A., Lu, S. S., and et al. (2003). Anabolic action of parathyroid hormone on cortical and cancellous bone differs between axial and appendicular skeletal sites in mice. Bone 32, 513–520. 136. Theiler, R., Bischoff, H., Tyndall, A., and Stahelin, H. B. (1998). Elevated PTH levels in hypovitaminosis D are more rapidly suppressed

137.

138.

139.

140.

141.

142.

143.

144.

145.

146.

147.

291

by the administration of 1,25dihydroxy-vitamin D3 than by vitamin D3. Int. J. Vitam. Nutr. Res. 68, 36–41. Berruti, A., Sperone, P., Fasolis, G., Torta, M., Fontana, D., Dogliotti, L., and Angeli, A. (1997). Pamidronate administration improves the secondary hyperparathyroidism due to “Bone Hunger Syndrome” in a patient with osteoblastic metastases from prostate cancer. Prostate 33, 252–255. Brahm, H., Ljunggren, O., Larsson, K., Lindh, E., and Ljunghall, S. (1994) Effects of infusion of parathyroid hormone and primary hyperparathyroidism on formation and breakdown of type I collagen. Calcif. Tissue Int. 55, 412–416. Ebeling, P. R., Jones, J. D., Burritt, M. F., Duerson, C. R., Lane, A. W., Hassager, C., Kumar, R., and Riggs, B. L. (1992). Skeletal responsiveness to endogenous parathyroid hormone in postmenopausal osteoporosis. J. Clin. Endocrinol. Metab. 75, 1033–1038. Ledger, G. A., Burritt, M. F., Kao, P. C., O’Fallon, W. M., Riggs, B. L. and Khosla, S. (1995). Role of parathyroid hormone in mediating nocturnal and age-related increases in bone resorption. J. Clin. Endocrinol. Metab. 80, 3304–3310. Schwietert, H. R., Groen, E. W., Sollie, F. A., and Jonkman, J. H. (1997). Single-dose subcutaneous administration of recombinant human parathyroid hormone [rhPTH(1-84)] in healthy postmenopausal volunteers. Clin. Pharmacol. Ther. 61, 360–376. Cosman, F., Morgan, D. C., Nieves, J. W., Shen, V., Luckey, M. M., Dempster, D. W., Lindsay, R., and Parisien, M. (1997). Resistance to bone resorbing effects of PTH in black women. J. Bone Miner. Res. 12, 958–966. Lindsay, R., Nieves, J., Formica, C., et al. (1997). Randomised controlled study of effect of parathyroid hormone on vertebral-bone mass and fracture incidence among postmenopausal women on oestrogen with osteoporosis. Lancet 350, 550–555. Kurland, E. S., Cosman, F., McMahon, D. J., Shen, V., Lindsay, R., Rosen, C. J., and Bilezikian, J. P. (2000). Changes in bone markers predict bone accural in osteoporotic men treated with parathyroid hormone. J. Bone Miner. Res. 85, 3069–3076. Neer, R. M., Arnaud, C. D., Zanchetta, J. R., and et al. (2001). Effect of parathyroid hormone (1–34) on fractures and bone mineral density in postmenopausal women with osteoporosis. New Engl. J. Med. 344, 1434–1441. Gallagher, J. C., Genant, H. K., Crans, G. G., Vargas, S. J., and Krege, J. H. (2005). Teriparatide reduces the fracture risk associated with increasing number and severity of osteoporotic fractures. J. Clin. Endocrinol. Metab. 90, 1583–1587. Black, D. M., Greenspan, S. L., Ensrud, K. E., Palmero, L., and et al. (2003). The effects of parathyroid hormone and alendronate alone or in combination in postmenopausal osteoporosis. New Engl. J. Med. 349, 1207–1215.

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Chapter 17

Interaction of Parathyroid Hormone-related Peptide with the Skeleton David Goltzman

Calcium Research Laboratory, Department of Medicine, Royal Victoria Hospital, McGill University, Montreal, H3A 1A1, Canada

I. Abstract II. Introduction III. Molecular Biology and Mechanism of Action

IV. The Skeletal Actions of PTHrP V. Summary References

I. ABSTRACT

lineage, resulting in enhanced bone formation and also, indirectly, in augmented osteoclastic bone resorption; however, unlike PTH which serves postnatally to defend against a decrease in extracellular calcium, endogenous PTHrP appears to play an important anabolic role in the postnatal skeleton. Overall, these findings point to a critical role for PTHrP in normal skeletal development and in its maintenance. Further elucidation of the molecular role and interactions of PTHrP may therefore reveal new facets of the pathogenesis of a variety of metabolic bone diseases and potentially point to new directions for therapeutic interventions.

Parathyroid hormone-related peptide (PTHrP) was discovered as a mediator of hypercalcemia associated with malignancy but is now known to be expressed by a large number of normal fetal and adult studies. The aminoterminal region of PTHrP reveals limited but significant homology with parathyroid hormone (PTH), resulting in the interaction of the first 34–36 residues of either peptide with a single seven-transmembrane spanning G proteinlinked receptor termed the PTH/PTHrP receptor or the PTH-1 receptor. PTHrP plays a critical role in chondrocyte biology and in endochondral bone formation. Deficient PTHrP action in utero results in a distorted epiphyseal growth plate and marked skeletal deformity with ensuing neonatal death due to respiratory compromise. The complex processes resulting in normal endochondral bone development involve additional factors such as the hedgehogsignaling pathway with which PTHrP interacts. PTHrP, like PTH, binds to receptors on cells of the osteoblast Dynamics of Bone and Cartilage Metabolism

II. INTRODUCTION The association of malignancies and hypercalcemia was first reported in 1936 [1] and the production and secretion by tumors of a circulating factor causing hypercalcemia was postulated by Fuller Albright in 1941 [2]. 293

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The hypercalcemia of malignancy syndrome is characterized by hypercalcemia and hypophosphatemia, biochemical features that are usually indicative of oversecretion of parathyroid hormone (PTH), the major regulator of calcium homeostasis. Initial studies that searched for immunoreactive PTH within the tumors, however, were equivocal or negative. However, PTH-like activity in tumor extracts was evident in assays that measured renal glucose-6-phosphate dehydrogenase (G6PD) activity by a cytochemical technique [3] and adenylate cyclase activity in osteoblast cell lines and in renal membranes. The latter assay proved to be critical for monitoring the purification and the final isolation of the PTH-like peptide from human lung [4], breast [5], and kidney [6] cancer cell lines. The finding of structural similarity of the amino-terminal region of this peptide (PTHrP, for PTH-related peptide) with the corresponding domain of PTH, and the resultant capacity of both molecules to interact with the same G protein-coupled cell-surface receptor (PTH/PTHrP or PTH-1 receptor) via this region [7, 8], appear to account for the ability of circulating, tumor-derived PTHrP to inappropriately activate receptors in classic PTH target organs such as bone and kidney and to elicit PTH-like bioactivity. By the same mechanism, PTHrP may also enhance osteoclastic activity when released locally by tumor cells that have already metastasized to bone [9, 10]. Further studies, however, have demonstrated the importance of PTHrP, not only as an osteolytic factor in malignancy but also as a paracrine/ autocrine, and probably an intracrine regulator, of many

Figure 1

physiologic processes, most notably of bone and cartilage metabolism. The pivotal role of the PTHrP signaling pathway in endochondral bone development and in adult skeletal homeostasis will be the focus of this review.

III. MOLECULAR BIOLOGY AND MECHANISM OF ACTION A. Organization of the PTHrP Gene The human gene for PTHrP has been mapped to the short arm of chromosome 12 [11], whereas the PTH gene is assigned to human chromosome 11. An evolutionary relationship between the PTHrP and PTH genes has been suggested, because human chromosomes 11 and 12 are thought to have arisen by a tetraploidization event of a common ancestral chromosome. Indeed, the PTHrP and PTH genes and their respective gene clusters have been maintained as syntetic groups in human, rat, and mouse genomes [12]. Considerably more complex than the PTH gene, however, the human PTHrP gene comprises at least seven exons, spans more than 15 kb of genomic DNA (Fig. 1), and utilizes at least three distinct promoters, two containing a classical TATA box and one including a GC-rich region [13–16]. Exon 1, subdivided into Ia, Ib, and Ic, and exon II encode different 5′ untranslated regions. Exon III encodes the prepro coding region and exon IV encodes most of the

Organization of the human, rat, mouse and chicken PTHrP genes. Bold numbers depict numbering of exons. Nonbold numbers depict amino acid positions in the precursor (denoted by – symbols) or in the mature peptide. Arrows indicate promoters. Open boxes indicate 5′ untranslated regions, dark gray boxes denote coding exons and light gray boxes denote exons encoding 3′ untranslated sequences. Exons joined by solid Vs denote obligatory splicing whereas exons joined by broken Vs denote alternative splicing.

Chapter 17 Interaction of Parathyroid Hormone-related Peptide with the Skeleton

mature peptide sequence. At the end of exon IV, the splice site interrupts codon exon IV, encodes a stop codon and a 3′ noncoding sequence. Exon VI encodes 36 additional amino acids, a stop codon, and a second 3′ noncoding region. Exon VII encodes two extra amino acids, a stop codon, and a third noncoding region. Hence, by the use of alternative splicing of exons, a multitude of human PTHrP RNA transcripts can occur, giving rise to three different isoforms of 139, 141, and 173 amino acids in length which are identical as far as amino acid 139. In contrast with this complexity, the rat [17], mouse [18], and chicken [19] PTHrP genes are considerably simpler in organization and consist of four or five exons and only a single promoter, corresponding to the downstream promoter in the human gene. Overall, this simpler organization predicts that, in most species, a single promoter will be found and the 141-amino acids isoform of the peptide will predominate. In the mouse, the mature peptide is 139 amino acids rather than 141 amino acids because of a sixbase pair deletion (corresponding to amino acids 130 and 131 of human and rat protein). Moreover, the chicken gene can encode two isoforms, one of 139 amino acids arising by read-through of exon III into exon IV, and one of 141 amino acids, which is the predominant form, arising from splicing to exon V. Human, rat, mouse, dog [20], and chicken peptides are virtually identical in the amino terminal and midregion of the molecule but diverge beyond residue 112. This striking conservation throughout this large evolutionary range suggested early on that PTHrP is of essential biological importance.

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increased PTHrP expression in the uterus, as does intrauterine occupancy in preterm myometrium [28], however the precise mechanism of these effects is unknown. Both stimulation and inhibition generally occur predominantly at the transcriptional level. 2. Post-translational Processing

With the availability of the PTHrP cDNA-based amino acid sequence, it became evident that the mature form of the protein is preceded by a 36-residue prepro sequence. This sequence has all the classic features of a signal peptide that would target the nascent protein to the rough endoplasmic reticulum. While the precise site of cleavage by the signal peptidase has yet to be determined, it is likely that it occurs at position −9, thereby generating a nascent propeptide extending from position –8. The evidence that the pro sequence is subsequently cleaved to generate the mature form of the protein at position Ala +1 is based on (1) direct sequencing of the secreted PTHrP form by three independent laboratories, (2) the presence of an endoproteolytic site for prohormone convertases of the furin and PC1/3 family proximal to it [29], and (3) the homology of this region with that of PTH. Closer examination of the mature PTHrP sequence reveals additional clusters of basic amino acids that could potentially serve as endoproteolytic processing sites (Fig. 2). Hence, cleavage appears to occur carboxy-terminal to arginine at position 37, most likely yielding the secretory form, PTHrP (1–36). A midregion fragment that begins at amino acid 38 and terminates in the 80–100 region has also been identified, both within cells and outside cells following

B. Regulation of PTHrP Production In contrast to PTH, which is synthesized almost exclusively in the parathyroid glands, PTHrP is expressed in many fetal and adult cells and tissues. 1. Regulation of Gene Expression

Several factors have been shown to regulate PTHrP gene expression in a variety of normal tissues or cells, at least in part, at the transcriptional level. In most cells, including for example cultured keratinocytes, breast epithelial cells, and smooth muscle cells, growth factors, such as EGF [21], IGF1 [22], and TGFb, stimulate PTHrP gene expression [23], whereas 1,25 (OH)2 vitamin D3 and low calcemic analogs of 1,25(OH)2 vitamin D3 inhibit this production [21, 24]. Other steroids including glucocorticoids, androgens [25] and estrogens [26] have also been shown to inhibit PTHrP expression in tissues expressing their cognate receptors. PTHP mRNA in lactating mammary tissue is stimulated by prolactin [27], and estrogen administration causes

Figure 2 PTHrP as a polyhormone. Distinct biological activities have been associated with the different regions of the peptide which are depicted. Numbers within the molecule indicate the sites where proteolytic processing is believed to occur. Numbers at the carboxyl terminal region (139, 141, 173) denote alternative carboxyl terminal amino acids.

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secretion [30]. An additional carboxy-terminal PTHrP fragment may exist that encompasses the highly conserved pentapeptide 107–111. It would seem, therefore, that PTHrP could function as a polyhormone precursor of a number of biologically active fragments with distinct biological actions. 3. Regulation of PTHrP Secretion

Both constitutive and regulated secretion of PTHrP have been described. Elevated calcium concentrations have been reported to stimulate PTHrP secretion via the calciumsensing receptor (CaSR) in prostate, breast, and Leydig cell cancer cell lines. Thus, calcium-activated CaSR may transactivate the EGF receptor by activation of a matrix metalloproteinase, which leads to cleavage of proheparin-binding EGF to heparin-binding EGF. This then leads to phosphorylation of EGF receptor kinase (ERK) 1/2 and resultant PTHrP secretion [31]. In contrast, in lactating mammary epithelium, low calcium levels have been reported to stimulate PTHrP secretion via the CaSR [32].

C. Receptors and Signal Transduction Pathways In view of the extensive post-translational processing that PTHrP undergoes, it would not be surprising if multiple receptors for the various forms of the protein exist. These can be divided primarily into four categories that recognize determinants in (1) the animo-terminal region of the protein, (2) the midregion form, (3) the nuclear localization sequence, and (4) the carboxyl-terminal form. The amino-terminal domain of PTHrP, because of limited but important homology with eight of the first 13 residues in PTH, binds to the classical PTH-1 receptor (PTHR-1) and activates both the adenylyl cyclase/protein kinase A pathway as well as the calcium/inositol phosphate/ protein kinase C pathway (Fig. 3). More recently, evidence for PTHR-1 stimulation of mitogen-activated protein kinase (MAPK) has also been reported [33, 34] and the sodium/hydrogen exchanger regulatory factor (NHERF) has been reported to interact with the carboxyl-terminal intracellular tail of the PTHR-1 to increase the formation of inositol triphosphate while reducing cyclic AMP formation [35]. The interaction between PTHrP and PTHR-1 is sufficient to confer functions of PTHrP which mimic those of PTH in bone and kidney. In fact, most of the well-characterized actions of PTHrP appear to be mediated by this receptor for the NH2-terminal domain of PTHrP. The expression of PTHR-1 has been observed in a wide variety of tissues that might well be PTHrP targets, although expression of the receptor seems highest in bone, cartilage, and kidney.

Figure 3

Interaction of PTHrP with the PTHR-1. The amino terminal domain of PTHrP (and PTH) binds to the seven-transmembrane spanning receptor, causing the guanyl nucleotide binding proteins Gsα and Gqα to activate adenylyl cyclase and phospholipase C respectively. Therefore cAMP or inositol triphosphate (IP3) and diacylglycerol (DAG) are produced. Cyclic AMP and DAG will stimulate protein kinase A (PKA) and protein kinase C (PKC) respectively and IP3 will increase levels of intracellular calcium (CA++). The βγ subunits of Gs and Gq, as well as PKC may also participate in increasing levels of mitogen-activated protein kinase (MAPK). The sodium/hydrogen exchanger regulatory factor (NHERF) appears to associate with the PTHR-1 to increase IP3 and may also reduce adenylyl cyclase activity by interacting with the Gi/o protein.

The existence of a “receptor” that recognizes the N-terminal domain of PTHrP but is distinct from the classic type 1 receptor has also been suggested [36, 37]. This high-capacity, low-affinity receptor was identified in keratinocytes and pancreatic β-cells and signals only through the protein kinase C pathway. Cloning and molecular characterization of this receptor, however, have not been accomplished. Even less is known about a putative receptor that binds the midregion form of PTHrP. The existence of such a receptor, likely linked to the protein kinase C pathway, is circumstantial and is based primarily on the finding that residues 67–86 of PTHrP are critical for the control of placental calcium transport [38]. While the receptor has not yet been identified, support for its existence has come from work with the PTHrP gene knockout mouse in which placental calcium transport is severely impaired and can only be restored by administration of PTHrP(1-86) and PTHrP(67-86) peptides. PTHrP(1-34) and PTH(1-84) have no effect [39]. Amino acids 87–107 of the mature form of PTHrP encode a nuclear localization signal (NLS) consisting of two basic clusters separated by a spacer region, analogous to the prototypical nucleoplasmin NLS [40, 41]. Studies have shown that elimination of the NLS from prepro

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PTHrP effectively abolished intranuclear localization of the recombinant protein. Moreover, the NLS alone was capable of translocating a heterologous cytoplasmic protein, β-galactosidase, to the nucleolus when expressed as a fusion protein in COS-7 cells. Selective protein import into the nucleus proceeds through the nuclear pore complex and requires the interaction between the NLS and an importin–beta1 receptor complex [42]. Receptors for the carboxyl-terminal form of PTHrP have been inferred from studies reporting that picomolar concentrations of this peptide can profoundly inhibit resorption in an isolated osteoclast bone resorption assay [43, 44]. In addition, the C-terminal peptide PTHrP(107139) has been shown to elicit a rise in cytosolic calcium in cultured hypocampal neurons [45]. It is possible, therefore, that novel receptors for carboxyl-terminal fragments might be expected in osteoclasts and neurons. The cloning and characterization of these receptors will be necessary in order to establish the physiological relevance associated with their activation.

D. Endocrine, Autocrine/Paracrine, and Intracrine Actions Although PTHrP was initially discovered through its “endocrine” effects (i.e., it is released from tumors into the circulation and it acts at distant sites such as the skeleton and kidneys), this mechanism of action is generally considered to be the exception rather than the rule. In fact, under normal circumstances, the only other endocrine effect ascribed to PTHrP is its influence on placental calcium transport, presumably following release from the fetal parathyroids. This scenario notwithstanding, PTHrP, unlike PTH, does not circulate in appreciable amounts in normal subjects [46]; rather, it is thought to exert its biological effects locally in a paracrine/autocrine fashion and in this manner may influence growth, differentiation, and differentiated function in a variety of cell types. For example, studies in cultured keratinocytes [40, 47] demonstrated a role for PTHrP in inhibiting growth and supporting differentiation, and overexpression of PTHrP in breast myoepithelial cells decreased ductal proliferation and branching morphogenesis [48, 49]. In some tissues such as mammary tissue, PTHrP is an epithelial-derived factor that acts in a paracrine manner on mesenchymal cells expressing the PTH-1 receptor to influence mammary gland development [49]. Similarly, PTHrP derived from the endothelium can act on vascular smooth muscle cells to regulate vascular tone and blood pressure. Alternatively, PTHrP can be secreted by vascular smooth muscle cells, and in turn, feed back in

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an autocrine mode to alter the contractility of these cells. Likewise, keratinocytes, pancreatic β cells, and uterine smooth muscle cells can both secrete and respond to PTHrP. Furthermore, PTHrP has also been reported to alter differentiated functions, such as smooth muscle relaxation and neurotransmission. In addition to the autocrine/paracrine effects, experimental evidence has been presented suggesting that PTHrP may also have an intracrine site of action. This conclusion was first based on the following observations: (1) endogenous PTHrP localizes to the nucleus of murine bone cells in vitro as well as in situ [46], (2) immunogold labeling for the endogenous peptide in situ was observed over the dense fibrillar component of nucleoli, a subnucleolar structure thought to represent the major sight of transcription of gene coding for rRNA, and (3) the nuclear actions of PTHrP have been subsequently confirmed in a human keratinocyte cell line (HaCaT) [50] and in cultured vascular smooth muscle cells [51]. In view of the fact that PTHrP contains a leader sequence, which directs the nascent peptide to the secretory apparatus of the cell in which it is biosynthesized, it is uncertain how such a peptide might be trafficked to the cytoplasm and have access to the nucleus. Three possibilities have been described. One is that PTHrP undergoes reverse transport from the endoplasmic reticulum to the cytoplasm, with at least some of PTHrP becoming subject to ubiquitination and proteosomal degradation [52]. A second is that, after secretion, PTHrP is internalized by the PTHR-1 [53] which itself has been reported to contain a nuclear localization signal, or by a distinct membrane receptor which has not yet been characterized [54]. In either case, how PTHrP would escape from endosomes subsequent to receptor-mediated endocytosis has not yet been determined. A third and perhaps the most likely possibility, is that, as has been demonstrated, some of the PTHrP is translated from an alternative non-AUG start site downstream from the initiator methionine, which excludes the leader sequence but allows the retention of the peptide, including the nuclear localization signal within the cell cytoplasm [55]. Studies in vascular smooth muscle cells [56] and keratinocytes [57] have shown that PTHrP expression occurs in a cell cycle-dependent manner with higher expression occurring in the G2 and M phases of the cycle. The cell cycle-dependent localization of PTHrP is regulated by the activity of the cyclin-dependent kinases (cdk) cdc2 and cdk2, which phosphorylate PTHrP at threonine85 within a consensus cdc2/cdk2 site. Phosphorylation increases as cells progress from G1 to G2 and M of the cell cycle and leads to decreased nuclear entry, perhaps by enhancing binding to a cytoplasmic retention factor.

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PTHrP appears to bind with high affinity to importin b1 and the GDP-bound protein, Ran. After translocation across the nuclear envelope, GTP-bound Ran may release PTHrP into the nucleus where it may act in the nucleolus to bind RNA [58], which may possibly regulate mRNA processing or mRNA transport. Functional effects of intranuclear PTHrP may then be to delay apoptosis, as shown in chondrocytic cells in vitro and to increase cell proliferation in vascular smooth muscle in vitro [51]. Further studies are, however, required to demonstrate the effects of nuclear localization of PTHrP in vivo. It would appear, therefore, that PTHrP influences a spectrum of cellular function by exerting its effects at, near, or in its site of synthesis.

IV. THE SKELETAL ACTIONS OF PTHrP A. PTHrP Signaling and Chondrocyte Biology 1. Chondrocyte Proliferation and Differentiation

Endochondral ossification is a complex, multistep process involving the formation of cartilaginous skeleton from aggregated mesenchymal cells and its subsequent replacement by bone. The cellular and molecular events that regulate the highly ordered progression of chondrocytes within the growth plate through the stages of proliferation and differentiation must be under precise spatial and temporal control. Ultimately, these events determine the extent and rate of skeletal growth. In the cartilaginous growth plate, PTHrP is expressed by perichondral, proliferating, and hypertrophic chondrocytes, and the PTHR-1 is expressed in proliferating and hypertrophic chondrocytes. The generation of mice homozygous for a disrupted PTHrP gene provided the first direct evidence of a physiological role for this protein in chondrocyte biology [59, 60]. PTHrP-null mice die in the immediate postnatal period, likely from respiratory failure in association with widespread abnormalities of endochondral bone development. Characterized by diminished proliferation, accelerated differentiation, and premature apoptotic death of chondrocytes, this form of ostoechondrodysplasia results in the untimely and rapid maturation of the skeleton [59, 60]. The critical role of PTHrP as an inhibitor of the chondrocyte differentiation program [61] has been further substantiated by the targeted overexpression of PTHrP in chondrocytes by means of the mouse collagen type II promoter [62]. This targeting induces a novel form of chondrodysplasia that is the antithesis of the PTHrP-null

phenotype and is characterized by a delay in endochondral ossification so profound that mice are born with cartilaginous endochondral skeletons. Therefore, PTHrP enhances chondrocyte proliferation and negatively regulates the switch from a proliferative immature chondrocyte to a post proliferative mature, hypertrophic chondrocyte. PTHrP blocks chondrocyte differentiation via the cAMP pathway and this process may involve phosphorylation and activation of the transcription factor SOX9, a master regulator of early chondrogenesis. 2. PTHrP and Chondrocyte Apoptosis

Hypertrophic chondrocytes in the growth plate are thought to undergo apoptosis immediately prior to ossification [63, 64] and, therefore, represent the terminal stage of differentiation in the chondrogenic lineage. PTHrP expression might be expected to delay, or even prevent, the progression to terminal differentiation and eventual programmed cell death. Studies with the chondrocytic cell line CFK2 overexpressing PTHrP demonstrated enhanced cell survival under conditions that prompted is apoptotic death [41]. In addition, quantitative analysis of the growth plate of PTHrP-null mice revealed significantly more apoptotic chondrocytes near the chondro-osseous junction compared to wild-type littermates [63]. Thus, PTHrP influences not only chondrocyte proliferation and differentiation, but also programmed cell death. In several cell types, apoptosis is regulated by the ratio of expression of the cell death inhibitor, Bcl-2, and the cell death inducer, Bax. Bcl-2 is expressed in growth plate chondrocytes in a pattern similar to that of PTHrP [65], with the highest levels detected in late proliferative and prehypertrophic chondrocyte. Both in vitro and in transgenic mice, PTHrP overexpression causes a marked increase in Bcl-2 with no detectable change in Bax levels. A shift of the Bcl-2 downstream of the Bcl-2/Bax ratio in favor of Bcl-2 delays terminal differentiation, prolongs chondrocyte survival, and leads to the accumulation of cells in their prehypertrophic stage. These observations place Bcl-2 downstream of PTHrP in the pathway that controls chondrocyte maturation and endochondral skeletal development. 3. INTERACTION OF PTHrP WITH OTHER CARTILAGE REGULATORY FACTORS

Indian hedgehog (Ihh), a member of a family of proteins important for embryonic patterning, is highly expressed in the prehypertrophic zone between proliferating and hypertrophic chondrocytes and in the upper hypertrophic chondrocytes [66, 67]. Ihh seems, in growth plate cartilage, to be necessary and sufficient for PTHrP gene expression (Fig. 4).

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and is essential for vascular invasion at the chondro-osseous junction. Runx2 can activate the Ihh/PTHrP pathway but at present it is unclear whether PTHrP alters Runx2 expression in chondrocytes. 4. Defining the Role of the PTH-1 Receptor

Figure 4 Effect of PTHrP and other factors on chondrocyte proliferation and differentiation in the growth plate. Stimulatory effects of the factors are shown by the solid arrows (t) and inhibitory effect by the solid symbol, . The effects of Indian hedgehog (Ihh), bone morphogenetic protein (BMP), the transcription factor Runx2, and of fibroblast growth factor (FGF) on each other are depicted by hatched arrows ( ) when stimulatory and by the hatched symbol, , when inhibitory.

Ihh stimulation of PTHrP therefore increases chondrocyte proliferation and decreases chondrocyte hypertrophy. This therefore reduces Ihh production, resulting in a negative feedback loop. Ihh, independently from PTHrP, also stimulates chondrocyte proliferation but may also enhance chondrocyte differentiation by blocking the effects of PTHrP [68]. The fibroblast growth factor (FGF) receptor, FGFR3, a tyrosine kinase receptor, is expressed in proliferating chondrocytes and can inhibit Ihh expression, thereby indirectly reducing PTHrP expression. Activation of FGFR3 inhibits chondrocyte proliferation independent of its effect on Ihh but may also increase chondrocyte differentiation, apoptosis, and vascular invasion at the chondro-osseous junction. FGF18 appears to be a major ligand for FGFR3 [69]. Studies with double mutants lacking both FGFR3 and PTHrP indicate that the stimulatory effect of PTHrP on chondrocyte proliferation supercedes the inhibitory effect of FGFR3. In contrast to FGF which inhibits Ihh, bone morphogenetic proteins (BMPs) form a positive autoregulatory feedback loop with Ihh, although a role for BMPs in stimulating proliferation and inhibiting differentiation independent of the Ihh/PTHrP axis has been suggested [66]. The transcription factor Runx2 (also known as Cbfa1) is expressed in early hypertrophic chondrocytes and enhances chondrocyte maturation in co-operation with Runx3 and/or core-binding factor b (CBFb), a co-transcription factor that forms heterodimers with Runx proteins. Runx2 also activates vascular endothelial growth factor (VEGF)

The PTH-1 receptor appears to mediate most PTHrP actions in the cartilaginous growth plate. Thus, the generation of PTH-1 receptor knockout mice provided the first evidence that this receptor mediates the cartilaginous effects of PTHrP [67]. The small number of homozygous mice that do survive to the peripartum period exhibit a phenotype similar to that observed in the PTHrP-negative mutants, although they are proportionally smaller. Skeletal alterations are characterized by accelerated differentiation of chondrocytes leading to a severe and lethal form of osteochondrodysplasia. In contrast, targeted overexpression of constitutively active PTH-1 receptor to the growth plate delays endochondral bone formation [70]. Of note, however, is the observation that most PTHR1-negative animals exhibit a more severe phenotype that the PTHrP-null mice characterized by early embryonic lethality (embryonic day E14.5). It was originally speculated that PTHrP synthesized by maternal decidual cells could complement the absence of fetal PTHrP but not that of its receptor. Alternatively, the more severe phenotype may implicate the presence of another member of the PTH/PTHrP ligand family that interacts with the common receptor. The existence of such a protein, however, remains speculative at present. Perhaps the most intriguing explanation for the early demise of these animals implicates a possible dual mechanism of action of PTHrP on cellular function, i.e., the amino-terminal end of the protein acting on the cell-surface receptor and the more carboxyl region acting within the nucleus. In the absence of the amino-terminal receptor, the unopposed nuclear effects of PTHrP would lead to severe imbalance of cellular proliferation and differentiation and hence to early lethality of the receptor-negative mutants. In contrast, ablation of the ligand would eliminate both membrane receptor- and nuclear-mediated activities, leading to a more “coordinated” impairment and a less severe phenotype. At present, however, it is reasonable to conclude that the small size and early death of the PTH-1 receptor-null mice remains unexplained. The critical role of the PTHR-1 in endochondral bone formation is emphasized, however, by the discovery that two human chondro-osseous dysplasias, Blomstrand’s lethal chondrodysplasia (BLC) [71, 72] and Jansen’s metaphyseal chondrodysplasia (JMC) are caused by receptor mutations. BLC appears to be autosomal recessive, is caused by inactivating mutations of PTHR-1, and is characterized

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by prenatal lethality, premature and abnormal bone mineralization and ossification, and shortened limbs. The proliferative zone in the mutant growth plate is virtually absent and endochondral bone formation is markedly advanced simulating the phenotype of the PTHR-1 knockout mice. JMC is a rare autosomal dominant disorder caused by an activating mutation of the PTHR-1 leading to ligand-independent (constitutive) activation of the receptor. It is characterized by short-limbed dwarfism secondary to severe abnormalities of the growth plate and by hypercalcemia [73–75]. Although laboratory findings of JMC are reminiscent of primary hyperparathyroidism, parathyroid glands appear normal and circulating PTH levels are normal or undetectable. A heterozygous missense mutation of PTHR-1, which causes constitutive activation of the receptor, has also been identified in enchondromatosis, characterized by multiple benign cartilage tumors [76].

B. PTHrP and Osteoblast Biology 1. PTHrP Haploinsufficiency Leads to Osteopenia

PTHrP and PTH-1 receptors are expressed in cells of the osteogenic lineage [77–83]. Although PTH is essential for normal trabecular bone formation in the fetus [84] and therefore plays an anabolic role in utero, accumulating evidence indicates that PTH subserves a role postnatally primarily in defending against a decline in blood calcium, whereas PTHrP assumes a role in promoting bone formation in place of PTH as the animal matures (Fig. 5). Thus, heterozygous PTHrP-null mice, while phenotypically normal at birth, by 3 months of age exhibit a form of osteopenia characterized by a marked decrease in trabecular thickness and connectivity [85]. Moreover, their bone marrow contains an abnormally high number of adipocytes. Since the same pluripotent stromal cells in the bone marrow compartment can give rise to adipocytes and osteoprogenitor cells [86], the increased number of adipocytes and osteopenia in these mice could be attributed to altered stem cell differentiation as a consequence of PTHrP haploinsufficiency. Furthermore increased apoptotic osteoblastic cells are also observed. Taken together, these findings, and a variety of in vitro studies, suggest that PTHrP is important for the orderly commitment of pluripotential bone marrow stromal cells toward the osteogenic lineage and for their subsequent maturation and survival. In these animals, a physiological concentration of PTH is unable to compensate for PTHrP haploinsufficiency. One possible explanation is that a defect in skeletal progenitor cells was caused by PTHrP deficiency in utero and was

Figure 5 Effects of PTHrP and of PTH on the cartilaginous growth plate and on trabecular bone in the fetus and postnatally. In the fetus and postnatally, PTHrP acts as a paracrine factor to increase proliferation and reduce differentiation in the growth plate. Postnatally, PTHrP as a paracrine factor is anabolic for trabecular bone. PTH, as a hormone, is anabolic for trabecular bone in the fetus. As its main role post natally, PTH functions to preserve calcium homeostasis by producing net bone resorption. Circles depict chondrocytes and black areas denote trabecular bone.

only manifest in the postnatal state. Alternatively, domains of PTHrP not shared with the PTH molecule may subserve unique functions that are important for normal adult skeletal development. Further evidence for the anabolic boneforming effect of PTHrP came from crosses of mice deficient in PTH due to targeted deletion of the PTH gene, with PTHrP heterozygous mice lacking one allele encoding PTHrP [87]. PTH knockout mice have increased trabecular bone volume with diminished bone turnover consistent with the primary role of PTH in resorbing bone in order to maintain calcium homeostasis. Nevertheless, introducing PTHrP haploinsufficiency in these mice reduced trabecular bone of the PTH knockout mice to below wild-type levels by decreasing osteoprogenitor cell recruitment and enhancing osteoblast apoptosis and thereby diminishing bone formation. Consequently PTHrP plays an important bone-forming role in the post-natal state. More recent studies employing a cell-specific knockout model, in which osteoblastic PTHrP was specifically deleted, have demonstrated that osteoblastic PTHrP per se plays a critical role in enhancing osteoblastic bone formation. 2. PTHrP and the Bone Remodeling Process

In common with PTH, PTHrP in the postnatal skeleton appears to regulate both bone formation and bone resorption and both actions occur after binding to cells of the osteoblast phenotype [88]. Anabolic effects appear to predominate when exposure of skeletal cells to PTHrP is

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intermittent, whereas continuous exposure appears to elicit a skeletal catabolic effect [89]. The underlying mechanism for this pattern is unclear at present. The anabolic effects of the NH2-terminal region of both PTHrP and PTH occur via their action on the PTH-1 receptor, however, at least some of the effects of PTHrP and PTH on bone formation may be partly mediated through cytokines or growth factors such as IGF-1 [90]. The effects of PTHrP (and PTH) on osteoclastic bone resorption also appear to be mediated by interaction with PTH-1 receptors. Multinucleated osteoclasts are derived by differentiation of mononuclear precursors presumably of the monocyte/macrophage lineage. The interaction of PTHrP with its receptor on the osteoblast appears to increase soluble or cell-bound cytokines, most important of which is the receptor activator of nuclear factor kappa beta ligand (RANKL) (Fig. 6). Simultaneously, PTHrP (and PTH) can reduce concentrations of the soluble RANKL antagonist, osteoprotegerin (OPG). The dual effect thereby facilitates the capacity of RANKL to bind to its cognate receptor on osteoclastic precursors and multinucleated mature osteoclasts to enhance bone resorption. Interestingly, reduction in osteoblastic PTHrP in conditional knockout experiments not only reduced bone formation but even more dramatically reduced bone resorption, thus illustrating a role for osteoblastic PTHrP as a “coupling” factor. Several important steroid modulators may influence the action of PTHrP and of PTH on bone anabolism and catabolism. Thus, glucocorticoids may increase PTH-1 mRNA receptor expression [90], PTH binding, and PTH-stimulated camp production, effects which could lead to a net increase in PTHrP-mediated bone resorption. Estrogen, on the

other hand, may inhibit PTHrP-and PTH-stimulated adenylate cyclase stimulation in osteoblasts [91, 92], and estrogen deficiency in postmenopausal women leads to increased skeletal catabolic effects of continuous PTH administration [93]. It is important to note that the capacity either of circulating PTH or of circulating or locally released PTHrP to exert anabolic or catabolic effects will, most likely, be determined not only by ambient levels of these two peptides but also by the state of occupancy of the receptors by either of these ligands, which may in turn alter receptor internalization and thus the efficacy of the peptides. 3. Potential Therapeutic Role in Osteoporosis

The anabolic effects of intermittent PTH administration on bone and its therapeutic potential in osteoporosis have been extensively explored [94]. With the recognition that PTHrP is the endogenous ligand for the PTH/PTHrP receptor in osteoblasts, its use as an anabolic agent has also been investigated. Indeed the anabolic action of intermittently administered PTHrP(1-36) has now been documented in women with post menopausal osteoporosis [95]. Further studies on the mechanism of PTHrP and PTH as skeletal anabolic agents are clearly needed in order to produce additional agents, which may have improved efficacy profiles and routes of administration.

V. SUMMARY The discovery of PTHrP has led to improved understanding of the pathogenesis of hypercalcemia of malignancy

Figure 6 Effect of PTHrP on osteoclastic bone resorption. PTHrP interacts with osteoblastic stromal cells to increase expression of receptor activator of nuclear factor kappa b ligand (RANKL) and to decrease expression of the soluble decoy receptor osteoprotegerin (OPG). RANKL can activate its receptor RANK which can then increase the production of active osteoclasts from hematopoietic stem cells, resulting in increased bone resorption.

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and may also be relevant for understanding malignancyinduced bone resorption even in the absence of hypercalcemia. Unexpectedly, and of equal importance, this humoral factor has been found to play a role in a number of normal developmental processes and in the differentiated function of a variety of normal tissues. Most notable is its crucial role in the development of the cartilaginous growth plate as demonstrated by gene disruption technology. Analyses of knockout mice have also emphasized the function of PTHrP in normal osteoblastic bone formation. Powerful new in vitro and in vivo molecular technologies are also providing insights into detailed intracellular signaling mechanisms of PTHrP and its interaction with other regulatory factors. Continued advances in this field should provide important new understanding of the pathophysiology of osteochondrodysplasias, osteopetrosis, hyperparathyroidism, and osteoporosis and should facilitate the development of new therapeutic tools for these disorders.

Acknowledgments This work on PTHrP was supported by the Canadian Institutes of Health Research and by the National Cancer Institute of Canada.

References 1. Gutman, A. B., Tyson, T. L., and Gutman E. B. (1936). Inorganic phosphorus and phosphatase activity in hyperparathyroidism, Paget’s disease, multiple myeloma and neoplastic disease of the bones. Arch. Intern. Med. 57, 379–413. 2. Albright, F. (1941). Case records of the Massachusetts General Hospital: Case 27461. N. Engl. J. Med. 225, 789–791. 3. Goltzman, D., Stewart, A. F., and Broadus, A. E. (1981). Malignancyassociated hypercalcemia: Evaluation with a sytochemical bioassay for parathyroid hormone. J. Clin. Endocrinol. Metab. 53, 899–904. 4. Moseley, J. M., Kubota, M., Diefenbach-Jagger, H., Wettenhall, R. E. H., Kemp, B. E., Suva, L. J., Rodda, C. P., Ebeling, P. R., Hudson, P. J., Zajac, J. D., and Martin, T. J. (1987). Parathyroid hormone-related protein purified from a human lung cancer cell line. Proc. Natl Acad. Sci. USA. 84, 5048–5052. 5. Stewart, A. F., Wu, T., Goumas, D., Burtis, W. J., and Broadus, A. E. (1987). N-terminal amino acid sequence of two novel tumor-derived adenylate cyclase-stimulating proteins: Identification of parathyroid hormone-like and parathyroid hormone-unlike domains. Biochem. Biophys. Res. Commun. 146, 672–678. 6. Strewler, G. J., Ster, P. H., Jacobs, J. W., Eveloff, L., Klein, R. F., Leung, S. C., Rosenblatt, M., and Nissenson, R. A. (1987). Parathyroid hormone-like protein from human renal carcinoma cells. Structural and functional homology with parathyroid hormone. J. Clin. Invest. 80, 1803–1807. 7. Juppner, H., Abou-Samra, A-B., Freeman, M., Kong, X.-F., Schipani, E., Richards, J., Kolakowski, L. F., Jr., Hock, J., Potts, J. T., Jr., Kronenberg, H. M., and Segre, G. V. (1991). A G protein-linked receptor for parathyroid hormone and parathyroid hormone-related peptide. Science 254, 1024–1026.

8. Abou-Samra, A.-B., Juppner, H., Force, T., Freeman, M. W., Kong, J. T., Jr., Kronenberg, H. M., and Segre, G. V. (1992). Expression cloning of a common receptor for parathyroid hormone and parathyroid hormone-related peptide from rat osteoblast-like cells: A single receptor stimulates intracellular accumulation of both camp and inositol trisphosphates and increases intracellular free calcium. Proc. Natl, Acad. Sci. USA. 89, 2732–2736. 9. Guise, T. A., Yin, J. J., Taylor, S. D., Kumagai, Y., Dallas, M., Boyce, B. F., Yoneda, T., and Mundy, G. R. (1996). Evidence for a causal role of parathyroid hormone-related peptide protein in the pathogenesis of human breast cancer-mediated osteolysis. J. Clin. Invest. 98, 1544–1549. 10. Rabbani, S. A., Gladu, J., Harakidas, P., Jamison, B., and Goltzman, D. (1999). Over-production of parathyroid hormone-related peptide results in increased osteolytic skeletal metastasis by prostate cancer cells in vivo. Int. J. Cancer. 80, 257–264. 11. Mangin, M., Webb, A. C., Dreyer, B, E., Posillico, J, T., Ikeda, K., Weir, E. C., Stewart, A. F., Bander, N. H., Milstone, L., Barton, D. E., Francke, U., and Broadus, A. E. (1988). Identification of a cDNA encoding a parathyroid hormone-like peptide from a human tumor associated with hypercalcemia of malignancy. Proc. Natl Acad. Sci. USA. 85, 597. 12. Hendy, G., Sakaguchi, A. Y., Yasuda, T., Weber, D. K., Wang, L.-M., Yoshida, M. C., Banville, D., and Goltzman, D. (1988). Gene for parathyroid hormone-like peptide is on rat chromosome 2. Biochem. Biophys. Res. Commun. 157, 558–562. 13. Mangin, M., Ikeda, K., Dreyer, B. E., and Broadus, A. E. (1989). Isolation and characterization of the human parathyroid hormone-like peptide gene. Proc. Natl Acad. Sci. USA. 86, 2408–2412. 14. Yasuda, T., Banville, D., Hendy, G. N., and Goltzman, D. (1989). Characterization of the human parathyroid hormone-like peptide gene. J. Biol. Chem. 264, 7720–7725. 15. Mangin, M., Ikeda, K., Dreyer, B. E., and Broadus. A. E. (1990). Identification of an up-stream promoter of the human parathyroid hormone-related peptide gene. Mol. Endocrinology 4, 851–858. 16. Vasavada, R. C., Wysolmerski, J. J., Broadus, A. E., and Philbrick, W. M. (1993). Identification and characterization of a GC-rich promoter of the human parathyroid hormone-related peptide gene. Mol. Endocrinology 7, 273–283. 17. Karaplis, A. C., Yasuda, T., Hendy, G. N., Goltzman, D., and Banville, D. (1990). Gene encoding parathyroid hormone-like peptide: Nucleotide sequences of the rat gene and comparison with the human homologue. Mol. Endocrinol. 4, 441–446. 18. Mangin, M., Ikeda, K., and Broadus, A. E. (1990). Structure of the mouse gene encoding parathyroid hormone-related peptide. Gene 95, 195–202. 19. Thiede, M., and Rutledge, S. J. (1990). Nucleotide sequences of a parathyroid hormone-related peptide expressed by the 10 day chicken embryo. Nucleic Acids Res. 18, 3062. 20. Rosol, T. J., Steinmeyer, C. L., McCauley, L. K., Grone, A., DeWille, J. W., and Capen, C. C. (1995). Sequence of the cDNAs encoding canine parathyroid hormone-related protein and parathyroid hormone. Gene 160, 241–243. 21. Kremer, R., Karaplis, A. C., Henderson, J., Gulliver, W., Banville, D., Hendy, H. N., and Goltzman, D. (1991). Regulation of parathyroid-like peptide in cultured normal human keratinocytes: Effect of growth factors and 1,25 dihydroxyvitamin D3 on gene expression and secretion. J. Clin. Invest. 87, 884–893. 22. Sebag, M., Henderson, J., Goltzman, D., and Kremer, R. (1994). Regulation of parathyroid hormone-related peptide production in normal human mammary epithelial cells in vitro. Am. J. Physiol. 267(3 Pt 1), C723–730. 23. Akilu, F., Park, M., Goltzman, D., and Rabbani, S. A. (1997). Induction of parathyroid hormone-related peptide by the Ras oncogene: Role of Ras farnesylation inhibitors as potential therapeutic agents for hypercalcemia of malignancy. Cancer Res. 57, 4515–4522.

Chapter 17 Interaction of Parathyroid Hormone-related Peptide with the Skeleton 24. Haq, M., Kremer, R., Goltzman, D., and Rabbani, S. A. (1993). A vitamin D analogue (EB1089) inhibits parathyroid hormone related protein production and prevents the development of malignancy-associated hypercalcemia in vivo. I. Clin. Invest. 911, 2416–2422. 25. Pizzi, H., Gladu, J., Carpio, L., Miao, D., Goltzman, D., and Rabbani S. A. (2003). Androgen regulation of parathyroid hormone-related peptide production in human prostate cancer cells. Endocrinology 144, 858–867. 26. Rabbani, S. A., Khalili, P., Arakelian, A., Pizzi, H., Chen, G., and Goltzman, D. (2005). Regulation of parathyroid hormone related peptide by estradiol (E2): effect on tumor growth and metastasis in vitro and in vivo. Endocrinology 146, 2885–2894. 27. Thiede, M. A. (1989). The mRNA encoding a parathyroid hormonelike peptide is produced in mammary tissue in response to elevations in serum prolactin. Mol. Endocrinology 3, 1443–1447. 28. Thiede, M. A., Daifotis, A. G., Weir, E. C., Brines, M. L., Burtis, W. J., Ikeda, K., Dreyer, B. E., Garfield, R. E., and Broadus, A. E. (1990). Intrauterine occupancy controls expression of the parathyroid hormone-related peptide gene in preterm rat myometrium. Proc. Natl Acad. Sci. USA. 87, 6969–6973. 29. Liu, B., Goltzman, D., and Rabbani, S. A. (1995). Processing of parathyroid hormone-related peptide (pro-PTHrP) by the prohormone convertase furin: Effect on biological activity. Am. J. Physiol. 268, E832-E838. 30. Soifer, N. E., Dee, K. E., Insogna, K. L., Burtis, W. J., Matovcik, L. M., Wu, T. L., Milstone, L. M., Broadus, A. E., Philbrick, W. M., and Stewart, A. F. (1992). Secretion of a novel mid-region fragment of parathyroid hormone-related protein by three different cell lines in culture. J. Biol. Chem. 267, 18236–18243. 31. Yano, S., Macleod, R. J., Chattopadhyay, N., Tfelt-Hensen, J., Kifor, O., Butters, R. R., and Brown, E. M. (2004). Calcium-sensing receptor activation stimulates parathyroid hormone-related protein secretion in prostate cancer cells: role of epidermal growth factor receptor transactivation. Bone 35(3), 664–672. 32. VanHouten, J., Dann, P., McGeoch, G., Brown, E. M., Krapcho, K., Neville, M., and Wysolmerski, J. J. (2004). The calcium-sensing receptor regulates mammary gland parathyroid hormone-related protein production and calcium transport. J. Clin. Invest 113, 598–608. 33. Chan, G. K., Deckelbaum, R. A., Bolivar, I., Goltzman, D., and Karaplis, A. C. (2001). PTHrP inhibits adipocyte differentiation by down-regulating PPAR gamma activity via a MAPK-dependent pathway. Endocrinology 142, 4900–4909. 34. Miao, D., Tong, X. K., Chan, G. K., Panda, D., McPherson, P. S., and Goltzman, D. (2001). Parathyroid hormone-related peptide stimulates osteogenic cell proliferation through protein kinase C activation of the Ras/mitogen-activated protein kinase signaling pathway. J. Biol Chem. 276, 32204–32213. 35. Mahon, M. J., Donowitz, M., Yun, C. C., and Segre, G. V. (2002). Na+ H+-exchange regulatory factor 2 directs parathyroid hormone receptor signalling. Nature 417, 858–861. 36. Orloff, J. J., Ganz, M. B., Ribaudo, A. E., Buris, W. J., Reiss, M., Milstone, L. M., and Stewart, A. F. (1992). Analysis of PTHrP binding and signal transduction mechanisms in benign and malignant squamous cells. Am. J. Physiol. 262, E599–E607. 37. Orloff, J. J., Kats, Y., Urena, P., Schipani, E., Vasavada, R. C., Philbrick, W. M., Behal, A., Abou-Samra, A.-B., Segre, G. V., and Juppner, H. (1995). Further evidence for a novel receptor for aminoterminal parathyroid hormone-related protein on keratinocytes and squamous carcinoma cell lines. Endocrinology 136, 3016–3023. 38. Care, A. E., Abbas, S. K., Pickard, D. W., Barri, M., Drinkhill, M., Findlay, J. B. C., White, I. R., and Caple, I. W. (1990). Stimulation of ovine placental transport of calcium and magnesium by mid-molecule fragments of human PTHrP. Exp. Physiol. 75, 605–608. 39. Kovacs, C. S., Lanske, B., Hunzelman, J. L., Guo, J., Karaplis, A. C., and Kronenberg, H. M. (1996). Parathyroid hormone-related peptide (PTHrP) regulated fetal-placental calcium transport through a receptor

40.

41.

42.

43.

44.

45.

46.

47.

48.

49.

50.

51.

52.

53.

54.

303

distinct from the PTH/PTHrP receptor. Proc. Natl Acad. Sci. USA. 93, 15233–15238. Kaiser, S. M., Laneuville, P., Bernier, S. M., Rhim, J. S., Kremer, R., and Goltzman, D. (1992). Enhanced growth of a human keratinocytes cell line induced by antisense RNA for parathyroid hormone-related peptide. J. Biol. Chem. 267, 13623–13628. Henderson, J. E., Amizurka, N., Warshawsky, H., Biasotto, D., Lanske, B. M. K., Goltzman, D., and Karaplis, A. C. (1995). Nucleolar localization of parathyroid hormone-related peptide enhances survival of chondrocytes under conditions that promote apoptotic cell death. Mol. Cell. Biol. 15, 4064–4075. Jans, D. A., Thomas, R. J., and Gillespie, M. T. (2003). Parathyroid hormone-related protein (PTHrP): a nucleocytoplasmic shuttling protein with distinct paracrine and intracrine roles. Vitam. Horm. 66, 345–384. Fenton, A. J., Kemp, B. E., Kent, G. N., Moseley, J. M., Zheng, M. H., Rowe, D. J., Britto, J. M., Martin, T. J., and Nicholson, G. C. (1991). A carboxyl-terminal peptide from the parathyroid hormone-related protein inhibits bone resorption by osteoclasts. Endocrinology 129, 1762–1768. Fenton, A. J., Kemp, B. E., Hammonds, J., Mitchelhill, K., Moseley, J. M., Martin, T. J., and Nicholson, G. C. (1991). A potent inhibitor of osteoclastic bone resorption within a highly conserved pentapeptide region of parathyroid hormone-related protein, PTHrP [107–111]. Endocrinology 129, 3424–3426. Fukayama, S., Tashjian, Jr, A., David, J. N., and Chisholm, J. C. (1995). Signaling by N-and C-terminal sequences of parathyroid hormone-related protein in hippocampal neurons. Proc. Natl Acad. Sci. USA 92, 10182–10186. Henderson, J., Shustik, C., Kremer, R., Rabbani, S. A., Hendy, G. N., and Goltzman, D. (1990). Circulating concentrations of parathyroid hormone-like peptide in malignancy and in hyperparathyroidism. J. Bone Miner. Res. 5, 105–113. Kaiser, S. M., Sebag, M., Rhim, J. S., Kremer, R., and Goltzman, D. (1994). Antisense-mediated inhibition of parathyroid hormone-related peptide production in a keratinocytes cell line impedes differentiation. Mol. Endocrinol. 8, 139–147. Wysolmerski, J. J., McCaughern-Carucci, J. F., Daifotis, A. G., Broadus, A. E., and Philbrick, W. M. (1995). Overexpression of parathyroid hormone-related protein or parathyroid hormone in transgenic mice impairs branching morphogenesis during mammary gland development. Development 12, 3539–3547. Wysolmerski, J. J., Philbrick, W. M., Dunbar, M. E., Lanske, B., Kronenberg, H., Karaplis, A., and Broadus, A. E. (1998). Rescue of the parathyroid hormone-related protein knockout mouse demonstrates that parathyroid hormone-related protein is essential for mammary gland development. Development 125, 1285–1294. Lam, M. H., Olsen, S. L., Rankin, W. A., Ho, P. W., Martin, T. J., Gillespie, M. T., and Moseley, J. M. (1997). PTHrP and cell division: Expression and localization of PTHrP in a keratinocytes cell line (HaCaT) during the cell cycle. J. Cell. Physiol. 173, 433–446. Massfelder, T., Dann, P., Wu, T. L., Vasavada, R., Helwig, J. J., and Steward, A. F. (1997). Opposing mitogenic and anti-mitogenic action of parathyroid hormone-related protein in vascular smooth muscle cells: a critical role for nuclear targeting. Proc. Natl Acad. Sci. USA. 94, 13630–13635. Meerovitch, K., Wing, S., and Goltzman, D. (1998). Proparathyroid hormone-related protein is associated with the chaperone protein BiP and undergoes proteasome-mediated degradation. J. Biol. Chem. 273, 21025–21030. Aarts, M. M., Rix, A., Guo, J., Bringhurst, R., and Henderson, J. E. (1999). The nucleolar targeting signal (NTS) of parathyroid hormone related protein mediates endocytosis and nucleolar translocation. J. Bone Miner. Res. 14, 1493–1503. Watson, P. H., Fraher, L. J., Hendy, G. N., Chung, U. I., Kisiel, M., Natale, B. V., and Hodsman, A. B. (2000). Nuclear localization of the

304

55.

56.

57.

58.

59.

60.

61.

62.

63.

64.

65.

66.

67.

68.

69.

type 1 PTH/PTHrP receptor in rat tissues. J. Bone Miner Res. 15, 1033–1044. Nguyen, M., He, B., and Karaplis, A. (2001). Nuclear forms of parathyroid hormone-related peptide are translated from non-AUG start sites downstream from the initiator methionine. Endocrinology 142, 694–703. Okano, K., Pirola, C. J., Wang, H. M., Forrester, J. S., Fagin, J. A., and Clemens, T. L. (1995). Involvement of cell cycle and mitogen-activated pathways in induction of parathyroid hormone-related protein gene expression in rat aortic smooth muscle cells. Endocrinology 136, 1782–1789. Lam, M. H., Olsen, S. L., Rankin, W. A., Ho, P. W., Martin, T. J., Gillespie, M. T., and Moseley, J. M. (1997). PTHrP and cell division: expression and localization of PTHrP in a keratinocyte cell line (HaCaT) during the cell cycle. J. Cell. Physiol. 173, 433–446. Aarts, M. M., Levy, D., He, B., Stregger, S., Chem, T., Richards, S., and Henderson, J. E. (1999). Parathyroid hormone-related protein interacts with RNA. J. Biol. Chem. 274, 4832–4838. Karaplis, A. C., Luz, A., Glowacki, J., Bronson, R. T., Tybulewicz, V. L. J., Kronenberg, H. M., and Mulligan, R. C. (1994). Lethal skeletal dysplasia from targeted disruption of the parathyroid hormone-related peptide (PTHrP) gene. Genes Dev. 8, 277–289. Amizuka, N., Henderson, J. E., Warshawsky, H., Goltzman, D., and Karaplis, A. C. (1994). Parathyroid hormone-related peptide (PTHrP)-depleted mice show abnormal epiphyseal cartilage development and altered endochondral bone formation. J. Cell Biol. 126, 1611–1623. Lee, K., Lanske, B., Karaplis, A. C., Deeds, J. D., Kohno, H., Nissenson, R. A., Kronenberg, H. M., and Segre, G. V. (196). Parathyroid hormone-related peptide delays terminal differentiation of chondrocytes during endochondral bone development. Endocrinology 137, 5055–5067. Weir, E. C., Philbrick, W. M., Amling, M., Neff, L. A., Baron, R., and Broadus, A. E. (1996). Targeted overexpression of parathyroid hormone-related peptide in chondrocytes causes chondrodysplasia and delayed endochondral bone formation. Proc. Natl Acad. Sci. USA. 93, 10240–10245. Amizuka, N., Henderson, J. E., Hoshi, K., Warshawsky, H., Ozawa, H., Goltzman, D., and Karaplis, A. C. (1996). Programmed cell death of chondrocytes and aberrant chondrogenesis in mice homozygous for parathyroid hormone-related peptide gene deletion. Endocrinology 137, 5055–5067. Farnum, C. E., and Wilsman, N. J. (1989). Condensation of hypertrophic chondrocytes at the chondro-osseous junction of growth plate cartilage in Yucatan swine: relationship to long bone growth. Am. Anat. 186, 346–358. Amling, M., Neff, L., Tanaka, S., Inoue, D., Kuida, K., Weir, E. C., Philbrick, W. M., Broadus, A. E., and Baron, R. (1997). Bcl-2 downstream of parathyroid hormone-related peptide in a signal pathway that regulates chondrocytes maturation during skeletal development. J. Cell. Biol. 136, 205–213. Provot, S., and Schipani, E. (2005). Molecular mechanisms of endochondral bone development. Biochem. Biophys. Res. Commun. 328, 658–665. Lanske, B., Karaplis, A. C., Lee, K., Luz, A., Vortkamp, A., Pirro, A., Karperien, M., Defize, L. H., Ho, C., Mulligan, R. C., Abou-Samra, A. -B., Juppner, H., Segre, G. V., and Kronenberg, H. M. (1996). PTH/PTHrP receptor in early development and Indian Hedgehog-regulated bone growth. Science 273, 663–666. Decklebaum, R. A., Chan, G., Miao, D., Goltzman, D., and Karaplis, A. C. (2002). Ihh enhances differentiation of CfK-2 chondrocytic cells and antagonizes PTHrP-mediated activation of PKA. J. Cell Sci. 115 (Pt 14), 3015–3025. Davidson, D., Blanc, A., Filion, D., Wang, H., Plut, P., Pfeffer, G., Buschmann, M. D., and Henderson, J. E. (2005). FGF18 signals

DAVID GOLTZMAN

70.

71.

72.

73. 74.

75.

76.

77.

78.

79.

80.

81.

82.

83.

84.

85.

86.

through FGFR3 to promote chondrogenesis. J. Biol. Chem. 27, 20509–20515. Schipani, E., Lanske, B., Hunzelman, J., Luz, A., Kovacs, C. S., Le, K., Pirro, A., Kronenberg, H. M., and Juppner, H. (1997). Targeted expression of constitutively active receptors for parathyroid hormone and parathyroid hormone-related peptide delays endochondral bone formation and rescues mice that lack parathyroid hormone-related peptide. Proc. Natl Acad. Sci. USA. 94, 13689–13694. Zhang, P., Jobert, A. S., Couvineau, A., and Sillve, C. (1998). A homozygous inactivating mutation in the parathyroid hormone/ parathyroid hormone-related peptide receptor causing Blomstrand chondrodysplasia. J. Clin. Endocrinol. 83, 3365–3368. Karaplis, A. C., He, B., Nguyen, M. T., Young, I. D., Semeraro, D., Ozawa, H., and Amizuka, N. (1998). Inactivating mutation in the human parathyroid hormone receptor type 1 gene in Blomstrand chondrodysplasia. Endocrinology 139, 5255–5258. Kruse, K. (1993). Calcium metabolism in the Jansen type of metaphyseal dysplasia. Eur. J. Pediatr. 1522, 912–915. Schipani, E., Kruse, K., and Juppner, H. (1995). A constitutively active mutant PTH-PTHrP receptor in Jansen-type metaphyseal chondrodysplasia. Science 268, 98–100. Schipani, E., Langman, C. B., Parfitt, A. M., Jensen, G. S., Kikuch, S., Kooh, S. W., Cole, W. G., and Juppner, H. (1996). Constitutive activated receptors for parathyroid hormone-related peptide in Jansen metaphyseal chondrodysplasia. N. Engl. J. Med. 335, 708–714. Hopyan, S., Gokgoz, N., Poon, R., Gensure, R. C., Yu, C., Cole, W. G., Bell, R. S., Juppner, H., Andrulis, I. L., Wunder, J. S., and Alman, B. A. (2002). A mutant PTH/PTHrP type 1 receptor in enchondromatosis. Nat. Genet. 30, 306–310. Lee, K., Deeds, J. D., Bond, A. T., Juppner, H., About-Samra, A.-B., and Segre, G. V. (1993). In situ localization of PTH/PTHrP receptor mRNA in the bone of fetal and young rats. Bone 14, 341–345. Lee, K., Deeds, J. D., Chiba, S., Un-No, M., Bond, A. T., and Segre, G. V. (1994). Parathyroid hormone induces sequential c-fos expression in bone cells in vivo: In situ localization of its receptor and c-fos messenger ribonucleric acids. Endocrinology 134, 441–450. Lee, K., Deeds, J. D., and Segre, G. V. (1995). Expression of parathyroid hormone related-peptide and its receptor messenger ribonucleic acids during fetal development of rats. Endocrinology 136, 453–463. Fermor, B., and Skerry, T. M. (1995). PTH/PTHrP receptor expression on osteoblasts and osteocytes but not resorbing bone surfaces in growing rats. J. Bone Miner. Res. 10, 1935–1943. Celic, S., Chilco, P. J., Zajac, J. D., Martin, T. J., and Findley, D. M. (1996). A type 1 collagen substrate increases PTH/PTHrP receptor mRNA expression and suppresses PTHrP mRNA expression in UMR106-06 osteoblast-like cells. J. Endocrinol. 150, 299–308. Suda, N., Gillespie, M. T., Traianedes, K., Zhou, H., Ho, P. W. M., Hards, D. K., Allan, E. H., Martin, T. J., and Moseley, J. M. (1996). Expression of parathyroid hormone-related protein in cells of osteoblast lineage. J. Cell. Physiol. 166, 94–104. McCauley, L. K., Koh, A. J., Beecher, C. A., Cui, Y., Rosol, T. l. J., and Franceschi, R. T. (1996). PTh/PthrP receptor is temporally regulated during osteoblast differentiation and is associated with collagen synthesis. J. Cell. Biochem. 61, 638–647. Miao, D., He, B., Karaplis, A. C., Goltzman, D. (2002). Parathyroid hormone is essential for normal fetal bone formation. J. Clin. Invest. 109, 1173–1182. Amizuka, N., Karaplis, A. C., Henderson, J. E., Warshawsky, H., Lipman, M. L., Matsuki, Y., Eijiri, S., Tanaka, M., Izumi, N., Ozawa, H., and Goltzman, D. (1996). Haploinsufficiency of parathyroid hormonerelated peptide (PTHrP) results in abnormal postnatal bone development. Dev. Biol. 175, 166–176. Owen, M., and Friedenstein, A. J. (1988). Stromal stem cells: marrowdeprived osteogenic precursors. Ciba Foundation Symposium 136, 42–60.

Chapter 17 Interaction of Parathyroid Hormone-related Peptide with the Skeleton 87. Miao, D., Li, J., Xue, Y., Su, H., Karaplis, A. C., and Goltzman, D. (2004). Parathyroid hormone-related peptide is required for increased trabecular bone volume in parathyroid hormone-null mice. Endocrinology 145, 3554–3562. 88. Rouleau, M. F., Mitchell, J., and Goltzman, D. (1990). Characteristics of the major parathyroid hormone target cell in the endosteal metaphysis of rat long bones. J. Bone Miner. Res. 5, 1043–1053. 89. Canalis, E., McCarthy, T. L., and Centrella, M. (1990). Differential effects of continuous and transient treatment with parathyroid hormone related peptide (PTHrP) on bone collagen synthesis. Endocrinology 126, 1806–1812. 90. Urena, P., Iida-Klein, A., Kong, X.-F., Juppner, H., Kronenberg, H. M., About-Samra, A.-B, and Segre, G. V. (1994). Regulation of parathyroid (PTH)/PTH-related peptide receptor messenger ribonucleic acid by glucocorticoids and PTH in ROS 17/2.8 and OK Cells. Endocrinology 134, 451–456. 91. Ernst, M., Heath, J. K., and Rodan, G. A. (1989). Estradiol effects on proliferation, messenger ribonucleric acid for collagen and insulin-like

92.

93.

94. 95.

305

growth factor-I, and parathyroid hormone-stimulated adenylate cyclase activity in osteoblastic cells from calvariae and long bones. Endocrinology 125, 825–833. Fukayama, S., and Tashjian, A. H. (1989). Direct modulation by estradiol of the response of human bone cells (SaOS-2) to human parathyroid hormone (PTH) and PTH-related protein. Endocrinology 124, 397–401. Joborn, C., Ljunghall, S., Larsson, K., Lindh, E., Naessen, T., Wide, L., Akerstrom, G., and Rastad, J. (1991). Skeletal responsiveness to parathyroid hormone in healthy females: Relationship to menopause and estrogen replacement. Clin. Endocrinol. 34, 335–339. Reeve, J. (1996). PTH: A future role in the management of osteoporosis? J. Bone Miner. Res. 11, 440–445. Horowitz, M. J., Tedesco, M. B., Gundberg, C., Garcia-Ocana, A, Stewart, A. F. (2003). Short-term, high-dose parathyroid hormonerelated protein as a skeletal anabolic agent for the treatment of postmenopausal osteoporosis. J. Clin. Endocrinol. Metab. 88, 569–575.

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Chapter 18

The Vitamin D Hormone and its Nuclear Receptor: Mechanisms Involved in Bone Biology Geert Carmeliet, Annemieke Verstuyf, Christa Maes, Guy Eelen, and Roger Bouillon

V. Pathology and Therapy Related to Vitamin D Availability, Metabolism, and Function VI. Conclusions References

I. Introduction II. Metabolism of Vitamin D III. Nuclear Vitamin D Receptor IV. Vitamin D and Bone Cells

I. INTRODUCTION

retinoid X receptor and associating specifically with vitamin D responsive elements (VDREs) in target genes. Upon binding of 1,25(OH)2D3, VDR releases corepressors and attracts general transcription factors and coactivators, some of them acetylating histones to create a permissive chromatin surrounding. At the cellular level, 1,25(OH)2D3 treatment inhibits osteoblast proliferation but induces osteoblast differentiation when given at the differentiation stage and protects osteoblastic cells from undergoing apoptosis. In addition, 1,25(OH)2D3 promotes osteoclast differentiation and alters chondrocyte development.

The active metabolite of vitamin D, 1α,25 dihydroxyvitamin D3 [1,25(OH)2D3], has multiple functions in humans and animals but its major role is related to bone metabolism and mineral homeostasis. Vitamin D is metabolized to its active form by two sequential hydroxylations on carbon-25 and carbon-1. The genomic action of 1,25(OH)2D3 is mediated by the nuclear vitamin D receptor (VDR) that binds its ligand with high affinity. VDR is a phosphoprotein that regulates gene expression by heterodimerizing with Dynamics of Bone and Cartilage Metabolism

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Vitamin D deficiency or mutations in the 25-hydroxyvitamin D-1α-hydroxylase gene or VDR gene lead to rickets and osteomalacia, a phenotype that is also observed in the corresponding knock-out mouse models. Although bone mineralization can be maintained in the absence of VDR or its ligand when sufficient mineral is supplied, the precise role of the vitamin D system in fine tuning of bone mineralization and homeostasis remains to be determined.

II. METABOLISM OF VITAMIN D A. Source and Synthesis of Vitamin D The secosteroid vitamin D3 can be produced endogenously in the skin by the action of UV light (290–300 nm), converting 7-dehydrocholesterol into previtamin D3 that is in thermoequilibrium with vitamin D3 (Fig. 1). A second source of vitamin D is the dietary intake, whereby 50% of vitamin D is absorbed by the enterocytes and transported via the chylomicrons to the general circulation. A first step in the activation pathway of vitamin D3 is the hydroxylation at carbon 25 catalyzed by a cytochrome P450 (CYP) enzyme and takes place primarily in the liver to produce 25-hydroxyvitamin D3 [25(OH)D3], the major circulating form of vitamin D3. Several CYPs, localized either in the inner mitochondrial membrane or in

Figure 1

the microsomes, can catalyze this reaction in vitro but CYP27A1 and CYP2R1 are the two viable candidates. Mutations in the human and mouse genes encoding the mitochondrial CYP27A1 protein impair bile acid synthesis, but have no consequences for vitamin D metabolism [1, 2]. On the other hand, molecular analysis of a patient with selective 25-hydroxyvitamin D deficiency and symptoms of vitamin D deficiency, including skeletal abnormalities, identified a mutation in the CYP2R1 gene (chromosome 11p15.2), encoding a microsomal enzyme [3]. Nevertheless, this patient still had measurable levels of 1,25(OH)2D3 and responded to vitamin D treatment. 25(OH)D3 is biologically inactive and requires further hydroxylation in the kidney to the active hormone 1,25(OH)2D3 by another mitochondrial cytochrome P450 enzyme, 25-hydroxyvitamin D-1α-hydroxylase (CYP27B1). The production of 1,25(OH)2D3 is regulated primarily at this final step by several factors including parathyroid hormone (PTH) (positive feedback), calcium, phosphate, and 1,25(OH)2D3 (negative feedback). The proximal renal tubule is the principal site of 1α-hydroxylase activity but expression has also been detected in several tissues including bone and cartilage. The CYP27B1 gene has been cloned in several species [4–8] and the human gene is mapped on chromosome 12q13.3. Mutations in the human gene are responsible for the rare autosomal recessive disease pseudovitamin D-deficiency rickets (PDDR) and will be discussed in Section V.A.2.

Sources, bioactivation of vitamin D3 and metabolism, actions of 1,25(OH)2D3. The active 1,25(OH)2D3 hormone is synthesized from the vitamin D precursor by two successive enzymatic hydroxylations on carbon-25 and carbon-1 in liver and kidney respectively. 1,25(OH)2D3 acts principally on intestine, kidney, and bone in order to keep serum calcium constant. The major pathway in catabolism is via 24-hydroxylase that has a broad tissue distribution.

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B. Transport of Vitamin D In blood, vitamin D and its active metabolites are transported by binding to a specific plasma protein vitamin Dbinding protein (DBP). The vitamin D-binding site of DBP is a cleft located at the surface of the molecule and partly in contact with the surrounding solvent as evidenced from the crystal structure of the human DBP [9]. Mice deficient in DBP had low levels of total vitamin D metabolites, but were otherwise normal, suggesting that intracellular concentrations of 1,25(OH)2D3 are still adequate [10]. However, when dietary vitamin D was reduced a clear functional 1,25(OH)2D3 deficiency state with osteomalacia developed rapidly in DBP-null mice, indicating that DBP provided a degree of protection against dietary-induced vitamin D deficiency. Unexpectedly, DBP did not create a buffer for 1,25(OH)2D3 as DBP-null mice were resistant to vitamin D toxicity, possibly reflecting a more rapid entry into metabolic breakdown pathways and a more rapid urinary excretion of 25(OH)D3. Indeed, after renal glomerular filtration of 25(OH)D3 in complex with DBP it is reabsorbed in the proximal tubular cells by the endocytic receptor megalin, rather than entering the cell by simple diffusion. The importance of this pathway was underlined by the phenotype of mice with a total or kidney-specific megalin gene defect [11, 12]. The latter mice show hypocalcemia and osteomalacia as a consequence of hypovitaminosis D, indicating that renal reuptake of the 25(OH)D3–DBP complex is essential to preserve vitamin D stores.

C. 24-Hydroxylase: Catabolism or Specific Function? An alternative hydroxylation of 25(OH)D3 occurs on carbon 24 by the multifunctional enzyme 24-hydroxylase (CYP24), mapped on human chromosome 20q13 [13]. This enzyme not only initiates the catabolic cascade of 25(OH)D3 and 1α,25(OH)2D3 [14, 15] by 24-hydroxylation but catalyzes also the dehydrogenation of the 24-OH group and performs 23-hydroxylation, resulting in 24-oxo1,23,25-(OH)3D3 [16]. This C24 oxidation pathway leads finally to calcitroic acid, which is the major end product of 1,25(OH)2D3 and this catabolic breakdown is regulated by 1,25(OH)2D3 itself. This indicates that CYP24 may play an important role in regulating the functions of 1,25(OH)2D3 in the target cells. In vivo evidence for the role of CYP24 in the catabolism of 1,25(OH)2D3 was provided by the generation of CYP24-null mice [17]. These mice were unable to clear 1,25(OH)2D3 from the bloodstream in response to 1,25(OH)2D3 treatment. Whether 24,25(OH)2D3 has a

physiological role in bone metabolism has previously been a matter of debate [18, 19]. CYP24-null mice, born from CYP24-null mothers, have impaired intramembranous bone formation that was however normalized after crossing them with vitamin D receptor (VDR)-null mice. This finding indicates that this phenotype was caused by increased 1,25(OH)2D3 levels acting via the VDR, and not by the absence of 24,25(OH)2D3.

III. NUCLEAR VITAMIN D RECEPTOR A. General Characteristics of the Protein and Gene The actions of 1,25(OH)2D3 are mediated by the nuclear vitamin D receptor (VDR), a member of the superfamily of steroid/thyroid hormone receptors [20]. The VDR has been cloned from several species, among which are humans, mouse, rat, chick, quail, Xenopus, zebrafish, rainbow trout, and Japanese flounder [21–28]. The recently cloned VDR from lamprey, an ancient vertebrate without calcified skeleton, indicates that the VDR predates the origin of a calcified skeleton [29]. VDR is a 55-kDa protein made up of 427 amino acids and is present at a low concentration (10–100 fmol/mg protein), consistent with the fact that it is a potent transcription regulatory molecule. The human VDR is encoded by a single gene mapped to chromosome 12q13. The gene is comprised of at least 14 exons; the noncoding 5′ end of the gene includes exons 1a, 1b, 1c, 1d, 1e, and 1f, whereas eight additional exons (exons 2–9) encode the structural portion of the VDR gene (Fig. 2) [30, 31]. For human VDR, a multitude of different transcripts were found that mostly vary in their 5′ untranslated region but encode the same 427-amino acid protein. Transcripts originating from the most distal promoter (near the 1f exon) were only found in calcitropic vitamin D target tissues (kidney, intestine, and parathyroid) [31] and illustrate that different receptor isoforms may contribute to tissuespecific effects.

B. Structure–Function Analysis A deeper insight into the structural and functional properties of the VDR protein and a better understanding of ligand-binding has been provided by site-directed mutation studies and by three-dimensional modeling of the protein by X-ray crystallography. From N- to C-terminus the VDR contains several functionally distinct domains which are listed below (Fig. 2).

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Figure 2

Gene and protein structure of the human vitamin D receptor (hVDR). The hVDR gene is composed of at least 14 exons and is mapped to chromosome 12q13. Indicated on the protein structure are the domains for DNA binding (DBD) and ligand binding (LBD) as well as the A/B domain and the hinge region. Two major phosphorylation sites are indicated by P.

1. The A/B-Domain

Unlike other nuclear receptors, the VDR contains a truncated A/B-domain which lacks the usual transactivation function (AF1). Therefore, little or no function was ascribed to this region. Evidence for a substantial role of the A/B-domain in transcriptional activity of the receptor came from recent studies on N-terminal variants of the VDR. Two of these variants with 50 or 23 amino acid extensions at their N-terminus display reduced transcriptional activity [32]. On the other hand, a polymorphic variant, called F/M4, lacking three amino acids at the N-terminal start, displays an increased transcriptional activity presumably due to a better interaction with the transcription factor TFIIB [33]. Multiple studies have linked the F/M4 polymorphism to increased bone mineral density [e.g. 34, 35]. 2. The C-Domain or DNA-Binding Domain

The DNA-binding domain (DBD) consists of two zincfinger modules within a 66 amino acid core sequence that is highly conserved among nuclear receptors [36]. The N-terminal module mediates specific DNA binding in the major groove of the DNA-binding site, whereas the C-terminal module is involved in binding a dimerization partner [37, 38]. In addition to this core sequence, the DBD contains a variable C-terminal extension (CTE) which is also involved in receptor dimerization and in directing the ligand-bound VDR, together with its dimerization partner to correctly spaced VDREs (see below) [39]. To function as a transcription factor, VDR needs to reside in the nucleus. In a screen for possible nuclear localization signals (NLS), two stretches of basic amino acids at both ends of the VDR region 79–105, which is partially located within the DBD, were shown to be crucial for nuclear accumulation [40]. A sequence of five basic amino acids positioned between the two zinc-finger modules is likely to function as an additional NLS [41].

3. THE D-DOMAIN OR HINGE-REGION

This region with poor sequence conservation confers flexibility to the receptor. An NLS of 20 amino acids in this region was found to facilitate nuclear transfer of the receptor [42]. 4. THE E-DOMAIN OR LIGAND-BINDING DOMAIN

The multifunctional ligand-binding domain (LBD) serves three major functions; binding of 1,25(OH)2D3, hetero- or homodimerization, and recruitment of regulatory factors needed for transcriptional activation. a. Ligand Binding-Function The VDR binds its ligand with extremely high affinity, in the range of 10−10 M. The ligand-binding pocket (LBP) comprises many segments spanning the entire C-terminal domain of 300 amino acids [43]. Great progress in understanding ligand binding and in identifying the crucial amino acids came from crystallization studies of the VDR in complex with 1,25(OH)2D3 [44]. Much like in other nuclear receptors the hydrophobic LBP of the VDR consists of 13 α-helices and a three-stranded β-sheet. In the LBP the A-ring of 1,25(OH)2D3 adopts a chair B conformation [45] and the connection between the A- and the C-ring lies in a hydrophobic channel formed by amino acids Ser275, Trp286, and Leu233. Contacts between specific amino acids lining the LBP and the hydroxyl groups of 1,25(OH)2D3 were found crucial for ligand binding. The 1α-OH group forms a hydrogen bond with Ser237 and with Arg274 whereas the 3-OH moiety is hydrogen-bonded with Ser278 and Tyr143. The 25-OH group connects through hydrogen bonds with His305 and His397. Studies with VDR constructs bearing mutations in the above-mentioned amino acids stress their importance in ligand binding [46]. b. Dimerization VDR forms homo- or heterodimers that can interact with specific DNA segments (VDREs, see below). The receptor

Chapter 18 The Vitamin D Hormone and its Nuclear Receptor

for 9-cis-retinoic acid, RXR, is the preferred dimerization partner for VDR [47–49] but also for other nuclear receptors and might therefore be a key element in cross-talk between nuclear receptor-mediated signaling [50, 51]. Part of the dimerization function of the receptor maps to two regions (amino acids 239–269 and 317–401) in the ligand-binding domain that are highly conserved across the steroid hormone receptor family [52]. In addition, regions in the DNA-binding domain are also involved in dimerization (see Section III.B.2). c. Transcriptional Activation Upon binding of 1,25(OH)2D3, VDR recruits RXR and this heterodimer can bind to specific DNA sequences in the promoter region of target genes called vitamin D responsive elements (VDREs) (Fig. 3). The consensus high-affinity VDRE consists of two hexameric half sites (PuG(G/T)TCA) arranged as a direct repeat spaced by three base pairs (DR3-type VDRE). Other natural VDREs exist, among which is an inverted palindrome of two hexameric binding sites known as IP9-type VDRE [53]. RXR occupies the 5′ half-site and VDR the 3′ half-site in cases of positive gene regulation. Reversal of the VDR-RXR polarity on the VDRE has been suggested to suppress gene expression (reviewed in reference [54]). Binding of the VDR–RXR dimer on the VDRE induces DNA-bending which facilitates transcription complex assembly [55].

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To contact the basal transcription complex, VDR–RXR releases corepressors and recruits coactivator molecules (Fig. 3). Upon ligand binding in the VDR-LBP, helix 12 closes off the LBP, much like a mousetrap, and exposes its activation function 2 (AF2) to which coactivators can bind through a conserved LXXLL motif in their amino acid sequence. The liganded VDR induces RXR to close off its LBP with helix 12 without binding of the cognate ligand 9-cis-retinoic acid. This “phantom ligand effect” allows RXR to recruit coactivators as well [56]. One type of attracted coactivators recruits histone acetyl-transferase (HAT) activity; they acetylate histone tails and create a permissive chromatin surrounding for gene transcription. This class contains CBP/p300 and the p160 family of proteins like SRC-1, GRIP1/TIF2, and ACTR (reviewed in reference [57]). Another type of coactivator is the multimeric vitamin D receptor interacting proteins (DRIP) complex, of which the 205-kDa subunit (DRIP205) interacts directly with the VDR–RXR heterodimer. The complex is not associated with HAT activity but instead recruits RNA polymerase II, the key enzyme needed for gene transcription [58]. Two other types of coactivators are NCoA-62/ski-interacting protein (SKIP) and WINAC. The former is thought to link transcriptional activation by nuclear receptors with mRNA splicing [59], whereas the latter interacts with VDR through the Williams’ syndrome

Figure 3 Simplified model of gene transcription by 1,25(OH)2D3. Ligand-bound VDR heterodimerizes with RXR and binds to VDRE sequence upstream of the target gene. Upon binding of 1,25(OH)2D3, VDR releases corepressors (not shown in figure) and attracts coactivator molecules (represented here by CBP/p300, p160, DRIP, and NCoA-62/SKIP) and general transcription factors (GTF, RNA polymerase II). Coactivators with histone acetyl transferase (HAT)-activity acetylate histones to create a permissive chromatin surrounding.

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transcription factor (WSTF) and displays ATP-dependent chromatin-remodeling activity [60]. VDR also directly contacts the N-terminal region of the general transcription factor IIB (TFIIB), a component of the basal transcription apparatus. The VDR–TFIIB interaction is not dependent upon 1,25(OH)2D3 and is disrupted by the ligand. A model is proposed in which unliganded VDR recruits TFIIB to the promoter region of vitamin D-responsive target genes where this transcription factor can be incorporated into the transcription apparatus after ligand-induced disruption from the VDR [61–63]. Recently TAFII-17, a subunit of the general transcription factor TFIID, was found to interact with VDR at low doses of 1,25(OH)2D3. Increased levels of TAFII-17 were linked to the bone-remodeling disorder Paget’s disease [64].

C. Modulation of Vitamin D Receptor 1. Phosphorylation, “Cross Talk”, and Receptor Activity

VDR is a phosphoprotein and the majority of the phosphorylation of VDR is on serine residues. Nevertheless the exact functional consequences of this phosphorylation have been difficult to determine. VDR becomes hyperphosphorylated upon 1,25(OH)2D3 treatment [65] which is partly catalyzed by casein kinase II that phosphorylates VDR at Ser-208 in the LBD and hereby amplifies VDR transcriptional activity [66]. Accordingly, a recent study indicated that 1,25(OH)2D3-dependent transcription was enhanced when phosphatase inhibitors were used. This enhancement may be due, at least in part, to an increased interaction between the VDR and the coactivator protein DRIP205 [67]. In addition, two 1,25(OH)2D3-independent phosphorylations of VDR antagonize this positive phosphorylation. Protein kinase C-mediated phosphorylation of Ser-51 decreased the DNA binding of the receptor [68]. Analogously, protein kinase A phosphorylates serines 182–185 in the LBD of VDR that seems to interfere with transactivation [69]. Taken together, VDR activity is both positively and negatively regulated depending on the type of phosphorylation and this might represent a mechanism whereby other cell-signaling systems interfere – cross talk – with the genomic (or nongenomic) actions of 1,25(OH)2D3 [50]. 2. Regulation of Vitamin D Receptor Abundance

Several studies have demonstrated that there is a strong correlation between the VDR level and the biological response in target cells [70]. A wide variety of factors, including PTH [71], pharmacological activators of the

protein kinase A pathway [72], transforming growth factor β (TGFβ) [73], estrogens [74], thyroid hormone [75], glucocorticoids [76], and retinoic acid [77] can alter VDR mRNA levels in a tissue-specific pattern (heterologous regulation). Interestingly, both cell cycle [78] and the differentiation state of cells in culture [79] influence the extent of VDR mRNA expression. Autoregulation of VDR expression has been demonstrated both in vitro and in vivo, again in a very tissuespecific fashion and could serve as a means of signal amplification. It is well established that 1,25(OH)2D3 increases the VDR content by stabilizing the receptor [80, 81]. The role of 1,25(OH)2D3 in the transcriptional regulation of the VDR gene is less evident and 1,25(OH)2D3-inducibility of the hVDR gene has not been found with the part of the promoter characterized [30, 82, 83].

IV. VITAMIN D AND BONE CELLS A. Genomic Versus Nongenomic Action of Vitamin D As a transcription factor, 1,25(OH)2D3 affects the expression of several genes in osteoblasts, osteoclasts, and chondrocytes, thereby regulating cellular growth and differentiation of these cells. Although over 50 genes have been reported to be regulated by 1,25(OH)2D3, only a small number have been reported to contain VDREs. Nongenomic responses to 1,25(OH)2D3 have also been described. They develop at the plasma membrane and comprise rapid (seconds to minutes) changes in ion channel activities, activation of second-messenger pathways, and elevation of cytosolic calcium concentrations [84, 85]. Recently, Zanello and Norman [86] described that the presence of a functional VDR was essential for the modulation of Cl− and Ca2+ channel activities by 1,25(OH)2D3 as well as for 1,25(OH)2D3-dependent rapid exocytose of osteoblast-secretory granules. Moreover, the classical VDR seems to be the 1,25(OH)2D3-binding protein associated with plasma membrane caveolae [87]. Analogously, inactivation of the VDR was found to abrogate rapid 1,25(OH)2D3mediated changes in intracellular Ca2+ in calvarial osteoblasts [88]. A new receptor conformational ensemble model proposes that the VDR has two ligand-binding pockets, which, when occupied by the appropriately shaped ligand will result in the onset of either rapid or genomic responses [89, 90]. Another study however reported that the rapid effect of 1,25(OH)2D3 on intracellular calcium and protein kinase C was independent of VDR [91], suggesting the existence of membrane-associated receptor.

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B. Effects of 1,25(OH)2D3 on Osteoblast Growth, Differentiation, and Apoptosis The general concept is that 1,25(OH)2D3 has a biphasic effect on osteoblasts: it abrogates or stimulates the normal developmental pathway or gene-expression profiles depending on whether it is given during the proliferation or differentiation stage. 1,25(OH)2D3 given during the proliferative period of rat calvaria osteoblast cultures inhibited proliferation and down-regulated collagen synthesis and alkaline phosphatase activity. No increase in osteocalcin expression was observed [92] and nodule formation was blocked [93] suggesting a stop in osteoblast differentiation. In contrast, 1,25(OH)2D3 treatment of mature osteoblasts resulted in up-regulation of some osteoblast-associated genes, such as osteopontin and osteocalcin and in stimulation of calcium accumulation [94]. At this stage of differentiation 1,25(OH)2D3 may promote further maturation of the osteoblast. However, differences in species, cell types, and cell culture conditions, as well as interactions with growth factor and cytokine signaling may result in diverse and sometimes contradictory biological actions of 1,25(OH)2D3. Despite this controversy, the general agreement is in favor of antiproliferative and prodifferentiative properties of 1,25(OH)2D3. In addition, 1,25(OH)2D3 protects osteoblastic cells from undergoing apoptosis, as shown for both human and rat osteoblastic cells treated with several apoptosis-inducing agents [95–97].

C. 1,25(OH)2D3-Regulated Gene Expression Related to Osteoblast Characteristics 1. Cell Cycle Genes

The antiproliferative effect of 1,25(OH)2D3 on several cell types including osteoblasts is due to inhibition of the transition from the G1 to the S-phase of the cell cycle [98, 99]. Cyclin-dependent kinase inhibitors like p21CIP1/WAF1 and p27KIP1 are believed to be the main mediators of this effect [100], although conflicting results on this matter exist [101–104]. A study on 1,25(OH)2D3-induced growth inhibition in murine osteoblasts revealed an early and persistent VDR-dependent down-regulation of cyclin D1 and several genes with key functions in the process of DNA replication. Similar findings in other cell types stress the general nature and the importance of this repressed gene expression in the antiproliferative effect of 1,25(OH)2D3 [105]. 2. TRANSCRIPTION FACTORS IN OSTEOBLAST DIFFERENTIATION

During osteoblast differentiation, 1,25(OH)2D3 regulates the expression of numerous genes belonging to different

families, some of them containing an identified functional VDRE. The expression of the transcription factor Runx2 that is essential for osteoblast differentiation [106], is down-regulated by 1,25(OH)2D3 in rodent osteoblastic cells via the VDR/RXR heterodimer recognizing a functional VDRE [107]. On the other hand, in primary human osteoblasts the effect of 1,25(OH)2D3 on Runx2 mRNA expression was found to be dependent on the duration of the treatment, being inhibitory after 1 h and stimulatory after 48 h [108]. 3. Extracellular Matrix Proteins

The expression of several noncollagenous extracellular matrix proteins is regulated by 1,25(OH)2D3 with osteocalcin being the best characterized. Transcription of the osteocalcin gene is principally regulated by Runx2 through multiple Runx2-binding sites in the promoter. 1,25(OH)2D3 stimulates the expression of osteocalcin strongly in human and rat osteoblastic cells. The VDRE in the rat osteocalcin gene is flanked by Runx2-binding sites and all these regulatory domains contribute to the extensive chromatin reorganization of the involved promoter regions at the time of gene activation [109–111]. In contrast, the expression of the mouse osteocalcin gene is inhibited by 1,25(OH)2D3 [112, 113], which may be explained by differences in the organization of Runx2 motifs in the promoter regions [114]. This illustrates that 1,25(OH)2D3 can play different roles in the regulation of the same gene in various species. Two other genes coding for extracellular matrix proteins have been shown to contain a functional VDRE involved in enhanced transcription by 1,25(OH)2D3: the mouse osteopontin [115] and the rat bone sialoprotein gene [116]. At the same time, genes encoding the proteinases MMP-13 and MT1-MMP [117, 118], functioning in extracellular matrix remodeling, are positively regulated by 1,25(OH)2D3 treatment. 4. Growth Factors

Not only local factors involved in bone remodeling but also their receptors and/or downstream effectors are modulated by 1,25(OH)2D3, but again different responses have been observed depending on the system used and on the stage of cell differentiation. As an example the effect on insulin-like growth factor I (IGF-I) production is either stimulatory or inhibitory, whereas on IGF receptors and IGF-binding proteins it is mainly stimulatory. TGFβ and TGFβ receptors are up-regulated by 1,25(OH)2D3, while Smad proteins that are downstream effectors of TGFβ signaling can interact (as a coactivator) with VDR (for review see reference [119]). A consistent stimulatory effect of 1,25(OH)2D3 is seen on vascular endothelial growth factor (VEGF) [120].

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D. Effects of 1,25(OH)2D3 on Osteoclastogenesis and Osteoclast Activity 1. Indirect Role of 1,25(OH)2D3 on Osteoclastogenesis

1,25(OH)2D3 is a potent stimulator of osteoclastic bone resorption by stimulating osteoclast formation and activity. It has been shown that 1,25(OH)2D3 acts as a fusigen for committed osteoclast precursors [121]. Unlike osteoblasts, osteoclasts do not express a VDR, but osteoclast precursors seem to contain the receptor. Therefore, it is unlikely that 1,25(OH)2D3 acts on mature osteoclasts directly but rather exerts its effects on osteoclastogenesis indirectly by modulating osteoblasts. Recently, it has been shown that 1,25(OH)2D3 acts directly on osteoblasts/ stromal cells to increase the production of membranebound receptor activator of nuclear factor (NF)-kB ligand (RANKL) [122], that plays an essential role in osteoclast differentiation and activation [123]. RANKL acts then via binding to its signal-transducing receptor RANK, present on (pre-)osteoclasts. This induction of RANKL is mediated by VDR/RXR binding to functional VDRE in the promoter region [124, 125]. In addition, up-regulation of macrophage-colony stimulating factor (M-CSF) [126] and suppression of the RANKL-decoy receptor OPG [122, 127, 128] in osteoblastic cells are involved in 1,25(OH)2D3enhanced osteoclast formation. This indirect osteoblastmediated mechanism may explain previous observation of coculture experiments with VDR-deficient mice: no osteoclasts were formed in response to 1,25(OH)2D3 when VDR-null osteoblastic cells were cocultured with wild-type (WT) spleen cells, while normal osteoclasts were present when WT osteoblastic cells and VDR-null spleen cells were cocultured [129]. Interestingly, the action of 1,25(OH)2D3 is not essential for osteoclast formation in vivo since VDR-deficient mice have normal number of osteoclasts [130]. In addition, the expression or signaling of several other cytokines and growth factors, which stimulate (interleukin-1 (IL-1), IL-6, IL-11) or inhibit (IL-4, TGFβ) bone resorption, are regulated by 1,25(OH)2D3 (reviewed in reference [131]). 2. 1,25(OH)2D3-Regulated Genes Involved in Osteoclast Activity

Several markers of the osteoclast phenotype are induced by 1,25(OH)2D3 in cultures of osteoclast precursors. One of them is the vitronectin receptor αvβ3 integrin that enables osteoclasts to adhere to bone-matrix proteins containing the RGD sequence, such as osteopontin, fibronectin, vitronectin, an event critical in the process of

bone resorption [132, 133]. Treatment of avian osteoclast precursors with 1,25(OH)2D3 increases αvβ3 expression [134]. The promoter sequence comprises three hexameric direct repeat half-sites separated by three and nine nucleotides to which, respectively, VDR/RXR and RAR/ RXR heterodimers bind [135]. Another 1,25(OH)2D3-regulated protein is the nonreceptor tyrosine kinase pp60c-src [136], that probably plays an important role in bone resorption since c-src-deficient mice develop osteopetrosis [137]. Chapel et al. [138] showed that 1,25(OH)2D3 accelerates pp60c-src kinasespecific activity without affecting its expression. 1,25(OH)2D3 also increases mRNA levels of carbonic anhydrase-II (CA-II) that produces hydrogen protons necessary to acidify the resorption lacuna. A VDRE consensus sequence has been identified in the promoter of the chicken CA-II gene [139], but the VDR/RXR complex binds to it with an inverse polarity compared to the classical VDRE. In contrast, the 1,25(OH)2D3-induced CA-II expression in murine marrow cultures is suggested not to be the result of increased transcription but rather a part of the differentiation process [140].

E. Effect on Chondrocytes Chondrocytes express the VDR and 1,25(OH)2D3 has a biphasic effect on chondrocyte proliferation, being stimulatory at physiological concentrations (10−12 M) and inhibitory at high concentrations (10−8 M) [141, 142]. Characteristics of chondrocyte differentiation, such as alkaline phosphatase expression, collagen production, and matrix calcification, are affected by 1,25(OH)2D3 treatment in vitro [142, 143]. The expression of and signaling by several factors modulating chondrocyte development, such as IGF-I and IGF-I receptor, is influenced by 1,25(OH)2D3 [142]. 1,25(OH)2D3 has also been reported to increase apoptosis in mouse hypertrophic chondrocytes [144].

V. PATHOLOGY AND THERAPY RELATED TO VITAMIN D AVAILABILITY, METABOLISM, AND FUNCTION A. Pathology Vitamin D is either primarily or secondarily involved in many metabolic bone diseases. Abnormalities in vitamin D availability, metabolism, or action are responsible for calciopenic rickets/osteomalacia. During development, defective chondrocyte mineralization occurs at the growth plates and contributes to disorganization of chondrocyte

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arrangement. This leads to epiphyseal widening, a typical hallmark of rickets, retardation of longitudinal bone growth and skeletal deformities. In osteomalacia, the characteristic pathology changes involve impaired mineralization of bone matrix or osteoid during bone remodeling, manifested by a decrease in the mineralizing surface and mineralization rate. This results in the accumulation of large surfaces of unmineralized osteoid. Focus will be on the types of rickets or osteomalacia caused by genetic defect while diseases that are acquired or secondary to renal or gastrointestinal dysfunction will only be briefly mentioned. 1. Vitamin D Deficiency

Classic nutritional rickets, caused by the simultaneous deprivation of sunlight exposure and dietary vitamin D [145] (Table I) has become uncommon because this vitamin is frequently added to dairy products. In the absence of such health policies, mild or severe cases of rickets are still endemic, especially in countries where calcium intake is low (e.g. northern China, many African countries) and/or exposure to sunshine is avoided for cultural reasons. Vitamin D deficiency is not a primary cause of osteoporosis but it can be secondarily involved. Indeed a low vitamin D status is quite common in the elderly and by inducing secondary hyperparathyroidism it certainly accelerates the negative balance of bone turnover [146]. Moreover, vitamin D metabolism or action is frequently disturbed in secondary osteoporosis. For instance, this fat-soluble vitamin is subject to malabsorption, and levels may be low in chronic liver disease and primary biliary cirrhosis. Chronic renal failure results in renal rickets and secondary hyperparathyroidism when compromised renal mass reduces 1α-hydroxylase activity. Some anticonvulsant drugs increase the turnover of vitamin D into inactive compounds, resulting in a decrease in serum levels of 1,25(OH)2D3.

Table I. 1.

Substrate availability

2.

Metabolism Genetic

Acquired 3.

Vitamin D action Genetic

Tumor-induced osteomalacia is a syndrome of renal phosphate wasting and low or inappropriate normal levels of 1,25(OH)2D3 [147]. 2. Hereditary Defects

Genetic defects causing calciopenic rickets include disorders with abnormal vitamin D metabolism or resistance to the effects of the active form of vitamin D. Genetic defects are found at several steps of this pathway but rickets are part of the clinical picture in only two disorders. A specific hereditary defect in the gene coding for the 25-hydroxyvitamin D-1α hydroxylase (1α-hydroxylase) or CYP27B1 impairs the final, critical step in the biosynthesis of 1,25(OH)2D3 resulting in vitamin D-dependent rickets type I (VDDR-I), also called pseudo-vitamin D-deficiency (PDDR) [148, 149]. This disease is characterized by failure to thrive, muscle weakness, skeletal deformities, hypocalcemia, secondary hyperparathyroidism and low serum 1,25(OH)2D3 concentrations caused by impaired activity of the 1α-hydroxylase [148]. Several mutations in the VDR are responsible for vitamin D-dependent rickets type II (VDDR-II), also called hereditary hypocalcemic vitamin D-resistant rickets (HVDRR) [150, 151]. The greatest number of these mutations can be localized to the DNA-binding zinc finger region and a smaller group to the ligand-binding domain, some of them affecting the heterodimerization of VDR with RXR. HVDRR results in a phenotype characterized by early-onset rickets, hypocalcemia, secondary hyperparathyroidism, and markedly increased 1,25(OH)2D3 levels. The alopecia observed in some kindreds with mutant VDRs has not been observed in vitamin D deficiency, suggesting that VDR not only mediates the bone mineral homeostatic actions of vitamin D but also plays a role in the differentiation of hair follicles. These patients do not respond to physiological

Pathology: Vitamin D-Related Calcium/Bone Diseases

Mechanism ↓ Intake ↓ Sun exposure

Pathology Rickets/Osteomalacia

DBP CYP24

KO mice KO mice

CYP27B1

KO mice Human-chr12q13

Chronic renal failure Oncogenic osteomalacia VDR

KO mice Human-chr12q13

More susceptible to osteomalacia Defect in intramembranous ossification Rickets HVDRR I Renal osteodystrophy Osteomalacia and hypophosphatemia Rickets HVDRR II

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doses of 1,25(OH)2D3, but the bone pathology is cured by frequent intravenous infusions of calcium [152] and is prevented by high oral doses of calcium [153]. Both of these two diseases display an autosomalrecessive trait, but clinical features and response to administrated 1,25(OH)2D3 are distinct. The phenotypes of mice with targeted ablation of 1α-hydroxylase or VDR exhibit the clinical abnormalities observed in the VDDR-I and -II patients respectively and offered opportunity to investigate in vivo functions of 1,25(OH)2D3 and the molecular basis of its actions. a. VDR-Null Mice: Model of Vitamin D-dependent Rickets Type II a.1. Mineral homeostasis and rickets of VDR-null mice VDR-null or VDR-mutant mice have been created by four different groups [131, 154–156]. They are born phenotypically normal, but show growth retardation starting after weaning. At that age, hypocalcemia, hyperparathyroidism, and hypophosphatemia develop and 1,25(OH)2D3 serum levels become 10- to 100-fold elevated, coincident with extremely low 24,25(OH)2D3 serum levels. Inspection and X-ray analysis of long bones showed a reduced length and the typical features of rickets including widening of epiphyseal growth plates, thinning of the cortex, cupping, and widening of the metaphysis. The time of onset of symptoms at 3 weeks of age suggests that the vitamin D endocrine system is principally required for maintaining bone mineral homeostasis when the organism is deprived of a consistent and plentiful supply of calcium, such as occurs after weaning in mammals. Amling and coworkers [157] studied histomorphometric and biomechanical parameters in Boston VDR-null mice, fed regular chow. They found by the age of 70 days an expanded length of

Figure 4

the growth plate and disorganization of the chondrocyte columns, a marked increase in bone volume due to increased osteoid, an increased number of osteoblasts, but normal number of osteoclasts, impaired mineral apposition rate, and reduced stiffness. However, normalization of mineral ion homeostasis (calcemia and phosphatemia) by feeding these mice a diet enriched with calcium, lactose, and phosphorus (the socalled “rescue diet”) prevented hyperparathyoidism and rickets [157, 158]. These findings are consistent with the therapeutic effect of high doses of calcium in VDRR type II (see Section V.A.2). In addition, the ratio of Ca/P in the diet is a major determinant for rescue of bone mineralization and turnover in VDR-null mice, with the optimal ratio being a factor of 2 [159, 160]. The underlying mechanism is suggested to be a Ca/P ratio-dependent intestinal transport of calcium and phosphorus. These data suggest that, in an ideal metabolic environment, the VDR is not essential for the development or maintenance of normal bone, although 1,25(OH)2D3 has been shown to have significant effects on the expression of genes by osteoblasts and osteoclasts. Thus the principal action of the VDR is its role in intestinal calcium absorption and/or renal calcium reabsorption. Intestinal active calcium absorption, assessed in two VDR-null strains, was reduced to one third of normal mice [155]. At the molecular level only the expression of the epithelial calcium channels TRPV6 and TRPV5 was severely impaired in the absence of a functional VDR, with less effect on calbindin-D9K and the plasma membrane calcium ATPase (PMCAIb), [155] (Fig. 4). The urinary calcium excretion in VDR-null mice under a normal diet is inappropriately high given the hypocalcemia

Gene targets of 1,25(OH)2D3 action in the enterocyte. In the absence of a functional VDR, active intestinal calcium absorption is impaired reflected at the molecular level by severely decreased gene expression of TRPV6 but only modest decreases in calbindin-D9K and PMCA1b expression. The thickness of the arrow reflects VDR-related changes.

Chapter 18 The Vitamin D Hormone and its Nuclear Receptor

in these mice [156, 161]. Normalization of serum calcium levels by the rescue diet resulted in a manifest increase of calcium excretion suggesting that renal tubular calcium reabsorption is impaired in VDR-null mice. The parameter that correlated best herewith was the expression of calbindin-D9K which was found to be decreased in all strains of VDR-null mice, both on the normal and the rescue diet [131, 155, 156, 161, 162]. a.2. Mechanisms underlying the disorganization of growth plate Rickets or the expansion and disorganization of the growth plate are a characteristic feature of VDR inactivation (Fig. 5). Analysis of growth plate morphology demonstrated normal resting and proliferating chondrocyte layers, but expansion of the hypertrophic chondrocyte layer. This latter finding could potentially be a consequence of an osteoclast defect, since 1,25(OH)2D3 has been shown to be a potent inducer of RANKL produced by osteoblasts (see Section IV.D). However, histomorphometric analyses have demonstrated normal numbers of osteoclasts in the metaphyses of several strains of VDR-null mice, indicating that other osteoclast-activating factors such as PTH and interleukin-1α are capable of inducing the formation of mature resorbing osteoclasts in vivo in the absence of VDR and that the VDR is not essential for osteoclastmediated bone resorption. This also suggests that the expansion of the hypertrophic chondrocyte layer is not secondary to an osteoclast defect. Osteoclasts are brought to the hypertrophic chondrocytes concomitant with vascular invasion. This process is initiated by secretion of VEGF, a signaling molecule produced by hypertrophic chondrocytes. No impairment of VEGF RNA production by these

Figure 5

317

cells was observed in VDR-null mice, indicating that hypertrophic chondrocytes were capable of signaling vascular invasion [163]. Alternatively, the expansion of the hypertrophic chondrocyte layer may be due to altered chondrocyte development in VDR-null mice, but present data are not conclusive. A recent study shows that in VDR-null mice, this feature is not secondary to increased chondrocyte proliferation or impaired differentiation but rather a consequence of decreased apoptosis [163]. The authors also postulate that hypophosphatemia, rather than hypocalcemia, is the major determinant of decreased chondrocyte apoptosis. However, excess PTH secretion in these mice might mimic the PTHrP paracrine effects on chondrocyte survival [164]. In contrast, no manifest alteration in the number of apoptotic cells or in the expression of apoptosis-related genes was detected in another VDR-null strain [165]. In addition, a functionally redundant VDR may be present. To investigate this possibility, mice deficient in both VDR and RXRγ were generated since RXRγ-null mice exhibit no discernible abnormalities [165]. The phenotype observed in double-null mutant mice was basically similar to those in VDR-null mice, except that growth plate development was more severely impaired and was not prevented by dietary supplementation. A selective impairment of hypertrophic chondrocyte development was observed despite normalized mineral homeostasis produced by the high-mineral diet. These findings indicate that the combined actions of VDR- and RXRγmediated signals are essential for normal development of growth plate cartilage and might be explained by a functional,

Molecular mechanisms of bone pathology in VDR-null mice. Inactivation of VDR leads to increased bone volume, mainly due to increased osteoid, which normalizes when a rescue diet with an adequate Ca/P ratio is given. The disorganization of the growth plate in VDR-null mice is not due to defective osteoclast formation or chondrocyte proliferation as both processes are normal in these mice. VEGF gene expression by hypertrophic chondrocytes is normal, suggesting normal vascular invasion of the growth plate. Rather, apoptosis of hypertrophic chondrocytes is impaired, possibly as a consequence of hypophosphatemia, although the effect of increased PTH levels can not be excluded.

318 but redundant, VDR-like receptor in growth plate chondrocytes. b. 1a-Hydroxylase-Null Mice: Model of Vitamin DDependent Type Rickets I Mice with inactivation of 1a-hydroxylase(1α(OH)ase) were generated and demonstrate failure to thrive [166, 167]. Regarding calcium homeostasis, 1α(OH)ase-null models demonstrate hypocalcemia, severe secondary hyperparathyroidism, hypophosphatemia, elevated serum alkaline phosphatase, and increased phosphaturia, analogous to the phenotype of the VDR-null mice. However, 1α(OH)ase-null mice genuinely lack the capacity to synthesize 1,25(OH)2D3: circulating levels of 1,25(OH)2D3 were undetectable, whereas circulating levels of 25(OH)D3 doubled. This emphasizes the non-redundant role played by the renal 1α(OH)ase in producing the hormonally active metabolite of vitamin D. Furthermore, this confirms that the phenotype of the 1α(OH)ase-null mice matches the clinical manifestations of VDDR-I [168]. Typical features of advanced rickets are observed histologically in bone of 1α(OH)ase-null mice: disorganization and widening of the columnar alignment of hypertrophic chondrocytes, resulting in increased width of the growth plate, impaired calcification of the hypertrophic cartilage and accumulation of osteoid in trabecular and cortical bone. Feeding the 1α(OH)ase-null mice a rescue diet normalizes serum calcium levels and trabecular bone volume, but not the width of the growth plate [169]. A phenotypic rescue, even with normalization of the growth plate, was also seen after treating the 1α(OH)ase-null mice with 1,25(OH)2D3, the treatment of choice for VDRR-I therapy.

B. 1,25(OH)2D3 and Analogs as Therapeutics in Bone Pathology The role of 1,25(OH)2D3 in the treatment of bone disorders characterized by defective mineralization, such as rickets and renal osteodystrophy, is well established. Moreover prevention or correction of subclinical 1,25(OH)2D3 deficiency can substantially reduce the incidence of hip and other fractures in senile osteoporosis, at least when given in combination with oral calcium supplementation [170]. The major goal is the reduction of PTH secretion and the subsequent decreased bone resorption. However, 1,25(OH)2D3 is also suggested as therapeutic for osteoporosis, a disorder characterized by decreased bone formation or at least a discrepancy between bone formation and resorption [171]. Indeed, a large New Zealand study demonstrated a major reduction in the number of new vertebral fractures during 1,25(OH)2D3

GEERT CARMELIET ET AL .

therapy [172]. However, the optimal therapeutic range for 1,25(OH)2D3 appears to be quite narrow and may vary among patients. An alternative therapy can be established with the vitamin D3 analog, 1α(OH)D3 (alfacalcidiol). 1α(OH)D3 is less effective on intestinal calcium absorption compared with its effect on bone because it must first be hydroxylated at the 25-position to become fully active [173]. Nevertheless, therapeutic dose monitoring is also needed to avoid side effects such as hypercalcemia and hypercalciuria. Efforts to develop new synthetic analogs for bone diseases have been modest. Presently some analogs are being evaluated in preclinical or human clinical phase II trials for the prevention or cure of osteoporosis. The vitamin D analog, 2β-(3-hydroxypropoxy)-1,25(OH)2D3 (ED-71), is characterized by high calcemic activity and strong binding to DBP (two-fold that of 1,25(OH)2D3) [174]. In normal or ovariectomized rats, ED-71 stimulates bone mass and bone formation rates [175] and is a more potent inhibitor of bone resorption than 1α(OH)D3 [176]. ED-71 is in final stages of clinical osteoporosis trials. Several other compounds such as 1α-(OH)D2 and 19-nor-1,25(OH)2D3 analogs as well as 2-methylene-19nor-20-epi-1α,25(OH)2D3 (2MD); 1αfluoro-25(OH)16,23diene-26,27bishomo-20-epi vitamin D3 (Ro 26-9228), and GS 1790 are now being evaluated for the same purpose in preclinical or early clinical settings. In vitro studies demonstrated that the analog 2MD was 100 times more potent in osteoclast-inducing activity compared with 1,25(OH)2D3, probably related to its induction of increased VDR interaction with RXR and co-activators (SRC-1 and DRIP 205) [177]. 2MD also induced numerous mineralized bone nodules (10−12 M) in primary human osteoblast precursors and fetal mouse calvarial cells, whereas 1α,25(OH)2D3 exhibited little effect even at 10−8 M [178]. 2MD increased total body bone mineral density in ovariectomized rats in a 23-week period, whereas 1,25(OH)2D3 only prevented the bone loss [178]. Three weeks of oral treatment of ovariectomized rats with Ro 269228 had a bone-protecting effect without inducing hypercalcemia. Analysis of biochemical and bone histomorphometrical indices suggested that Ro 26-9228 inhibited bone resorption and increased the number of differentiated osteoblasts. Gene expression was differentially regulated in duodenum and bone in analog-treated rats [179]. This tissue-selective action of the analog has been explained by significant interactions of Ro 26-9228-VDR with RXR and cofactors such as GRIP and DRIP in osteoblasts but much less in intestinal CaCo-2 cells [180]. Bone disease in patients with chronic renal failure is due to a complex set of mechanisms such as impaired 1,25(OH)2D3 synthesis, vitamin D resistance, secondary hyperparathyroidism, and abnormal mineral handling

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Chapter 18 The Vitamin D Hormone and its Nuclear Receptor

(hyperphosphatemia, aluminum or fluoride excess, acidosis). Whereas 1α-(OH)D3 and 1,25(OH)2D3 are widely used for the prevention and cure of renal osteodystrophy, several analogs have been evaluated for this indication with the aim of better PTH suppression with less risk for inducing hypercalcemia or hyperphosphatemia [181]. In the United States, 19-nor-1,25(OH)2D2 (paricalcitol, Zemplar®) and 1α(OH)D2 (doxercalciferol, Hecterol®) are available for the treatment of secondary hyperparathyroidism in renal-failure patients. 22-oxacalcitriol (OCT, maxacalcitol, Oxarol®) and 1,25-dihydroxy-26,26,26, 27,27,27-hexafluorovitamin D3 (falecalcitriol, Hornel®, and Fulstan®) have been approved in Japan. 22-Oxacalcitriol has similar effects as 1,25(OH)2D3 on cell proliferation and inhibition of PTH secretion in vitro and in vivo [182] in experimental animals and patients with chronic renal failure [183]. Similarly, the vitamin D2 analogs 19-nor1,25(OH)2D2 [184] and 1α(OH)D2 [185] and the analog falecalcitriol [186] have been evaluated in vivo for suppressing PTH levels, together with the effects on calcium and phosphate levels [187]. A double-blind, randomized, multicenter study, comparing the efficacy of paricalcitol and calcitriol in renal disease patients undergoing hemodialysis demonstrated that paricalcitol reduced PTH concentration more rapidly, with fewer sustained episodes of either hypercalcemia or increased Ca × P product than 1,25(OH)2D3 therapy [188]. An uncontrolled, retrospective assessment of a clinical data base of about 67000 patients undergoing hemodialysis and receiving either paricalcitol or calcitriol, demonstrated that paricalcitol was associated with a lower mortality rate (especially by cardiovascular events) over the 36-month follow-up period (18%) than calcitriol (22.3%). However, such an uncontrolled study has serious limitations and properly designed, randomized, controlled trials are still needed to confirm these data [189].

VI. CONCLUSIONS Insight into the molecular mechanisms of 1,25(OH)2D3 action, via binding to VDR and interacting with VDRE promoting altered transcriptional activity, has enlarged considerably during the last years. Many elements of this complex machinery are not yet fully identified; the mechanism of negatively regulated genes by 1,25(OH)2D3 especially remains an open question. Certainly the participation of co-activators or co-repressors has been established and their functional significance elucidated. In addition, the knowledge of the three-dimensional structure of DBP and of the VDR provided valuable information for the understanding of ligand–protein and protein–protein interactions.

The development of several KO mice (DBP, megalin, VDR, 1α-hydroxylase, and 24-hydroxylase) strengthens the importance of 1,25(OH)2D3–VDR system in mineral and bone homeostasis. The exact function of 1,25(OH)2D3 in bone metabolism awaits new physiological in vivo models with tissue-specific inactivation of the gene of interest.

References 1. Calli, J. J., and Russell, D. W. (1991). Characterization of human sterol 27-hydroxylase. A mitochondrial cytochrome P-450 that catalyzes multiple oxidation reaction in bile acid biosynthesis. J. Biol. Chem. 266, 7774–7778. 2. Rosen, H., Reshef, A., Maeda, N., Lippoldt, A., Shpizen, S., Triger, L., Eggertsen, G., Bjorkhem, I., and Leitersdorf, E. (1998). Markedly reduced bile acid synthesis but maintained levels of cholesterol and vitamin D metabolites in mice with disrupted sterol 27-hydroxylase gene. J. Biol. Chem. 273, 14805–14812. 3. Cheng, J. B., Levine, M. A., Bell, N. H., Mangelsdorf, D. J., and Russell, D. W. (2004). Genetic evidence that the human CYP2R1 enzyme is a key vitamin D 25-hydroxylase. Proc. Natl Acad. Sci. USA 101, 7711–7715. 4. St-Arnaud, R., Messerlian, S., Moir, J. M., Omdahl, J. L., and Glorieux, F. H. (1997). The 25-hydroxyvitamin D 1-alpha-hydroxylase gene maps to the pseudovitamin D-deficiency rickets (PDDR) disease locus. J. Bone Miner. Res. 12, 1552–1559. 5. Takeyama, K., Kitanaka, S., Sato, T., Kobori, M., Yanagisawa, J., and Kato, S. (1997). 25-hydroxyvitamin D3 1α-hydroxylase and vitamin D synthesis. Science 277, 1827–1830. 6. Monkawa, T., Yoshida, T., Wakino, S., Shinki, T., Anazawa, H., DeLuca, H. F., Suda, T., Hayashi, M., and Saruta, T. (1997). Molecular cloning of cDNA and genomic DNA for human 25-hydroxyvitamin D3 1α-hydroxylase. Biochem. Biophys. Res. Comm. 239, 527–533. 7. Shinki, T., Shimada, H., Wakino, S., Anazawa, H., Hayashi, M., Saruta, T., DeLuca, H. F., and Suda, T. (1997). Cloning and expression of rat 25-hydroxyvitamin D3-1α-hydroxylase cDNA. Proc. Natl Acad. Sci. USA 94, 1290–1295. 8. Fu, G. K., Lin, D., Zhang, M. Y., Bikle, D. D., Shackleton, C. H. L., Miller, W. L., and Portale, A. A. (1997). Cloning of human 25-hydroxyvitamin D-1α-hydoxylase and mutations causing vitamin Ddependent rickets type 1. Molec. Endocrinol. 11, 1961–1970. 9. Verboven, C., Rabijns, A., De Maeyer, M., Van Baelen, H., Bouillon, R., and De Ranter, D. (2002). A structural basis for the unique binding features of the human vitamin D-binding protein. Nat. Struct. Biol. 9, 131–136. 10. Safadi, F. F., Thornton, P., Magiera, H., Hollis, B. W., Gentile, M., Haddad, J. G., Liebhaber, S. A., and Cooke, N. E. (1999). Osteopathy and resistance to vitamin D toxicity in mice null for vitamin D binding protein. J. Clin. Invest. 103, 239–251. 11. Nykjaer, A., Dragun, D., Walther, D., Vorum, H., Jacobsen, C., Herz, J., Melsen, F., Christensen, E. I., and Willnow, T. E. (1999). An endocytic pathway essential for renal uptake and activation of the steroid 25-(OH) Vitamin D3. Cell 96, 507–515. 12. Leheste, J. R., Melsen, F., Wellner, M., Jansen, P., Schlichting, U., Renner-Müller, I., Andreassen, T. T., Wolf, E., Bachmann, S., Nykjaer, A., and Willnow, T. E. (2003). Hypocalcemia and osteopathy in mice with kidney-specific megalin gene defect. FASEB J. 17, 247–249. 13. Ohyama, Y., Noshiro, M., Eggertsen, G., Gotoh, O., Kato, Y., Bjorkhem, I., and Okuda, K. (1993). Structural characterization of the

320

14.

15.

16.

17.

18.

19.

20.

21.

22.

23.

24.

25.

26.

27.

28.

29.

30.

gene encoding rat 25-hydroxyvitamin D3 24-hydoxylase. Biochemistry 32, 76–82. Lohnes, D., and Jones, G. (1987). Side chain metabolism of vitamin D3 in osteosarcoma cell line UMR-106. Characterization of products. J. Biol. Chem. 262, 14394–14401. Reddy, G. S., and Tserng, K. Y. (1989). Calcitroic acid, end product of renal metabolism of 1,25-dihydroxyvitamin D3 through C24 oxidation pathway. Biochemistry 28, 1763–1769. Akiyoshi-Shibata, M., Sakaki, T., Ohyama, Y., Noshiro, M., Okuda, K., and Yabusaki, Y. (1994). Further oxidation of hydroxycalcidiol by calcidiol 24-hydroxylase. A study with the mature enzyme expressed in Escherichia Coli. Eur. J. Biochem. 224, 335–343. St.-Arnaud, R., Arabian, A., Travers, R., Barletta, F., Raval-Pandya, M., Chapin, K., Depovere, J., Mathieu, C., Christakos, S., Demay, M. B., and Glorieux, F. H. (2000). Deficient mineralization of intramembranous bone in vitamin D-24-hydroxylase-ablated mice is due to elevated 1,25dihydroxyvitamin D and not to the absence of 24,25-dihydroxyvitamin D. Endocrinology 141, 2658–2666. Yamaura, M., Nakamura, T., Nagai, Y., Yoshishara, A., and Suzuki, K. (1993). Reduced mechanical competence of bone by ovariectomy and its prevention with 24R,25-dihydroxyvitamin D3 administration in beagles. Calcif. Tissue Int. 25, 49–56. Schwartz, Z., Bonewald, L. F., Caulfield, K., Brooks, B., and Boyan, B. D. (1993). Direct effects of transforming growth factor-beta on chondrocytes are modulated by vitamin D metabolites in a cell maturation-specific manner. Endocrinology 132, 1544–1552. Mangelsdorf, D. J., Thummel, C., Beato, M., Herrlich, P., Schütz, G., Umesono, K., Blumberg, B., Kastner, P., Mark, M., Chambon, P., and Evans, R. M. (1995). The nuclear receptor superfamily: the second decade. Cell 83, 835–839. Li, Y. C., Bergwitz, C., Jüppner, H., and Demay, M. B. (1997). Cloning and characterization of the vitamin D receptor from Xenopus laevis. Endocrinology 138, 2347–2353. McDonnell, D. P., Mangelsdorf, D. J., Pike, J. W., Haussler, M. R., and O’Malley, B. W. (1987). Molecular cloning of complementary DNA encoding the avian receptor for vitamin D. Science 235, 1214–1217. Elaroussi, M. A., Prahl, J. M., and DeLuca, H. F. (1994). The avian vitamin D receptors: primary structures and their origins. Proc. Natl Acad. Sci. USA 91, 11596–11600. Kamei, Y., Kawada, T., Fukuwatari, T., Ono, T., Kato, S., and Sugimoto, E. (1995). Cloning and sequencing of the gene encoding the mouse vitamin D receptor. Gene 152, 281–282. Burmester, J., Wiese, R., Maeda, N., and DeLuca, H. F. (1988). Structure and regulation of the rat 1,25-dihydroxyvitamin D3 receptor. Proc. Natl Acad. Sci. USA 85, 9499–9502. Baker, A. R., McDonnell, D. P., Hughes, M., Crisp, T. M., Mangelsdorf, D. J., Haussler, M. R., Pike, J. W., Shine, J., and O’Malley, B. W. (1988). Cloning and expression of full-length cDNA encoding human vitamin D receptor. Proc. Natl Acad. Sci. USA 85, 3294–3298. Suzuki, T., Suzuki, N., Srivastava, A. S., and Kurokawa, T. (2000). Identification of cDNAs encoding two subtypes of vitamin D receptor in flounder, Paralichthys olivaceus. Biochem. Biophys. Res. Commun. 270, 40–45. Sugiura, S. H., and Ferraris, R. P. (2004). Dietary phosphorus-responsive genes in the intestine, pyloric ceca, and kidney of rainbow trout. Am. J. Physiol. 287, R541–550. Whitfield, G. K., Dang, H. T., Schluter, S. F., Bernstein, R. M., Bunag, T., Manzon, L. A., Hsieh, G., Dominguez, C. E., Youson, J. H., Haussler, M. R., and Marchalonis, J. J. (2003). Cloning of a functional vitamin D receptor from the lamprey (Petromyzon marinus), an ancient vertebrate lacking a calcified skeleton and teeth. Endocrinology 144, 2704–2716. Miyamoto, K., Kesterson, R. A., Yamamoto, H., Taketani, Y., Nishiwaki, E., Tatsumi, S., Inoue, Y., Morita, K., Takeda, E., and

GEERT CARMELIET ET AL .

31.

32.

33.

34.

35.

36. 37. 38.

39.

40.

41.

42.

43.

44.

45.

46.

47.

48.

49.

Pike, J. W. (1997). Structural organization of the human vitamin D receptor chromosomal gene and its promoter. Mol. Endocrinol. 11, 1165–1179. Crofts, L. A., Hancock, M. S., Morrison, N. A., and Eisman, J. A. (1998). Multiple promoters direct the tissue-specific expression of novel N-terminal variant human vitamin D receptor gene transcripts. Proc. Natl Acad. Sci. USA. 95, 10529–10534. Sunn, K. L., Cock, T. A., Crofts, L. A., Eisman, J. A., and Gardiner, E. M. (2001). Novel N-terminal variant of human VDR. Mol. Endocrinol. 15, 1599–1609. Jurutka, P. W., Remus, L. S., Whitfield, G. K., Thompson, P. D., Hsieh, J. C., Zitzer, H., Tavakkoli, P., Galligan, M. A., Dang, H. T., Haussler, C. A., and Haussler, M. R. (2000). The polymorphic N terminus in human vitamin D receptor isoforms influences transcriptional activity by modulating interaction with transcription factor IIB. Mol. Endocrinol. 14, 401–420. Ames, S. K., Ellis, K. J., Gunn, S. K., Copeland, K. C., and Abrams, S. A. (1999). Vitamin D receptor gene Fok1 polymorphism predicts calcium absorption and bone mineral density in children. J. Bone Miner. Res. 14, 740–746. Harris, S. S., Eccleshall, T. R., Gross, C., Dawson-Hughes, B., and Feldman, D. (1997). The vitamin D receptor start codon polymorphism (FokI) and bone mineral density in premenopausal American black and white women. J. Bone Miner. Res. 12, 1043–1048. Khorasanizadeh, S., and Rastinejad, F. (2001). Nuclear-receptor interactions on DNA-response elements. Trends Biochem. Sci. 26, 384–390. Freedman, L. P. (1992). Anatomy of the steroid receptor zinc finger region. Endocr. Rev. 13, 129–145. Rastinejad, F., Perlmann, T., Evans, R. M., and Sigler, P. B. (1995). Structural determinants of nuclear receptor assembly on DNA direct repeats. Nature 375, 203–211. Shaffer, P. L., and Gewirth, D. T. (2002). Structural basis of VDR-DNA interactions on direct repeat response elements. EMBO J. 21, 2242–2252. Luo, Z., Rouvinen, J., and Maenpaa, P. H. (1994). A peptide C-terminal to the second Zn finger of human vitamin D receptor is able to specify nuclear localization. Eur. J. Biochem. 223, 381–387. Hsieh, J. C., Shimizu, Y., Minoshima, S., Shimizu, N., Haussler, C. A., Jurutka, P. W., and Haussler, M. R. (1998). Novel nuclear localization signal between the two DNA-binding zinc fingers in the human vitamin D receptor. J. Cell. Biochem. 70, 94–109. Michigami, T., Suga, A., Yamazaki, M., Shimizu, C., Cai, G., Okada, S., and Ozono, K. (1999). Identification of amino acid sequence in the hinge region of human vitamin D receptor that transfers a cytosolic protein to the nucleus. J. Biol. Chem. 274, 33531–33538. Allegretto, E. A., Pike, J. W., and Haussler, M. R. (1987). C-terminal proteolysis of the avian 1,25-dihydroxyvitamin D3 receptor. Biochem. Biophys. Res. Commun. 147, 479–485. Rochel, N., Wurtz, J. M., Mitschler, A., Klaholz, B., and Moras, D. (2000). Crystal structure of the nuclear receptor for vitamin D bound to its natural ligand. Mol. Cell. 5, 173–179. Bouillon, R., Okamura, W. H., and Norman, A. W. (1995). Structurefunction relationships in the vitamin D endocrine system. Endocr. Rev. 16, 200–257. Choi, M., Yamamoto, K., Masuno, H., Nakashima, K., Taga, T., and Yamada, S. (2001). Ligand recognition by the vitamin D receptor. Bioorg. Med. Chem. 9, 1721–1730. Sone, T., Ozono, K., and Pike, J. W. (1991). A 55-kilodalton accessory factor facilitates vitamin D receptor DNA binding. Mol. Endocrinol. 5, 1578–1586. Kliewer, S. A., Umesono, K., Mangelsdorf, D. J., and Evans, R. M. (1992). Retinoid X receptor interacts with nuclear receptors in retinoic acid, thyroid hormone and vitamin D3 signaling. Nature 355, 446–449. Carlberg, C., Bendik, I., Wyss, A., Meier, E., Sturzenbecker, L. J., Grippo, J. F., and Hunziker, W. (1993). Two nuclear signalling pathways for vitamin D. Nature 361, 657–660.

Chapter 18 The Vitamin D Hormone and its Nuclear Receptor 50. Christakos, S., Raval-Pandya, M., Werny, J., and Yang, W. (1996). Genomic mechanisms involved in the pleiotropic actions of 1,25dihydroxyvitamin D3. Biochem. J. 316, 361–371. 51. Carlberg, C. (1995). Mechanisms of nuclear signalling by vitamin D3. Interplay with retinoid and thyroid hormone signalling. Eur. J. Biochem. 231, 517–527. 52. Jin, C. H., Kerner, S. A., Hong, M. H., and Pike, J. W. (1996). Transcriptional activation and dimerization functions in the human vitamin D receptor. Mol. Endocrinol. 10, 945–957. 53. Schrader, M., Nayeri, S., Kahlen, J. P., Muller, K. M., and Carlberg, C. (1995). Natural vitamin D3 response elements formed by inverted palindromes: polarity-directed ligand sensitivity of vitamin D3 receptor-retinoid X receptor heterodimer-mediated transactivation. Mol. Cell. Biol. 15, 1154–1161. 54. Haussler, M. R., Whitfield, G. K., Haussler, C. A., Hsieh, J. C., Thompson, P. D., Selznick, S. H., Dominguez, C. E., and Jurutka, P. W. (1998). The nuclear vitamin D receptor: biological and molecular regulatory properties revealed. J. Bone Miner. Res. 13, 325–349. 55. Kimmel-Jehan, C., Darwish, H. M., Strugnell, S. A., Jehan, F., Wiefling, B., and DeLuca, H. F. (1999). DNA bending is induced by binding of vitamin D receptor-retinoid X receptor heterodimers to vitamin D response elements. J. Cell. Biochem. 74, 220–228. 56. Bettoun, D. J., Burris, T. P., Houck, K. A., Buck, D. W. 2nd, Stayrook, K. R., Khalifa, B., Lu, J., Chin, W. W., and Nagpal, S. (2003). Retinoid X receptor is a nonsilent major contributor to vitamin D receptor-mediated transcriptional activation. Mol. Endocrinol. 17, 2320–2328. 57. Rachez, C., and Freedman, L. P. (2000). Mechanisms of gene regulation by vitamin D3 receptor: a network of coactivator interactions. Gene. 246, 9–21. 58. Rachez, C., Gamble, M., Chang, C. P., Atkins, G. B., Lazar, M. A., and Freedman, L. P. (2000). The DRIP complex and SRC-1/p160 coactivators share similar nuclear receptor binding determinants but constitute functionally distinct complexes. Mol. Cell. Biol. 20, 2718–2726. 59. Auboeuf, D., Honig, A., Berget, S. M., and O’Malley, B. W. (2002). Coordinate regulation of transcription and splicing by steroid receptor coregulators. Science 298, 416–419. 60. Kitagawa, H., Fujiki, R., Yoshimura, K., Mezaki, Y., Uematsu, Y., Matsui, D., Ogawa, S., Unno, K., Okubo, M., Tokita, A., Nakagawa, T., Ito, T., Ishimi, Y., Nagasawa, H., Matsumoto, T., Yanagisawa, J., and Kato, S. (2003). The chromatin-remodeling complex WINAC targets a nuclear receptor to promoters and is impaired in Williams syndrome. Cell 113, 905–917. 61. MacDonald, P. N., Sherman, D. R., Dowd, D. R., Jefcoat, S. C. Jr., and DeLisle, R. K. (1995). The vitamin D receptor interacts with general transcription factor IIB. J. Biol. Chem. 270, 4748–4752. 62. Blanco, J. C. G., Wang, I. M., Tsai, S. Y., Tsai, M. J., O’Malley, B. W., Jurutka, P. W., Haussler, M. R., and Ozato, K. (1995). Transcription factor TFIIB and the vitamin D receptor cooperatively activate liganddependent transcription. Proc. Natl Acad. Sci. USA 92, 1535–1539. 63. Masuyama, H., Jefcoat, S. C. Jr., and MacDonald, P. N. (1997). The N-terminal domain of trancription Factor IIB is required for direct interaction with the vitamin D receptor and participates in vitamin Dmediated transcription. Mol. Endocrinol. 11, 218–228. 64. Kurihara, N., Reddy, S. V., Araki, N., Ishizuka, S., Ozono, K., Cornish, J., Cundy, T., Singer, F. R., and Roodman, G. D. (2004). Role of TAFII-17, a VDR binding protein, in the increased osteoclast formation in Paget’s Disease. J. Bone Miner. Res. 19, 1154–1164. 65. Brown, T. A., and DeLuca, H. F. (1990). Phosphorylation of the 1,25-dihydroxyvitamin D3 receptor. A primary event in 1,25dihydroxyvitamin D3 action. J. Biol. Chem. 265, 10025–10029. 66. Jurutka, P. W., Hsieh, J-C., Nakajima, S., Haussler, C. A., Whitfield, G. K., and Haussler, M. R. (1996). Human vitamin D receptor phosphorylation by casein kinase II at Ser-208 potentiates transcriptional activation. Proc. Natl Acad. Sci. USA 93, 3519–3524.

321

67. Bareletta, F., Freedman, L. P., and Christakos, S. (2002). Enhancement of VDR-mediated transcription by phosphorylation: correlation with increased interaction between the VDR and DRIP205, a subunit of the VDR-interacting protein coactivator complex. Mol. Endocrinol. 16, 301–314. 68. Hsieh, J-C., Jurutka, P. W., Nakajima, S., Galligan, M. A., Haussler, C. A., Shimizu, Y., Shimizu, N., Whitfield, G. K., and Haussler, M. R. (1993). Phosphorylation of the human vitamin D receptor by protein kinase C. Biochemical and functional evaluation of the serine 51 recognition site. J. Biol. Chem. 268, 15118–15126. 69. Nakajima, S., Yamagata, M., Sakai, N., and Ozono, K. (2000). Effect of cyclic adenosine 3′,5′-monophosphate and protein kinase A on ligand-dependent transactivation via the vitamin D receptor. Mol. Cell. Endocrinol. 159, 45–51. 70. Chen, T. L., and Feldman, D. (1981). Regulation of 1,25dihydroxyvitamin D3 receptors in cultured mouse bone cells. Correlation of receptor concentration with the rate of cell division. J. Biol. Chem. 256, 5561–5566. 71. Pols, H. A. P., van Leeuwen, J. P. T. M., Schilte, J. P., Visser, T. J., and Birkenhager, J. C. (1988). Heterologous up-regulation of the 1,25dihydroxyvitamin D3 receptor by parathyroid hormone (PTH) and PTH-like peptide in osteoblast-like cells. Biochem. Biophys. Res. Commun. 156, 588–594. 72. van Leeuwen, J. P. T. M., Birkenhager, J. C., Wijngaarden, T. V. V., vanden Bemd, G. J. C. M., and Pols, H. A. P. (1992). Regulation of 1,25dihydroxyvitamin D3 receptor gene expression by parathyroid hormone and cAMP-agonists. Biochem. Biophys. Res. Commun. 185, 881–886. 73. Staal, A., Van Wijnen, A. J., Desai, R. K., Pols, H. A., Birkenhager, J. C., DeLuca, H. F., Denhardt, D. T., Stein, J. L., van Leeuwen, J. P., Stein, G. S., and Lian, J. B. (1996). Antagonistic effects of transforming growth factor-beta on vitamin D3 enhancement of osteocalcin and osteopontin transcription: reduced interactions of vitamin D receptor/ retinoid X receptor complexes with vitamin D response elements. Endocrinology 137, 2001–2011. 74. Walters, M. R. (1981). An estrogen-stimulated 1,25-dihydroxyvitamin D3 receptor in rat uterus. Biochem. Biophys. Res. Commun. 103, 721–726. 75. Mahonen, A., Pirskanen, A., and Maenpaa, P. H. (1991). Homologous and heterlologous regulation of 1,25-dihydroxyvitmain D-3 receptor mRNA levels in human osteosarcoma cells. Biochim. Biophys. Acta 188, 111–118. 76. Chen, T. L., Cone, C. M., Morey-Holton, E., and Feldman, D. (1983). 1,25-Dihydroxyvitmain D3 receptors in cultured rat osteoblasts-like cells. Glucocorticoid treatment increases receptor content. J. Biol. Chem. 258, 4350–4355. 77. Petkovitch, P. M., Heersche, J. N. M., Tinker, D. O., and Jones, G. (1984). Retinoic acid stimulates 1,25-dihydroxyvitamin D3 binding in rat osteosarcoma cells. J. Biol. Chem. 259, 8274–8290. 78. Krishnan, A. V., and Feldman, D. (1991). Stimulation of 1,25dihydroxyvitamin D3 receptor gene expression in cultured cells by serum and growth factors. J. Bone Miner. Res. 6, 1099–1107. 79. Zhao, X., and Feldman, D. (1993). Regulation of vitamin D receptor abundance and responsiveness during differentiation of HT-29 human colon cancer cells. Endocrinology 132, 1808–1814. 80. Wiese, R. J., Uhland-Smith, A., Ross, T. K., Prahl, J. M., and DeLuca, H. F. (1992). Up-regulation of the vitamin D receptor in response to 1,25-dihydroxyvitamin D3 results from ligand-induced stabilization. J. Biol. Chem. 267, 20082–20086. 81. Arbour, N. C., Prahl, J. M., and DeLuca, H. F. (1993). Stabilization of the vitamin D receptor in rat osteosarcoma cells through the action of 1,25(OH)2D3. Mol. Endocrinol. 7, 1307–1312. 82. Jehan, F., and DeLuca, H. F. (1997). Cloning and characterization of the mouse vitamin D receptor promoter. Proc. Natl Acad. Sci. USA 94, 10138–10143. 83. Byrne, I. M., Flanagan, L., Tenniswood, M. P., and Welsh, J. (2000). Identification of a hormone-responsive promoter immediately upstream

322

84.

85.

86.

87.

88.

89.

90.

91.

92.

93.

94.

95.

96.

97.

98.

99.

of exon 1c in the human vitamin D receptor gene. Endocrinology 141, 2829–2836. Caffrey, J. M., and Farach-Carson, M. C. (1989). Vitamin D3 metabolites modulate dihydropyridine-sensitive calcium currents in clonal rat osteosarcoma cells. J. Biol. Chem. 264, 20265–20274. Zanello, L. P., and Norman, A. W. (1997). Stimulation by 1α,25(OH)2-vitamin D3 of whole cell chloride currents in osteoblastic ROS 17/2.8 cells. A structure-function study. J. Biol. Chem. 272, 22617–22622. Zanello, L. P., and Norman, A. W. (2004). Rapid modulation of osteoblast ion channel responses by 1α,25(OH)2-vitamin D3 require the presence of a functional vitamin D nuclear receptor. Proc. Natl Acad. Sci. USA 101, 1589–1594. Huhtakangas, J. A., Olivera, C. J., Bishop, J. E., Zanello, L. P., and Norman, A. W. (2004). The vitamin D receptor is present in caveolaeenriched plasma membranes and binds 1α,25(OH)2-vitamin D3 in vivo and in vitro. Mol. Endocrinol. 18, 2660–2671. Erben, R. G., Soegiarto, D. W., Weber, K., Zeitz, U., Lieberherr, M., Gniadecki, R., Moller, G., Adamski, J., and Balling, R. (2002). Deletion of deoxyribonucleic acid binding domain of the vitamin D receptor abrogates genomic and non genomic functions of vitamin D. Mol. Endocrinol. 16, 1524–1537. Mizwicki, M. T., Keidel, D., Bula, C. M., Bishop, J. E., Zanello, L. P., Wurtz, J. M., Moras, D., Norman, A. W. (2004). Identification of an alternative ligand-binding pocket in the nuclear vitamin D receptor and its functional importance in 1α,25(OH)2-vitamin D3 signalling. Proc. Natl Acad. Sci. USA 101, 12876–12881. Norman, A. W., Mizwicki, M. T., and Norman, D. P. G. (2004). Steroid-hormone rapid actions, membrane receptors and a conformational ensemble model. Nat. Rev. Drug Discov. 3, 27–41. Wali, R. K., Kong, J., Sitrin, M. D., Bissonnette, M., and Li, Y. C. (2003). Vitamin D receptor is not required for the rapid actions of 1,25dihydroxyvitamin D3 to increase intracellular calcium and activate protein kinase C in mouse osteoblasts. J. Cell. Biochem. 88, 794–801. Ishida, H., Bellows, C. G., Aubin, J. E., and Heersche, J. N.M. (1993). Characterizaion of the 1,25(OH)2D3-induced inhibition of bone nodule formation in long-term cultures of fetal rat calvaria cells. Endocrinology 132, 61–66. Owen, T. A., Aronow, M. S., Barone, L. M., Bettencourt, B., Stein, G. S., and Lian, J. B. (1991). Pleiotropic effects of vitamin D on osteoblast gene expression are related to the proliferative and differentiated state of the bone cell phenotype: Dependency upon basal levels of gene expression, duration of exposure and bone matrix competency in normal rat osteoblast cultures. Endocrinology 128, 1496–1504 Matsumoto, T., Igarashi, C., Taksuchi, Y., Harada, S., Kikuchi, T., Yamamoto, H., and Ogata, E. (1991). Stimulation by 1,25-dihydroxyvitamin D3 of in vitro mineralization induced by osteoblast-like MC3T3-E1 cells. Bone 12, 27–32. Hansen, C. M., Hansen, D., Holm, P. K., and Binderup, L. (2001). Vitamin D compounds exert anti-apoptotic effects in human osteosarcoma cells in vitro. J. Steroid Biochem. Mol. Biol. 77, 1–11. Duque, G., El Abdaimi, K., Henderson, J. E., Lomri, A., and Kremer, R. (2004). Vitamin D inhibits Fas ligand-induced apoptosis in human osteoblasts by regulating components of both the mitochondrial and Fas-related pathways. Bone 35, 57–64. Morales, O., Samuelsson, M. K., Lindgren, U., and Haldosen, L. A. (2004). Effects of 1α,25-dihydroxyvitamin D3 and growth hormone on apoptosis and proliferation in UMR 106 osteoblast-like cells. Endocrinology 145, 87–94. Wang, Q. M., Jones, J. B., and Studzinski, G. P. (1996). Cyclindependent kinase inhibitor p27 as a mediator of the G1-S phase block induced by 1,25-dihydroxyvitamin D3 in HL60 cells. Cancer Res. 56, 264–267. Verlinden, L., Verstuyf, A., Convents, R., Marcelis, S., Van Camp, M., and Bouillon, R. (1998). Action of 1,25(OH)2D3 on the cell cycle

GEERT CARMELIET ET AL .

100.

101.

102.

103.

104.

105.

106.

107.

108.

109.

110.

111.

112.

113.

114.

genes, cyclin D1, p21 and p27 in MCF-7 cells. Mol. Cell. Endocrinol. 142, 57–65. Liu, M., Lee, M. H., Cohen, M., Bommakanti, M., and Freedman, L. P. (1996). Transcriptional activation of the Cdk inhibitor p21 by vitamin D3 leads to the induced differentiation of the myelomonocytic cell line U937. Genes Dev. 10, 142–153. Akutsu, N., Lin, R., Bastien, Y., Bestawros, A., Enepekides, D. J., Black, M. J., and White, J. H. (2001). Regulation of gene expression by 1α,25-dihydroxyvitamin D3 and its analog EB1089 under growthinhibitory conditions in squamous carcinoma cells. Mol. Endocrinol. 15, 1127–1139. Hager, G., Formanek, M., Gedlicka, C., Thurnher, D., Knerer, B., and Kornfehl, J. (2001). 1,25(OH)2 vitamin D3 induces elevated expression of the cell cycle-regulating genes P21 and P27 in squamous carcinoma cell lines of the head and neck. Acta Otolaryngol. 121, 103–109. Hershberger, P. A., Modzelewski, R. A., Shurin, Z. R., Rueger, R. M., Trump, D. L., and Johnson, C. S. (1999). 1,25-Dihydroxycholecalciferol (1,25-D3) inhibits the growth of squamous cell carcinoma and down-modulates p21Waf1/Cip1 in vitro and in vivo. Cancer Res. 59, 2644–2649. Wade, W. N., Willingham, M. C., Koumenis, C., and Cramer, S. D. (2002). p27Kip1 is essential for the antiproliferative action of 1,25dihydroxyvitamin D3 in primary, but not immortalized, mouse embryonic fibroblasts. J. Biol. Chem. 277, 37301–37306. Eelen, G., Verlinden, L., Van Camp, M., Van Hummelen, P., Marchal, K., De Moor, B., Mathieu, C., Carmeliet, G., Bouillon, R., and Verstuyf, A. (2004). The effects of 1alpha,25-dihydroxyvitamin D3 on the expression of DNA replication genes. J. Bone Miner. Res. 19, 133–146. Ducy, P., Zhang, R., Geoffroy, V., Ridall, A. L., and Karsenty, G. (1997). Osf2/Cbfa1: a transcriptional activator of osteoblast differentiation. Cell 89, 747–754. Drissi, H., Pouliot, A., Koolloos, C., Stein, J. L., Lian, J. B., Stein, G. S., and van Wijnen, A. J. (2002). 1,25-(OH)2-vitamin D3 suppresses the bone-related Runx2/Cbfa1 gene promoter. Exp. Cell Res. 274, 323–333. Viereck, V., Siggelkow, H., Tauber, S., Raddatz, D., Schutze, N., and Hufner, M. (2002). Differential regulation of Cbfa1/Runx2 and osteocalcin gene expression by vitamin-D3, dexamethasone, and local growth factors in primary human osteoblasts. J. Cell. Biochem. 86, 348–356. Javed, A., Gutierrez, S., Montecino, M., van Wijnen, A. J., Stein, J. L., Stein, G. S., and Lian, J. B. (1999). Multiple Cbfa/AML sites in the rat osteocalcin promoter are required for basal and vitamin D-responsive transcription and contribute to chromatin organization. Mol. Cell. Biol. 19, 7491–7500. Sierra, J., Villagra, A., Paredes, R., Cruzat, F., Gutierrez, S., Javed, A., Arriagada, G., Olate, J., Imschenetzky, M., Van Wijnen, A. J., Lian, J. B., Stein, G. S., Stein, J. L., and Montecino, M. (2003). Regulation of the bone-specific osteocalcin gene by p300 requires Runx2/Cbfa1 and the vitamin D3 receptor but not p300 intrinsic histone acetyltransferase activity. Mol. Cell. Biol. 23, 3339–3351. Gutierrez, S., Liu, J., Javed, A., Montecino, M., Stein, G. S., Lian, J. B., and Stein, J. L. (2004). The vitamin D response element in the distal osteocalcin promoter contributes to chromatin organization of the proximal regulatory domain. J. Biol. Chem. 279, 43581–43588. Lian, J. B., Shalhoub, V., Aslam, F., Frenkel, B., Green, J., Hamrah, M., Stein, G. S., and Stein, J. L. (1997). Species-specific glucocorticoid and 1,25-dihydroxyvitamin D responsiveness in mouse MC3T3-E1 osteoblasts: dexamethasone inhibits osteoblast differentiation and vitamin D down-regulates osteocalcin gene expression. Endocrinology 138, 2117–2127. Zhang, R., Ducy, P., and Karsenty, G. (1997). 1,25-dihydroxyvitamin D3 inhibits osteocalcin expression in mouse through an indirect mechanism. J. Biol. Chem. 272, 110–116. Ducy, P., and Karsenty, G. (1995). Two distinct osteoblast-specific cis-acting elements control expression of a mouse osteocalcin gene. Mol. Cell. Biol. 15, 1858–1869.

Chapter 18 The Vitamin D Hormone and its Nuclear Receptor 115. Noda, M., Vogel, R. L., Craig, A. M., Prahl, J., Deluca, H. F., and Denhardt, D. T. (1990). Identification of a DNA sequence responsible for binding of the 1,25-dihydroxyvitamin D3 receptor and 1,25dihydroxyvitamin D3 enhancement of mouse secreted phophoprotein 1 (Spp-1 or osteopontin) gene expression. Proc. Natl Acad. Sci. USA 87, 9995–9999. 116. Kim, R. H., Li, J. J., Ogata, Y., Yamauchi, M., Freedman, L. P., and Sodek, J. (1996). Identification of a vitamin D3-response element that overlaps a unique inverted TATA box in the rat bone sialoprotein gene. Biochem. J. 318, 219–226. 117. Uchida, M., Shima, M., Chikazu, D., Fujieda, A., Obara, K., Suzuki, H., Nagai, Y., Yamato, H., and Kawaguchi, H. (2001). Transcriptional induction of matrix metalloproteinase-13 (collagenase-3) by 1α,25dihydroxyvitamin D3 in mouse osteoblastic MC3T3-E1 cells. J. Bone Miner. Res. 16, 221–230. 118. Luo, X. H., and Liao, E. Y. (2003). 1α,25-dihydroxyvitamin D3 regulates the expression of membrane-type matrix metalloproteinase-1 in normal human osteoblast-like cells. Endocr. Res. 29, 353–362. 119. Van Driel, M., Pols, H. A. P., and van Leeuwen, J. P. T. M. (2004). Osteoblast differentiation and control by vitamin D and vitamin D metabolites. Curr. Pharm. Des. 10, 2535–2555. 120. Wang, D. S., Miura, M., Demura, H., and Sato, K. (1997). Anabolic effects of 1,25-dihydroxyvitamin D3 on osteoblasts are enhanced by vascular endothelial growth factor produced by osteoblasts and by growth factors produced by endothelial cells. Endocrinology 138, 2953–2962. 121. Takahashi, N., Udagawa, N., Tanaka, S., Murakami, H., Owan, I., Tamura, T., and Suda, T. (1993). Postmitotic osteoclast precursors are mononuclear cells which express macrophage-associated phenotypes. Dev. Biol. 163, 212–221. 122. Nakashima, T., Kobayashi, Y., Yamasaki, S., Kawakami, A., Eguchi, K., Sasaki, H., and Sakai, H. (2000). Protein expression and functional difference of membrane-bound and soluble receptor activator of NF-kappaB ligand: modulation of the expression by osteotropic factors and cytokines. Biochem. Biophys. Res. Commun. 275, 768–775. 123. Lacey, D. L., Timms, E., Tan, H. L., Kelley, M. J., Dunstan, C. R., Burgess, T., Elliott, R., Colombero, A., Elliott, G., Scully, S., Hsu, H., Sullivan, J., Hawkins, N., Davy, E., Capparelli, C., Eli, A., Qian, Y. X., Kaufman, S., Sarosi, I., Shalhoub, V., Senaldi, G., Guo, J., Delaney, J., and Boyle, W. J. (1998). Osteoprotegerin ligand is a cytokine that regulates osteoclast differentiation and activation. Cell 93, 165–176. 124. Kitazawa, R., and Kitazawa, S. (2002). Vitamin D3 augments osteoclastogenesis via vitamin D-responsive element of mouse RANKL gene promoter. Biochem. Biophys. Res. Commun. 290, 650–655. 125. Kitazawa, S., Kajimoto, K., Kondo, T., and Kitazawa, R. (2003). Vitamin D3 supports osteoclastogenesis via functional vitamin D response element of human RANKL gene promoter. J. Cell. Biochem. 89, 771–777. 126. Rubin, J., Fan, X., Thornton, D., Bryant, R., and Biskobing, D. (1996). Regulation of murine osteoblast macrophage colony-stimulating factor production by 1,25(OH)2D3. Calcif. Tissue Int. 59, 291–296. 127. Horwood, N. J., Elliott, J., Martin, T. J., and Gillespie, M. T. (1998). Osteotropic agents regulate the expression of osteoclast differentiation factor and osteoprotegerin in osteoblastic stromal cells. Endocrinology 139, 4743–4746. 128. Kondo, T., Kitazawa, R., Maeda, S., and Kitazawa, S. (2004). 1α,25dihydroxyvitamin D3 rapidly regulates the mouse osteoprotegerin gene through dual pathways. J. Bone Miner. Res. 19, 1411–1419. 129. Takeda, S., Yoshizawa, T., Nagai, Y., Yamato, H., Fukumoto, S., Sekine, K., Kato, S., Matsumoto, T., and Fujita, J. (1997). Stimulation of osteoclast formation by 1,25-dihydroxyvitamin D requires its binding to vitamin D receptor (VDR) in osteoblastic cells: studies using VDR knockout mice. Endocrinology 140, 1005–1008. 130. Yoshizawa, T., Handa, Y., Uematsu, Y., Takesa, S., Sekine, K., Yoshihara, Y., Kawakami, T., Arioka, K., Sato, H., Uchiyama, Y.,

131.

132. 133. 134.

135.

136.

137.

138.

139.

140.

141.

142.

143.

144.

145.

146. 147.

148.

149.

323 Masushige, S., Fukamizu, A., Matsumoto, T., and Kato, S. (1997). Mice lacking the vitamin D receptor exhibit impaired bone formation, uterine hypoplasia and growth retardation after weaning. Nat. Genet. 16, 391–396. Gurlek, A., Pittelkow, M. R., and Kumar, R. (2002). Modulation of growth factor/cytokine synthesis and signaling by 1α,25dihydroxyvitamin D3: implications in cell growth and differentiation. Endocr. Rev. 23, 763–786. Roodman, G. D. (1996). Advances in bone biology: the osteoclast. Endocr. Rev. 17, 308–332. Rodan, S. B., and Rodan, G. A. (1997). Integrin function in osteoclasts. J. Endocrinol. 154, S47–S56. Mimura, H., Cao, X., Ross, F. P., Chiba, M., and Teitelbaum, S. L. (1994). 1,25-dihydroxyvitamin D3 transcriptionally activates the β3-integrin subunit gene in avian osteoclast precursors. Endocrinology 134, 1061–1066. Cao, X., Ross, F. P., Zhang, L., MacDonald, P. N., Chappel, J., and Teitelbaum, S. L. (1993). Cloning of the promoter for the avian integrin β3 subunit gene and its regulation by 1,25-dihydroxyvitamin D3. J. Biol. Chem. 268, 27371–27380. Yoneda, T., Niewolna, M., Lowe, C., Izbicka, E., and Mundy, G. R. (1993). Hormonal regulation of pp60c-src expression during osteoclast formation in vitro. Mol. Endocrinol. 7, 1313–1318. Soriano, P., Montgomery, C., Geske, R., and Bradley, A. (1991). Targeted disruption of the c-src proto-oncogene leads to osteopetrosis in mice. Cell 64, 693–702. Chappel, J., Ross, F. P., Abu-Amer, Y., Shaw, A., and Teitelbaum, S. L. (1997). 1,25-dihydroxyvitamin D3 regulates pp60c-src activity and expression of a pp60c-src activating phosphatase. J. Cell. Biochem. 67, 432–438. Quelo, I., Kahlen, J. P., Rascle, A., Jurdic, P., and Carlberg, C. (1994). Identification and characterization of a vitamin D3 response element of chicken carbonic anhydrase-II. DNA Cell Biol. 13, 1181–1187. Biskobing, D. M., Fan, D., Fan, X., and Rubin, J. (1997). Induction of carbonic anhydrase II expression in osteoclast progenitors requires physical contact with stromal cells. Endocrinology 138, 4852–4857. Klaus, G., Weber, L., Rodriguez, J., Fernandez, P., Klein, T., Grulich-Henn, J., Huegel, U., Ritz, E., and Mehls, O. (1998). Interaction of IGF-I and 1,25(OH)2D3 on receptor expression and growth stimulation in rat growth plate chondrocytes. Kidney Int. 53, 1152–1161. Krohn, K., Haffner, D., Hugel, U., Himmele, R., Klaus, G., Mehls, O., and Schaefer, F. (2003). 1,25(OH)2D3 and dihydrotestoterone interact to regulate proliferation and differentiation of epiphyseal chondrocytes. Calcif. Tissue Int. 73, 400–410. Schwartz, Z., Schlader, D. L., Ramirez, V., Kennedy, M. B., and Boyan, B. D. (1989). Effects of vitamin D metabolites on collagen production and cell proliferation of growth zone and resting zone cartilage cells in vitro. J. Bone Miner. Res. 4, 199–207. Donohue, M. M., and Demay, M. B. (2002). Rickets in VDR null mice is secondary to decreased apoptosis of hypertrophic chondrocytes. Endocrinology 143, 3691–3694. Schmidt-Gayk, H., Bouillon, R., and Roth, H. J. (1997). The determination of vitamin D metabolites (calcidiol and calcitriol) and their clinical significance. Scand. J. Clin. Lab. Invest. 57, 35–45. Bouillon, R., Carmeliet, G., and Boonen, S. (1997). Ageing and calcium metabolism. Baill. Clin. Endocrinol. Metab. 11, 341–365. Brame, L. A., White, K. E., and Econs, M. J. (2004). Renal phosphate wasting disorders: clinical features and pathogenesis. Semin. Nephrol. 24, 39–47. Fu, G. K., Lin, D., Zhang, M. Y., Bikle, D. D., Shackleton, C. H. L., Miller, W. L., and Portale, A. A. (1997). Cloning of human 25-hydroxyvitamin D-1α-hydroxylase and mutations causing vitamin D-dependent rickets type 1. Molec. Endocrinol. 11, 1961–1970. Kitanaka, S., Takeyama, K., Murayama, A., Sato, T., Okumura, K., Nogami, M., Hasegawa, Y., Niimi, H., Yanagisawa, J., Tanaka, T., and Kato, S. (1998). Inactivating mutations in the 25-hydroxyvitamin D3

324

150.

151.

152.

153. 154.

155.

156.

157.

158.

159.

160.

161.

162.

163.

164.

GEERT CARMELIET ET AL . 1α-hydroxylase gene in patients with pseudovitamin D-deficiency rickets. N. Engl. J. Med. 338, 653–661. Fraser, D., Kooh, S. W., Kind, H. P., Holick, M. F., Tanaka, Y., and DeLuca, H. F. (1973). Pathogenesis of hereditary vitamin D-dependent rickets. An inborn error of vitamin D metabolism involving defective conversion of 25-hydroxyvitamin D to 1α,25-hydroxyvitamin D. N. Engl. J. Med. 289, 817–822. Malloy, P. J., Pike, J. W., and Feldman, D. (1999). The vitamin D receptor and the syndrome of hereditary 1,25-dihydroxyvitamin Dresistant rickets. Endocr. Rev. 20, 156–188. Balsan, S., Garabedian, M., Larchet, M., Gorski, A. M., Cournot, G., Tau, C., Bourdeau, A., Silve, C., and Ricour, C. (1986). Long-term nocturnal calcium infusions can cure rickets and promote normal mineralization in hereditary resistance to A,25-dihydroxyvitamin D. J. Clin. Invest. 77, 1661–1667. Hochberg, Z., Tiosano, D., and Even, L. (1992). Calcium therapy for calcitriol-resistant rickets. J. Pediatr. 121, 803–808. Li, Y. C., Pirro, A. E., Amling, M., Delling, G., Baron, R., Bronson, R., and Demay, M. B. (1997). Targeted ablation of the vitamin D receptor: an animal model of vitamin D-dependent rickets type II with alopecia. Proc. Natl Acad. Sci. USA 94, 9831–9835. Van Cromphaut, S. J., Dewerchin, M., Hoenderop, J. G. J., Stockmans, I., Van Herck, E., Kato, S., Bindels, R. J. M., Collen, D., Carmeliet, P., Bouillon, R., and Carmeliet, G. (2001). Duodenal calcium absorption in vitamin D receptor-knockout mice: Functional and molecular aspects. Proc. Natl Acad. Sci. USA 98, 13324–13329. Erben, R. G., Soegiarto, D. W., Weber, K., Zeitz, U., Lieberherr, M., Gniadecki, R., Moller, G., Adamski, J., and Balling, R. (2002). Deletion of deoxyribonucleic acid binding domain of the vitamin D receptor abrogates genomic and nongenomic functions of vitamin D. Mol. Endocrinol. 16, 1524–1537. Amling, M., Priemel, M. H. T., Chapin, K., Rueger, J. M., Baron, R., and Demay, M. B. (1999). Rescue of the skeletal phenotype of vitamin D receptor ablated mice in the setting of normal mineral ion homeostasis: formal histomorphometric and biomechanical analyses. Endocrinology 140, 4982–4987. Li, Y. C., Amling, M., Pirro, A. E., Priemel, M., Meuse, J., Baron, R., Delling, G., and Demay, M. B. (1998). Normalization of mineral ion homeostasis by dietary means prevents hyperparathyroidism, rickets, and osteomalacia, but not alopecia in vitamin D receptor-ablated mice. Endocrinology 139, 4391–4396. Masuyama, R., Nakaya, Y., Tanaka, S., Tsurukami, H., Nakamura, T., Watanabe, S., Yoshizawa, T., Kato, S., and Suzuki, K. (2001). Dietary phosphorus restriction reverses the impaired bone mineralization in vitamin D receptor knockout mice. Endocrinology 142, 494–497. Masuyama, R., Nakaya, Y., Katsumata, S., Kajita, Y., Uehara, M., Tanaka, S., Sakai, A., Kato, S., Nakamura, T., and Suzuki, K. (2003). Dietary calcium and phosphorus ratio regulates bone mineralization and turnover in vitamin D receptor knockout mice by affecting intestinal calcium and phosphorus absorption. J. Bone Miner. Res. 18, 1217–1226. Li, Y. C., Bolt, M. J. G., Cao, L. P., and Sitrin, M. D. (2001). Effects of vitamin D receptor inactivation on the expression of calbindins and calcium metabolism. Am. J. Physiol. Endocrinol. Metab. 281, E558–E564. Li, Y. C., Pirro, A. E., and Demay, M. B. (1998). Analysis of vitamin D-dependent calcium-binding protein messenger ribonucleic acid expression in mice lacking the vitamin D receptor. Endocrinology 139, 847–851. Donohue, M. M., and Demay, M. B. (2002). Rickets in VDR null mice is secondary to decreased apoptosis of hypertrophic chondrocytes. Endocrinology 143, 3691–3694. Amling, M., Neff, L., Tanaka, S., Inoue, D., Kuida, K., Weir, E., Philbrick, W. M., Broadus, A. E., and Baron, R. (1997). Bcl-2 lies downstream of parathyroid hormone-related peptide in a signaling

165.

166.

167.

168. 169.

170.

171. 172.

173.

174.

175.

176.

177.

178.

179.

180.

pathway that regulates chondrocyte maturation during skeletal development. J. Cell Biol. 136, 205–213. Yagishita, N., Yamamoto, Y., Yoshizawa, T., Sekine, K., Uematsu, Y., Murayama, H., Nagai, Y., Krezel, W., Chambon, P., Matsumoto, T., and Kato, S. (2001). Aberrant growth plate development in VDR/RXRγ double null mutant mice. Endocrinology 142, 5332–5341. Dardenne, O., Prud’homme, J., Arabian, A., Glorieux, F. H., and St-Arnaud, R. (2001). Targeted inactivation of the 25-hydroxyvitamin D3-1α-hydroxylase gene (CYP27B1) creates an animal model of pseudovitamin D-deficiency rickets. Endocrinology 142, 3135–3141. Panda, D. K., Miao, D., Tremblay, M. L., Sirois, J., Farookhi, R., Hendy, G. N., and Goltzman, D. (2001). Targeted ablation of the 25-hydroxyvitamin D 1α-hydroxylase enzyme: Evidence for skeletal, reproductive, and immune dysfunction. Proc. Natl. Acad. Sci. USA 98, 7498–7503. Bouillon, R. (1998). The many faces of rickets. N. Engl. J. Med. 338, 653–661. Panda, D. K., Miao, D., Bolivar, I., Li, J., Huo, R., Hendy, G. N., and Goltzman D. (2004). Inactivation of the 25-hydroxyvitamin D 1α-hydroxylase and vitamin D receptor demonstrates independent and interdependent effects of calcium and vitamin D on skeletal and mineral homeostasis. J. Biol. Chem. 279, 16754–16766. Chapuy, M., Arlot, M., Duboeuf, F., Brun, J., Crouzet, B., Arnaud, S., Delmas, P. D., and Meunier, P. J. (1992). Vitamin D3 and calcium to prevent hip fractures in elderly women. N. Eng. J. Med. 327, 1637–1642. Civitelli, R. (1995). The role of vitamin D metabolism in the treatment of osteoporosis. Calcif. Tissue Int. 57, 409–414. Tilyard, M. W., Spears, G. F., Thomson, J., and Dovey, S. (1992). Treatment of postmenopausal osteoporosis with calcitriol or calcium. N. Engl. J. Med. 326, 357–362. Akesson, K., Lau, K-H. W., and Baylink, D. J. (1997). Rationale for active vitamin D analog therapy in senile osteoporosis. Calcif. Tissue Int. 60, 100–105. Okano, T., Tsugawa, N., Masuda, S., Takeuchi, A., Kabayashi, T., Takita, Y., and Nishii, Y. (1989). Regulatory activities of 2β(3-hydroxypropoxy)-1α,25-dihydroxy- vitamin D3, a novel synthetic vitamin D3 derivative, on calcium metabolism. Biochem. Biophys. Res. Commun. 163, 1444–1449. Tsurukami, H., Nakamura, T., Suzuki, K., Sato, K., Higuchi, Y., and Nishii, Y. (1994). A novel synthetic vitamin D analogue, 2β(3-hydroxypropoxy)1α,25-dihydroxyvitamin D3 (ED-71), increases bone mass by stimulating the bone formation in normal and ovariectomized rats. Calcif. Tissue Int. 54, 142–149. Uchiyama, Y., Higuchi, Y., Takeda, S., Masaki, T., Shira-Ishi, A., Sato, K., Kubodera, N., Ikeda, K., and Ogata, E. (2002). ED-71, a vitamin D analog, is a more potent inhibitor of bone resorption than alfacalcidol in an estrogen-deficient rat model of osteoporosis. Bone 30, 582–588. Yamamoto, H., Shevde, N. K., Warrier, A., Plum, L. A., DeLuca, H. F., and Pike, J. W. (2003). 2-Methylene-19-nor-(20S)1,25-dihydroxyvitamin D3 potently stimulates gene-specific DNA binding of the vitamin D receptor in osteoblasts. J. Biol. Chem. 278, 31756–31765. Shevde, N. K., Plum, L. A., Clagett-Dame, M., Yamamoto, H., Pike, J. W., and DeLuca, H. F. (2002). A potent analog of 1α,25dihydroxyvitamin D3 selectively induces bone formation. Proc. Natl. Acad. Sci. USA 99, 13487–13491. Peleg, S., Uskokovic, M., Ahene, A., Vickery, G., and Avnur, Z. (2002). Cellular and molecular events associated with the bone-protecting activity of the noncalcemic vitamin D analog Ro-26–9228 in osteopenic rats. Endocrinology 143, 1625–1636. Peleg, S., Ismail, A., Uskokovic, M. R., and Avnur, Z. (2003). Evidence for tissue- and cell-type selective activation of the vitamin D receptor by Ro-26–9228, a noncalcemic analog of vitamin D3. J. Cell. Biochem. 88, 267–273.

Chapter 18 The Vitamin D Hormone and its Nuclear Receptor 181. Brown, A. J., Dusso, A. S., and Slatopolsky, E. (2002). Vitamin D analogues for secondary hyperparathyroidism. Nephrol. Dial. Transplant. 17 Suppl 10, 10–19. 182. Brown, A. J., Ritter, C. R., Finch, J. L., Morrissey, J., Martin, K., Murayama, E., Nishii, Y., and Slatopolsky, E. (1989). The noncalcemic analogue of vitamin D, 22-oxacalcitriol, suppresses parathyroid hormone synthesis and secretion. J. Clin. Invest. 84, 728–732. 183. Kurokawa, K., Akizawa, T., Suzuki, M., Akita, T., Ogata, E., and Slatopolsky, E. (1996). Effect of 22-oxacalcitriol on hyperparathyroidism of dialysis patients: results of a preliminary study. Nephrol. Dial. Transplant. 11, 121–124. 184. Lindberg, J., Martin, K. J., Gonzalez, E. A., Acchiardo, S. R., Valdin, J. R., and Soltanek, C. (2001). A long-term, multicenter study of the efficacy and safety of paricalcitol in end-stage renal disease. Clin. Nephrol. 56, 315–323. 185. Maung, H. M., Elangovan, L., Frazao, J. M., Bower, J. D., Kelley, B. J., Acchiardo, S. R., Rodriguez, H. J., Norris, K. C., Sigala, J. F., Rutkowski, M., Robertson, J. A., Goodman, W. G., Levin, B. S., Chesney, R. W., Mazess, R. B., Kyllo, D. M., Douglass, L. L.,

186.

187.

188.

189.

325 Bishop, C. W., and Coburn, J. W. (2001). Efficacy and side effects of intermittent intravenous and oral doxercalciferol (1α-hydroxyvitamin D2) in dialysis patients with secondary hyperparathyroidism: a sequential comparison. Am. J. Kidney Dis. 37, 532–543. Nishizawa, Y., Morii, H., Ogura, Y., and DeLuca, H. F. (1991). Clinical trial of 26,26,26,27,27,27-hexafluoro-1,25-dihydroxyvitamin D3 in uremic patients on hemodialysis: preliminary report. Contrib. Nephrol. 90, 196–203. Akiba, T., Marumo, F., Owada, A., Kurihara, S., Inoue, A., Chida, Y., Ando, R., Shinoda, T., Ishida, Y., and Ohashi, Y. (1998). Controlled trial of falecalcitriol versus alfacalcidol in suppression of parathyroid hormone in hemodialysis patients with secondary hyperparathyroidism. Am. J. Kidney Dis. 32, 238–246. Sprague, S. M., Llach, F., Amdahl, M., Taccetta, C., and Batlle, D. (2003). Paricalcitol versus calcitriol in the treatment of secondary hyperparathyroidism. Kidney Int. 63, 1483–1490. Teng, M., Wolf, M., Lowrie, E., Ofsthun, N., Lazarus, J. M., and Thadhani, R. (2003). Survival of patients undergoing hemodialysis with paricalcitol or calcitriol therapy. N. Engl. J. Med. 349, 446–456.

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Chapter 19

Sex Steroid Effects on Bone Metabolism David G. Monroe, Ph.D. Thomas C. Spelsberg, Ph.D. and S. Khosla, M.D.

Biochemistry and Molecular Biology, Mayo Clinic College of Medicine, Rochester, MN Professor of Biochemistry, Mayo Clinic College of Medicine, Rochester, MN

I. Abstract II. Introduction III. Molecular Structures, Synthesis, Mechanism of Action of Major Sex Steroids, and Transcriptional Coregulator Function IV. Effects of Sex Steroids on Bone Cells and Bone Turnover

V. Effects of Estrogens and Androgens on Bone Metabolism in Men Versus Women VI. Effects of Sex Steriods on Extraskeletal Calcium Homeostasis VII. Summary References

I. ABSTRACT

significant effects on the male skeleton, estrogen may also play a key role in males, and conversely, androgens have important skeletal effects in women. These dual roles for estrogen and androgens have long been known in reproductive tissues, and now also appear to be true for the skeleton. Sex steroids also have important, indirect effects on bone metabolism via their effects on intestinal calcium absorption and renal calcium handling. While much has been learned in recent years on sex steroid effects on bone, this is a rapidly evolving area and future studies are likely to reveal new insights, both into the mechanisms of sex steroid action on bone, as well as the relative contributions of estrogens versus androgens towards bone metabolism in both sexes.

Sex steroids have major effects on bone and calcium metabolism. Thus, hypogonadism in either sex is associated with rapid bone loss, which is preventable with gonadal steroid replacement. In this chapter, we review the molecular structures, biosynthesis, and mechanism of action of the major clinically relevant sex steroids. The identification of estrogen and androgen receptors in bone cells was followed by an explosion of studies on the effects of sex steroids, and in particular, estrogen, on osteoblasts and osteoclasts. This has led to the identification of candidate autocrine and paracrine mediators of estrogen and androgen action in bone, although important gaps remain in our understanding of the direct effects of sex steroids on bone cell function. Recent genetic, epidemiologic, and direct interventional evidence has also challenged traditional notions of the relative importance of estrogen on bone metabolism in men. While androgens clearly have Dynamics of Bone and Cartilage Metabolism

II. INTRODUCTION Estrogens and androgens play major roles in skeletal homeostasis. Estrogen deficiency following the menopause 327

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is recognized as the major cause of postmenopausal osteoporosis. The effects of estrogen on bone and overall calcium homeostasis have been the subject of intensive investigation. Androgens also have important effects on bone, although the relative contributions of estrogens and androgens towards bone metabolism in both sexes are still under investigation. In this chapter, we review the molecular structures, synthesis, and role of the major sex steroids in bone and calcium metabolism.

III. MOLECULAR STRUCTURES, SYNTHESIS, MECHANISM OF ACTION OF MAJOR SEX STEROIDS, AND TRANSCRIPTIONAL COREGULATOR FUNCTION A. Sex Steroid Structure and Synthesis Figure 1 provides an overview of the major steps in the biosynthetic pathway for the sex steroids [1, 2]. In the ovary, estrone and estradiol are formed from androstenedione and

testosterone, respectively. This reaction is mediated by the enzyme, aromatase, which is a cytochrome P450 enzyme (P450 aromatase) present in the ovary [3]. This enzyme is also present in numerous other locations, including the testis, adipose tissue, and bone cells (osteoblasts and osteoclasts) [4, 5]. During ovulation and pregnancy, progesterone is secreted by the ovaries, although its role in bone metabolism is not as well defined as that of estrogens or androgens. Individual cell types in these tissues are often responsible for the synthesis of each of the species of steroids. The steroidogenic pathway is essentially similar in the testes, with the exception that testosterone is the major secretory product, although there is some conversion of both androstenedione and testosterone to estrone and 17β-estradiol, respectively (Fig. 1). In androgen target tissues, which include bone [6], testosterone is further converted to dihydrotestosterone (DHT). Unlike testosterone, DHT cannot be aromatized to estrogens. The adrenal glands are a significant source of weak androgens, principally dehydroepiandrosterone (DHEA) and androstenedione (Fig. 1). DHEA is further converted by the enzyme steroid sulfotransferase to dehydroepiandrosterone

Figure 1 Biosynthetic pathways involved in the formation of estrogens, androgens, and progestins. (Adapted from Monroe et al. [10].)

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sulfate (DHEAS). DHEA and DHEAS represent the major circulating adrenal androgens. In peripheral tissues, these can be further converted to the more potent sex steroids, testosterone and 17β-estradiol, although there is some evidence that adrenal androgens may have independent and direct effects on some tissues, including bone [7]. In the circulation, the major sex steroids, testosterone and estradiol, are bound to serum-binding proteins, and only 1–3% of the total circulating sex steroids are free in solution. Approximately 35–55% of the sex steroids are bound to albumin, which constitutes a high-capacity, lowaffinity reservoir of the steroids. Both the free fraction and the albumin-bound fraction have access to target tissues [8] and thus represent the “bioavailable” sex steroid. Since sex hormone-binding globulin (SHBG) has a high affinity for testosterone and estradiol, the fraction of these sex steroids bound to SHBG is likely unavailable to target tissues. In contrast to testosterone and estradiol, adrenal androgens do not bind SHBG to a significant degree, and more than 90% of adrenal androgens circulate loosely bound to albumin [9].

B. Mechanisms of Action of Steroids Considerable work has defined the mechanisms of action of sex steroids in target tissues (for reviews, see references [1, 10–12]). Sex steroids rapidly diffuse into cells through the plasma membrane where they bind their cognate receptors, which are proteins of high molecular weight present in low concentrations (approximately 5000–40 000 molecules per cell), primarily in the nucleus. The steroid hormones bind their specific receptor and modify the tertiary structure of the receptor producing

Figure 2

a novel conformation. The result of this conformational change is dissociation from specific chaperone proteins [13, 14], homo- or heterodimerization [15–17], several post-translational modifications including phosphorylation [18–20], and recruitment of specific transcriptional coregulator protein complexes [21]. In the classical pathway, the liganded ER then binds estrogen response elements (EREs) and this leads to the activation of specific genes (Fig. 2A). However, the ER is also known to function through pathways that do not involve direct binding to DNA, such as through protein–protein interactions (Fig. 2B) [22]. In addition, recent evidence has implicated estrogen signaling through a membrane receptor, with subsequent activation of specific cellular kinase pathways [23] (Fig. 2C); the latter appear to be directly involved in estrogen regulation of cellular apoptosis [24, 25]. Two highly related isoforms of the estrogen receptor exist (termed ERα and ERβ) that are encoded by unique genes. Both estrogen receptor isoforms bind 17β-estradiol with similar affinities and utilize similar DNA response elements, whereas androgens bind to the androgen receptor and use their own unique DNA response elements. Interestingly, while almost all other ligand-dependent hormone receptors have two isoforms of the receptor (i.e. ER, GR, PR), only one known androgen receptor isoform exists. The ligand-dependent steroid hormone receptors, including the estrogen and androgen receptors, are classified as type I nuclear hormone receptors. Type I receptors are soluble and recognize their respective DNA-binding elements only when activated by ligand. Type II receptors include receptors that bind nonsteroidal compounds such as thyroid hormone (thyroid receptor; TR), retinoic acid (RAR, RXR), and vitamin D (VDR). These receptors are

Classical and nonclassical pathways of estrogen action. See text for details. (Adapted from Manolagas et al. [188].)

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bound to DNA in the absence of ligand where they function to repress gene transcription. Activation of transcription is elicited through ligand binding. In this chapter, we will concentrate on the type I receptors (specifically estrogen and androgen receptors). All type I receptors share a similar domain structure [26]. While the DNA-binding domains of ERα and ERβ are similar, their transactivation domains, tissue distribution, and molecular sizes differ [21]. The N-terminal region of the receptor shows less conservation of sequence among the steroid hormone receptors and is involved in transcriptional activation functions. The ERβ species is highly expressed in prostate, ovary, brain, and bladder, but is found in limiting quantities in the uterus, kidney, pituitary, and epididymis (these tissues contain high levels of the ERα species [27]). Recent studies also indicate that bone cells express both ERα and ERβ [28, 29]. Several studies are now indicating that coexpression of ERα and ERβ, leading to heterodimerization, has unique functions apart from either ER isoform alone. Finally, a number of studies using mice with deletion of either ERα or ERβ indicate that ERα is likely the major ER species mediating estrogen effects on bone [30], with ERβ either substituting for ERα [31] or perhaps enhancing ERα action under certain conditions [30].

Figure 3

C. Transcriptional Coregulator Function Nuclear hormone receptor coactivators are involved in enhancing the ligand-dependent transcriptional signal of nuclear hormone receptors (see Fig. 3). SRC1, the founding member of the SRC family, was originally identified in a yeast two-hybrid screen of a human B-lymphocyte cDNA library using the ligand-binding domain of progesterone receptor (PR) as bait [32]. Further analysis demonstrated that SRC1 exhibits coactivation properties, not only with PR but also with numerous other nuclear hormone receptors including ER [21, 33]. Two additional members of the SRC family, SRC2 [34] and SRC3 [35], were also identified which interact with and coactivate a wide variety of nuclear hormone receptors including ER. Gene deletion experiments in mice demonstrate that, while homozygous deletion of SRC1 and SRC2 results in partial hormone resistance [36], the phenotype of SRC1 and SRC2 knockout animals otherwise appear largely normal. However, recent studies using SRC1 knockout mice have demonstrated that loss of SRC1 leads to impaired estrogen action, principally in trabecular bone [37]. Interestingly, the SRC3 mutant demonstrates dwarfism and a more severe reproductive phenotype [38]. Numerous other transcriptional

General pathway for E2 and SERM action. The ligand enters the cell through diffusion where it encounters inactive ER monomers. Homo- or heterodimerization can occur with ERα and ERβ. Depending on the type of estrogen response element present at any given promoter, the ER dimer binds either directly or can bind indirectly through protein tethering (AP1 or fos/jun complex, in this case) and causing activation of gene transcription.

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coregulators involved in either activation or repression of nuclear receptor function have been described elsewhere [21, 39]. The SRC family of proteins mediate the interactions with nuclear receptors through a centrally located receptorinteracting domain (RID), which contains three LXXLL motifs necessary for interaction with nuclear receptors [40, 41]. Binding of ligand to ER induces specific conformational changes, providing a unique surface for association of SRCs through the RID. Although all the SRCs are capable of binding ER, it is unclear whether the different SRCs preferentially interact with a particular ER dimer (e.g. ERα, ERβ, or ERα/β). A recent study suggests that SRC1 overexpression preferentially enhances ERβ transcriptional activation of a transiently transfected ERE construct, whereas SRC2 overexpression preferentially enhances ERα transcriptional activation in human bone cells [42]. However, whether these preferences are due to preferential binding or another phenomenon (e.g. posttranslational modifications) is unknown.

IV. EFFECTS OF SEX STEROIDS ON BONE CELLS AND BONE TURNOVER A. Estrogen Estrogen deficiency is recognized as the most relevant factor in the pathogenesis of postmenopausal bone loss [43, 44]. Recent studies have begun to elucidate the molecular mediators of estrogen deficiency on bone resorption. The identification of both ERα and ERβ on human osteoblasts [45] (Fig. 4) and osteoclasts [46–49] has demonstrated that estrogen likely has direct effects on bone, as opposed to more indirect effects of systemic

Figure 4 Demonstration of the presence of ERs (left) and ARs (right) in human bone cells. (Adapted from Colvard et al. [115].)

estrogen deficiency. The modulation of ER-dependent processes within both osteoblasts and osteoclasts has been utilized clinically through estrogen replacement therapy, which has been shown in numerous studies to be effective in preventing and treating osteoporosis [50]. A summary of the effects of estrogen on osteoblasts and osteoclasts is described in Figure 5. The OPG/RANKL/RANK system has become recognized as one of the major regulatory mechanisms influencing osteoclastogenesis [51–54] (see Fig. 5). Osteoblasts express surface RANKL that binds to surface RANK receptors on osteoclast precursor cells, leading to differentiation and activation of these precursors into mature osteoclasts. A soluble decoy receptor, osteoprotegerin (OPG), is secreted from the preosteoblast/stromal cell that inhibits osteoclastogenesis by competing with RANKL for RANK. Estrogen can exert its antiresorptive effect by up-regulating OPG [55–58], thus inhibiting bone resorption. Estrogen can also decrease the expression of RANKL on the surface of osteoblasts leading to a similar inhibition of bone resorption [59]. Estrogens also have direct effects on the responsiveness and formation of osteoclasts [60, 61], thereby decreasing bone resorption. In addition to the effects of estrogen on the OPG and RANKL expression, estrogen also regulates the production of additional cytokines in osteoblasts or bone marrow mononuclear cells, thus modulating osteoclastic activity in a paracrine fashion [62]. There is now an increasing body of evidence that bone-resorbing cytokines, such as interleukin (IL)-1, IL-6, tumor necrosis factor α (TNF-α),

Figure 5 osteocytes.

Summary of estrogen effects on osteoblasts, osteoclasts, and

332 macrophage-colony stimulating factor (M-CSF), and prostaglandins may be potential candidates for mediating the bone loss following estrogen deficiency. Both IL-1 and M-CSF production are increased in estrogen-deficient model systems [63, 64], which can be inhibited using specific antagonists [65–67]. Additionally, the boneresorptive effects of TNF-α are well documented [68] and can be reversed using a soluble type I TNF receptor [69]. Numerous other studies indicate that IL-6 plays a key role in mediating bone loss following estrogen deficiency [70–74]. However, it is likely that, in vivo, multiple cytokines act cooperatively in inducing bone resorption following estrogen deficiency, and that a single cytokine may only partially account for the effects of estrogen deficiency on the skeleton. In contrast to the inhibitory effects of estrogen on bone resorption, the effects of estrogen on bone formation and on osteoblast proliferation and differentiation are less clear. Since estrogen deficiency is associated with both increased bone resorption and impaired compensatory bone formation [75], it is likely that estrogen has significant effects on bone formation. In support of this, estrogen has been shown to increase production of IGF-I [76] and transforming growth factor-β [77] by osteoblastic cells in vitro. Also, estrogen has been reported to stimulate fibroblast [78] and osteoblast [76] type I collagen synthesis, which represents >90% of the biosynthetic capacity of these cells. However, estrogen treatment has been reported both to stimulate [79] and to inhibit [80] bone formation in experimental animals, so this issue is far from resolved. In vitro, estrogen effects on osteoblast proliferation and differentiation markers have been variable, depending on the model system used [76, 81, 82]. Since these initial studies were conducted before ERβ was discovered, many of these apparent discrepancies may be due to the ER isoform expressed and the receptor concentration, which can greatly influence both the magnitude and direction of the estrogen-dependent response. A number of more recent studies, using osteoblastic cell systems expressing ERα, have found a consistent inhibitory effect of estrogen on proliferation [83–85], whereas no effect of estrogen was seen using cells expressing ERβ [83]. Additionally, Waters et al. [86] found that osteoblastic matrix formation, mineralization, and the expression and activity of bone marker genes are not only dependent on the ER isoform, but also on the stage of osteoblastic differentiation (e.g. matrix synthesis, early/late mineralization stages). Interestingly, Cao et al. [87] found that in the MG-63 osteosarcoma line that ERβ mediates the synthesis of bone matrix proteins. Collectively, these data suggest that estrogen directly regulates osteoblast proliferation and differentiation, although the net effect of estrogen on

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osteoblasts likely depends on factors such as species differences, cell system heterogeneity, differentiation stage, ER isoform expression, and receptor concentration. The advent of microarray technology has advanced the understanding of osteoblastic gene expression. A number of studies using an osteoblastic cell model expressing either ERα or ERβ has demonstrated that estrogen differentially regulates a unique subset of genes through either ER isoform [83, 88, 89] (see Fig. 6). These unique estrogendependent regulatory patterns are most likely due to differences in recruitment of transcriptional coregulator molecules. Indeed, as noted earlier, recent studies have indicated that ERα and ERβ have preferential affinities for the coregulators SRC1 and SRC2 in transient transfection assays in the hFOB osteoblast model [42]. Recent evidence also suggests that, in addition to the osteoblast, the osteocyte may also be a target for estrogen action. Estrogen withdrawal associated with GnRH therapy has, for example, been shown to induce apoptosis of osteocytes in iliac bone [90]. Furthermore, estrogen treatment inhibits osteocyte apoptosis induced by pro-apoptotic stimuli [23]. Since osteocytes may be involved in mechanosensing and transducing loading responses [91], these effects of estrogen deficiency could impair the skeletal response to loading. In addition to effects on bone resorption and possibly on bone formation in the adult skeleton, estrogen clearly also has major effects on skeletal development. Thus, the increase in estrogen levels at the onset of puberty in girls is associated with the pubertal increase in growth velocity as well as the ultimate closure of the epiphyseal growth plate and cessation of linear growth [92]. The pubertal growth spurt in girls is also associated with an increase in bone mass through a combination of increases in bone length, bone diameter, cortical bone width, and cancellous bone mass.

Figure 6 Microarrray analysis of U2OS cells stably expressing either ERα or ERβ (U2OS-ERα, U2OS-ERβ) treated with 10 nM E2 for 24 h. The data demonstrate that ~80% of genes regulated by either ER isoform are regulated uniquely (e.g. not regulated by the other ER isoform). Data taken from [83].

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The mechanism of action of estrogen on growth plate cartilage is at present poorly understood. It appears, however, that estrogen stimulates maturation of cartilage without increasing the growth rate [93, 94]. In addition, the effects of estrogen on cartilage growth and maturation are likely mediated by interactions with growth hormone and insulin-like growth factor-I. Thus, although estrogen stimulates growth hormone secretion, it also inhibits IGF-I synthesis in the growth plate [95]. Therefore, the longitudinal growth rate may be influenced by the relative actions of estrogen and growth hormone on the growth plate. Finally, studies on selective estrogen receptor modulators (SERMs), such as raloxifene, are providing new insights into possible alternate pathways for estrogen action in bone. These compounds have tissue-specific actions and can serve as either estrogen antagonists or agonists. For example, they can affect the skeleton without stimulating breast or endometrial tissue [96]. Although they bind the ER, they induce conformational changes in the ER that are different from estradiol that affects transcriptional coregulator interactions [21, 97–99]. The SERM–ER complex may modulate the expression of genes via alternate, nonclassical pathways, rather than activating estrogen-responsive genes via the classical steroid pathway. For example, estrogen regulates the expression of the IL-6 gene, not by direct interaction of the estrogen/ER complex with DNA, but by binding and inhibiting the activity of transcription factors, such as NF-κβ and C/EBPβ, which are required for IL-6 gene transcription [70, 100, 101] (Fig. 2). It is possible that SERMs may also activate this pathway in bone cells. Similarly, raloxifene has been shown to activate the gene for TGF-β3 in human osteosarcoma MG63 cells transfected with an ERα lacking the DBD. This effect may be mediated by a raloxifene response element in DNA that is distinct from the estrogen response element [102, 103]. Both ERα and ERβ interact with AP-1 sites (fos/jun) [104]; however, the ER isoforms have different affinities and transcriptional activation potentials for various SERMs [105, 106]. When estradiol is present, ERα activates transcription from the AP-1 site, whereas ERβ inhibits transcription from this site. In contrast, in the presence of the SERM raloxifene, ERβ stimulates transcription from the AP-1 site, whereas ERα inhibits it [104]. Further elucidation of the mechanism of action of these SERMs is likely to provide new insights into estrogen action in bone and in other tissues.

B. Progesterone The effects of progesterone on bone metabolism are much less clear than those of estrogen. However, progesterone

receptors have been identified in primary human osteoblastic cells, human osteosarcoma cells, and immortalized fetal osteoblast cells [107, 108], although estrogen is generally required to induce progesterone receptors in these cells, as in other cell systems [109, 110]. In addition, progesterone has been shown to increase the proliferation and differentiation of human osteoblastic cells [111]. Although progesterone can stimulate transcriptional activation of bone markers such as type I collagen, osteocalcin [112] and other bone-related cytokines such as IGF-II [111], transforming growth factor TGFβ1–3 [113], progesterone does not appear to have consistent effects on postmenopausal bone loss in humans [112, 114]. Furthermore, while estrogen treatment of primary human osteoblasts stimulates OPG mRNA production [55–58], progesterone had no significant effect on OPG expression [57]. Thus, the effects of progesterone on the human skeleton likely differ significantly from those of estrogen. Indeed, since progesterone is a known antagonist of estrogen in certain cell processes, and an agonist in others, it is possible that the major role of progesterone is to modulate estrogen action in the processes of bone metabolism.

C. Androgens Human bone cells have been demonstrated to have specific androgen-binding sites [115], and the androgen receptor concentrations found have been similar to estrogen receptor concentrations in these cells (Fig. 4). Moreover, the number of specific androgen-binding sites in osteoblasts (1000–3000 sites/cell) is similar to the number of binding sites present in other androgen-responsive tissues, such as the prostate [116]. In addition to the androgen receptor, bone cells have also been shown to have 5α-reductase activity [117], which is responsible for the conversion of testosterone to DHT, the major biologically active metabolite of testosterone in most tissues. Testosterone may also be aromatized to estrogens in various tissues by the microsomal enzyme, aromatase, and there is also evidence that bone cells possess an aromatase activity [118]. As in the case of estrogen, there is evidence that androgens inhibit bone resorption. Thus, orchiectomy, like ovariectomy, is associated with increased bone resorption and rapid bone loss [119]. In addition, Tenover [120] has shown that 3 months of testosterone treatment of men with borderline serum testosterone levels resulted in a 28% reduction in urinary hydroxyproline excretion, an index of bone resorption. In another study of eugonadal men with osteoporosis, who were treated with 6 months of intramuscular testosterone, testosterone therapy was associated

334 with significant reductions in bone resorption markers [121]. Finally, Riggs et al. [122] performed bone biopsies in 29 women with postmenopausal osteoporosis before and after either estrogen or oxandrolone (a synthetic androgen) treatment and found similar decreases in resorption surfaces, and comparable changes in bone resorption rate assessed by calcium kinetics. Androgen receptors have also been identified on osteoclasts, and androgens, like estrogen, have been shown to directly decrease bone resorption in vitro by avian and human osteoclast-like cells [123]. In addition to these direct effects on osteoclasts, androgens have also been shown to regulate the production of a number of bone-resorbing factors by osteoblasts or marrow stromal cells. Thus, both DHT and testosterone have been shown to reduce PGE2 production in calvarial organ cultures exposed to stimulation with parathyroid hormone or IL-1 [124]. Similarly, androgens have been shown to inhibit IL-6 production by stromal and osteoblastic cells as well as the stimulation of osteoclastogenesis by marrow osteoclast precursors [125, 126]. While androgens have generally been considered anabolic for bone, the evidence for this in adults is relatively sparse. Thus, in the studies noted above, Tenover [120] found that testosterone treatment had no effect on the bone formation marker, serum osteocalcin. In contrast, Raisz et al. [127] found that postmenopausal women treated with estrogen plus 2.5 mg of methyltestosterone had a 24% higher serum osteocalcin level after 3 months of treatment, as compared to a 40% lower osteocalcin level in the women treated with estrogen alone. In addition, Baran et al. [128] performed bone biopsies in a hypogonadal man before and after 6 months of testosterone treatment and noted increases in relative osteoid volume, total osteoid surface, linear extent of bone formation, and bone mineralization. Similarly, Cantatore et al. [129] administered synthetic androgens in postmenopausal women and found increases in makers of bone formation, at least over the short term (3–6 months), and Morley et al. [130] also noted that 3 months of testosterone therapy in elderly hypogonadal men resulted in a significant increase in serum osteocalcin levels. Thus, at least over the short term, androgens may have an anabolic effect on bone, although further studies are needed to address this issue. As in the case of estrogen, in vitro studies of androgen effects on osteoblast proliferation and differentiation have yielded inconsistent results. Androgens have been shown to have mitogenic effects on normal and transformed osteoblast-like cells in most [81, 131], but not all systems [132, 133]. With respect to osteoblast differentiation, androgens have been shown to increase alkaline phosphatase activity in primary cultures of osteoblast-like cells and the

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percentage of alkaline phosphatase-positive cells, suggesting a shift towards a more differentiated phenotype [131, 134, 135]. On the other hand, studies using a fetal human osteoblast cell line stably transfected with the androgen receptor have found a decrease in alkaline phosphatase activity in these cells following DHT exposure [133]. Androgen effects on type I collagen synthesis have been variable, with some studies showing a stimulatory effect [81, 132, 134], and others finding no effect [124], or even a decrease [133] in collagen synthesis. Finally, studies using primary osteoblast cultures have suggested that some of the effects of androgens in these cells may be mediated, at least in part, by increased TGF-β production, or by alterations in the insulin-like growth factor/insulin-like growth factor-binding protein system [136, 137]. In addition to testosterone and DHT, there is some evidence that adrenal androgens may have significant effects on bone metabolism. Thus, Durbridge et al. [138] found that, in rats, adrenalectomy alone resulted in loss of metaphyseal trabecular bone of an extent similar to that produced by oophorectomy. These investigators interpreted these data as being consistent with an important role for adrenal androgens in the maintenance of skeletal mass in rats. In addition, Turner et al. [139] have shown that, in rats, treatment with DHEA reduced the loss of cancellous bone following oophorectomy, indicating that adrenal androgens may prevent the bone loss induced by estrogen deficiency. DHEA, which is essentially a prohormone, can be converted locally in various tissues to androstenedione and then into potent androgens and/or estrogens, and it has generally been assumed that the effects of adrenal androgens on target tissues are due to the effects of these androgens or estrogens [140]. However, the recent description of specific binding sites for DHEA [141] raises the possibility that DHEA or similar compounds may have direct effects on bone. In support of this, Bodine et al. [7] have shown that DHEA caused a rapid reduction (30 minutes post treatment) in steady-state levels of c-fos mRNA in normal human osteoblastic cells, whereas testosterone or DHT had no effects on c-fos expression. However, all three agents resulted in significant increases in TGF-β activity in these cells. These studies demonstrate that bone cells have the ability to respond to DHEA, and that the effects of DHEA may or may not be identical to the effects of testosterone, depending on the specific metabolism and perhaps independent action of DHEA in bone cells. Perhaps the major role of androgens in bone metabolism occurs at puberty. As in females, adolescence in males is associated with rapid longitudinal growth, and marked increases in axial and appendicular bone mass [142]. The increase in bone mass is associated with increases in markers of bone formation and is closely linked to pubertal

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stage and to testosterone levels [92, 143], suggesting that testicular androgen production plays a major role in mediating the pubertal increase in bone mass. In addition, however, the increase in adrenal androgens that occurs before the onset of puberty also likely affects the acquisition of bone mass. Thus, longitudinal growth rate increases during adrenarche [144] and bone mass has been shown to increase before the onset of sexual development, likely related to the rise in adrenal androgens [145]. Androgens are also clearly responsible for the sexual dimorphism of the skeleton that is evident during adolescence: the male skeleton is larger in most dimensions compared to the female skeleton, the diameter and cortical thickness of long bones is greater in men than in women, and vertebral size is larger in men [146]. Despite these unequivocal effects of androgens on skeletal growth, however, it remains unresolved which of these effects are due to direct androgen effects on bone and which are due to indirect effects. For example, like estrogen, testosterone can influence growth hormone secretion [147] or the local production of IGF-I or its binding proteins [148], and thus affect bone mass indirectly. It is likely, therefore, that androgen effects on bone during adolescence are due both to direct and indirect effects.

V. EFFECTS OF ESTROGENS AND ANDROGENS ON BONE METABOLISM IN MEN VERSUS WOMEN Based on the sexual dimorphism of the skeleton, it had generally been believed that estrogens were the major determinant of bone metabolism in women and androgens primarily affected bone metabolism in men. Recent evidence, however, indicates considerable overlap of the effects of sex steroids on bone in men and women. Data from several “experiments of nature” are consistent, for example, with the concept that estrogen plays a major role in bone metabolism in men. Smith et al. [149] described a male with homozygous mutations in the ERα gene who, even in the presence of normal testosterone levels, had unfused epiphyses and marked osteopenia, along with elevated indices of bone turnover. Subsequently, Morishima et al. [150] and Carani et al. [151] reported clinical findings in two males with homozygous mutations in the aromatase gene. In both instances, bone mineral density was significantly reduced and bone turnover markers were markedly elevated despite normal testosterone levels. Treatment with testosterone did not significantly affect bone metabolism in one patient, whereas treatment with estrogen markedly increased bone mineral density in both

335 patients [151, 152]. Moreover, a number of observational studies [153–157] have demonstrated that serum estrogen levels (particularly bioavailable estrogen levels) are better predictors of bone density in men at all measured sites except at some cortical bone sites in the appendicular skeleton. In addition, rates of bone loss in elderly men are also associated with serum estrogen levels [158–160]. Direct evidence for the key role played by estrogen in bone metabolism in men was provided by Falahati-Nini et al. [161] who suppressed endogenous testosterone and estrogen production in a group (n = 59) of older men (mean age, 68 years) using a combination of a GnRH agonist and an aromatase inhibitor. Physiologic estradiol and testosterone levels were maintained by placing the men on the respective patches. After baseline measurements of bone turnover markers, the men were then randomized to one of four groups in order to rigorously delineate the relative contributions of estrogen and testosterone towards regulating bone resorption and formation. Group A (−T, −E) had both patches withdrawn; Group B (−T, +E) had the testosterone patch withdrawn but continued the estrogen patch; Group C (+T, −E) had the estrogen patch withdrawn, but continued the testosterone patch; and Group D (+T, +E) continued both patches. All subjects were continued on GnRH and the aromatase inhibitor. Figure 7A shows the changes in the bone resorption markers, urine deoxypyridinoline (Dpd) and N-telopeptide of type I collagen (NTx), after the subjects had been on the variable treatments for a period of 3 weeks. As is evident, significant increases in both urinary Dpd and NTx excretion, Group A (−T, −E), were prevented completely by continuing T and E replacement [Group D (+T, +E)]. E alone (Group B) was almost completely able to prevent the increase in bone resorption, whereas T alone (Group C) was much less effective. Using a two-factor ANOVA model, the effects of E on urinary Dpd and NTx excretion were highly significant (P = 0.005 and 0.0002, respectively). E accounted for 70% or more of the total effect of sex steroids on bone resorption in these older men, while T could account for no more than 30% of the effect. Using a somewhat different design, Leder et al. [162] have confirmed an independent effect of T on bone resorption, although the data in the aggregate clearly favor a more prominent effect of E on the control of bone resorption in men. Figure 7B shows the corresponding changes in the bone formation markers, serum osteocalcin, and PINP. The reductions in both osteocalcin and PINP levels with the induction of sex steroid deficiency (Group A) were prevented with continued E and T replacement (Group D). Interestingly, serum osteocalcin, which is a marker of function of the mature osteoblast and osteocyte, was maintained

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Figure 7 Percent changes in (A) bone resorption markers (urinary deoxypyridinoline [Dpd] and N-telopeptide of type I collagen [NTx]) and (B) bone formation markers (serum osteocalcin and N-terminal extension peptide of type I collagen [PINP]) in a group of elderly men (mean age 68 years) made acutely hypogonadal and treated with an aromatase inhibitor (Group A), treated with E alone (Group B), T alone (Group C), or both (Group D). See text for details. Asterisks indicate significance for change from baseline: *, P < 0.05; **, P < 0.01; ***, P < 0.001. The estrogen and testosterone effects were analyzed using two-factor ANOVA models: Dpd: E effect, P = 0.005; T effect, P = 0.232; NTx: E effect, P = 0.0002; T effect, P = 0.085; Osteocalcin: E effect, P = 0.002; T effect, P = 0.013; PINP: E effect, P = 0.0001; T effect, P = 0.452. (Adapted from Falahati-Nini et al. [161].)

by either E or T (ANOVA P values of 0.002 and 0.013, respectively). By contrast, serum PINP, which represents type I collagen synthesis throughout the various stages of osteoblast differentiation, was maintained by E (ANOVA P value 0.0001), but not T. Collectively, these findings provided conclusive proof of an important (and indeed, dominant) role for E in bone metabolism in the mature skeleton of adult men. Similar findings were subsequently reported by Taxel et al. [163] in a study of 15 elderly men treated with an aromatase inhibitor for 9 weeks, where suppression of E production resulted in significant increases in bone resorption markers and a suppression of bone formation markers. Similarly, several lines of evidence indicate that androgens may have important effects on bone in women. In premenopausal women, serum androgen levels are significantly related to bone mineral density [155, 164], suggesting that androgens may contribute to the development of peak bone mass in women. Moreover, hyperandrogenic women with hirsutism have increased bone mass relative to control women [165]. Finally, testosterone levels correlate

with bone mass and rates of fall in bone mass in perimenopausal women [164].

VI. EFFECTS OF SEX STEROIDS ON EXTRASKELETAL CALCIUM HOMEOSTASIS In addition to direct effects on bone cells and on bone remodeling, it is becoming clear that sex steroids may affect bone turnover indirectly through an action on extraskeletal calcium homeostasis, including effects on intestinal calcium absorption, renal calcium handling, and perhaps direct effects on PTH secretion. Thus, Gallagher et al. [166] found that estrogen treatment increased both serum total 1,25(OH)2D levels and calcium absorption in postmenopausal osteoporotic women, although this effect may have been due to the increase in serum PTH in response to the estrogen-induced decrease in bone resorption. However, in perimenopausal women before and

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6 months after oophorectomy, Gennari et al. [167] found that the increase in calcium absorption in response to treatment with 1,25(OH)2D was blunted in the presence of estrogen deficiency, suggesting a direct effect of estrogen on intestinal calcium absorption. Of note, estrogen receptors have been found in an untransformed cell line from rat small intestinal crypts (IEC-6 cells) as well as epithelial cells from rat intestine using both classical receptor-binding techniques and RT-PCR [168]. Thus, estrogen may affect bone turnover both directly by acting on bone cells and indirectly by altering intestinal calcium absorption and hence serum PTH levels. Estrogen receptors have also been demonstrated in renal tubules [169], and estrogen has been reported to increase adenylate cyclase activity of renal membranes from hens [170] and from cultured opossum kidney cells [171]. Using estimates based on regression analysis, Nordin et al. [172] found that early postmenopausal women had a “renal calcium leak” that they attributed to estrogen deficiency. McKane et al. [173] assessed renal calcium transport by direct measurements both at baseline and during administration of a saturating dose of PTH in early postmenopausal women before and after 6 months of estrogen treatment. They demonstrated a PTH-independent decrease in tubular calcium reabsorption in the estrogendeficient women compared to the estrogen-replete women, consistent with a direct effect of estrogen on renal calcium conservation. In addition to the indirect effects on serum PTH levels, there is evidence that estrogen may directly regulate PTH secretion. Some [174], but not other [175], investigators have demonstrated the presence of estrogen receptors in parathyroid glands. Cosman et al. [176] reported that chronic estrogen replacement therapy in elderly women decreased maximal PTH secretory rate, as assessed by measurement of serum PTH during EDTA-induced hypocalcemia, although Vincent et al. [177] failed to demonstrate an effect of acute estrogen treatment or withdrawal on PTH secretion. Studies in experimental animals and parathyroid cells in vitro suggest that estrogen increases PTH secretion [178]. Thus, estrogen administration to oophorectomized rats increased PTH mRNA in the parathyroid glands four-fold [174], and in vitro estrogen treatment of surgically removed human hyperplastic parathyroid glands increased PTH secretion in a dose- and time-dependent manner [177]. Some of the contrasting effect of estrogen on PTH secretion may be related to acute versus chronic effects, although more work is needed to address this issue. There are several lines of evidence suggesting that androgens may also have similar, indirect effects on bone metabolism. Thus, there are data indicating that androgens

may significantly enhance intestinal calcium absorption. Hope et al. [179] reported that duodenal active calcium transport decreased in male rats following orchiectomy and testosterone treatment reversed this. In early studies, Lafferty et al. [180] found that calcium absorption increased following short-term (2–3 months) testosterone treatment in three subjects studied, although these changes did not persist with long-term therapy. Need et al. [181] studied 27 women with postmenopausal osteoporosis and found a significant increase in the hourly fractional rate of radiocalcium absorption from (mean ± SEM) 0.79 ± 0.06 to 0.93 ± 0.05 following 3 months of therapy with the anabolic steroid, nandrolone decanoate. Some evidence also suggests that androgens may affect renal calcium transport. The androgen receptor gene is expressed in kidney epithelial cells [182] and androgens have been shown to modulate calcium fluxes in mouse kidney cortex preparations [183]. Reifenstein and Albright [184] initially noted that testosterone propionate decreased the urinary excretion of calcium. Several studies using synthetic androgens have also suggested an effect of androgens on renal calcium handling. Thus, Chesnut et al. [185] noted a 32% decrease in urinary calcium excretion in 23 women with postmenopausal osteoporosis following 8–32 months of therapy with the anabolic steroid, stanozolol. Similarly, Need et al. [181] indirectly estimated renal tubular calcium reabsorption and found that this increased significantly in 27 women with postmenopausal osteoporosis treated for 3 months with nandrolone decanoate. Finally, in recent studies, Mason et al. [186] showed that DHT administration increased the tubular reabsorption of calcium in oophorectomized rats. Thus, both estrogen and androgens may affect bone metabolism indirectly via alterations in overall calcium homeostasis and subsequent changes in serum PTH levels. Indeed, loss of these indirect effects of sex steroids in elderly women and in some men may account, in large part, for the age-related increase in serum PTH levels in both sexes [187]. This, in turn, results in an age-related increase in bone turnover and, in the setting of a concomitant defect in osteoblast function, in bone loss.

VII. SUMMARY Sex steroids clearly have major effects on all aspects of bone metabolism. These include effects on longitudinal growth at the time of puberty, the acquisition of peak bone mass, and the regulation of bone turnover in adults. While there are several candidate factors for mediating the autocrine and paracrine effects of estrogen and, to a lessor extent, androgens, on osteoclasts and osteoblasts, the precise

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mechanism of direct sex steroid effects on the skeleton remains to be clearly defined. Estrogen likely has a significant effect on bone not just in females, but also in males. Conversely, androgens also have important skeletal effects in both sexes. In addition, sex steroids also affect bone turnover indirectly, via extraskeletal effects on calcium homeostasis. Collectively, these direct and indirect actions of sex steroids on bone and calcium metabolism are critical for normal bone mass acquisition and maintenance.

Acknowledgment Supported by Research Grants AG-004875, AR-027065 and AR-30582 from the National Institutes of Health, U.S. Public Health Service.

References 1. Monroe, D. G., and Spelsberg, T. (2004). General mechanisms of steroid receptors and receptor coregulator action. In: “Advances in Molecular and Cell Biology” (V. Miller, M. Hay, eds.), 34th ed., Vol. 34, pp. 39–48. Elsevier B.V., Amsterdam. 2. Hsueh, A. J. W., and Billig, H. (1995). Ovarian hormone synthesis and mechanism of action. In: “Endocrinology” (L. J. DeGroot, M. Besser, H. G. Burger, J. L. Jameon, D. L. Loriaux, J. C. Marshall, W. D. Odell, J. T. Potts, A. H. Rubenstein, eds.), pp. 2019–2030. W.B. Saunders, Philadelphia, PA. 3. Simpson, E. R., Mahendroo, M. S., Means, G. D., Kilgore, M. W., Hinshelwood, M. M., Graham-Lorence, S., Amarneh, B., Ito, Y., Fisher, C. R., Michael, M. D., Mendelson, C. R., and Bulun, S. E. (1994). Aromatase cytochrome P450, the enzyme responsible for estrogen biosynthesis. Endocr. Rev. 15(3), 342–355. 4. Simpson, E. R. (2003). Sources of estrogen and their importance. J. Steroid Biochem. Mol. Biol. 86(3–5), 225–230. 5. Nawata, H., Tanaka, S., Takayanagi, R., Sakai, Y., Yanase, T., Ikuyama, S., and Haji, M. (1995). Aromatase in bone cell: association with osteoporosis in postmenopausal women. J. Steroid Biochem. Mol. Biol. 53(1–6), 165–174. 6. Dinneen, S., Alzaid, A., Miles, J., and Rizza, R. (1993). Metabolic effects of the nocturnal rise in cortisol on carbohydrate metabolism in normal humans. J. Clin. Invest. 92(5), 2283–2290. 7. Bodine, P. V., Riggs, B. L., and Spelsberg, T. C. (1995). Regulation of c-fos expression and TGF-beta production by gonadal and adrenal androgens in normal human osteoblastic cells. J. Steroid Biochem. Mol. Biol. 52(2), 149–158. 8. Manni, A., Pardridge, W. M., Cefalu, W., Nisula, B. C., Bardin, C. W., Santner, S. J., and Santen, R. J. (1985). Bioavailability of albumin-bound testosterone. J. Clin. Endocrinol. Metab. 61(4), 705–710. 9. Cekan, S. Z., Xing, S., and Ritzen, M. (1984). On the binding of steroid sulfates to albumin. Experientia 40(9), 949–951. 10. Monroe, D. G., and Spelsberg, T. C. (2004). Gonadal steroids and receptors. In: “Primer on the Metabolic Bone Diseases and Disorders of Mineral Metabolism” (M. J. Favus, ed), 5th ed., pp. 32–38. Lippincott, Williams and Wilkins, Philadelphia, PA. 11. Monroe, D. G., and Spelsberg, T. C. (2003). A case for estrogen receptors on cell membranes and nongenomic actions of estrogen. Calcif. Tissue Int. 72(3), 183–184. 12. Monroe, D. G., Secreto, F. J., and Spelsberg, T. (2003). Overview of estrogen action in osteoblasts: Role of the ligand, receptor, and the coregulators. J. Musculoskeletal Neuronal. Int. 3(4), 357–362.

13. Pratt, W. B., and Toft, D. O. (2003). Regulation of signaling protein function and trafficking by the hsp90/hsp70-based chaperone machinery. Exp. Biol. Med. (Maywood) 228(2), 111–133. 14. Smith, D. F., and Toft, D. O. (1993). Steroid receptors and their associated proteins. Mol. Endocrinol. 7(1), 4–11. 15. Cowley, S. M., Hoare, S., Mosselman, S., and Parker, M. G. (1997). Estrogen receptors alpha and beta form heterodimers on DNA. J. Biol. Chem. 272(32), 19858–19862. 16. Jisa, E., and Jungbauer, A. (2003). Kinetic analysis of estrogen receptor homo- and heterodimerization in vitro. J. Steroid Biochem. Mol. Biol. 84(2–3), 141–148. 17. Tsai, S. Y., Carlstedt-Duke, J., Weigel, N. L., Dahlman, K., Gustafsson, J. A., Tsai, M. J., and O’Malley, B. W. (1988). Molecular interactions of steroid hormone receptor with its enhancer element: evidence for receptor dimer formation. Cell 55(2), 361–369. 18. Driggers, P. H., and Segars, J. H. (2002). Estrogen action and cytoplasmic signaling pathways. Part II: the role of growth factors and phosphorylation in estrogen signaling. Trends Endocrinol. Metab. 13(10), 422–427. 19. Lannigan, D. A. (2003). Estrogen receptor phosphorylation. Steroids 68(1), 1–9. 20. Takimoto, G. S., Tasset, D. M., Eppert, A. C., and Horwitz, K. B. (1992). Hormone-induced progesterone receptor phosphorylation consists of sequential DNA-independent and DNA-dependent stages: analysis with zinc finger mutants and the progesterone antagonist ZK98299. Proc. Natl Acad. Sci. USA 89(7), 3050–3054. 21. Smith, C. L., and O’Malley, B. W. (2004). Coregulator function: a key to understanding tissue specificity of selective receptor modulators. Endocr. Rev. 25(1), 45–71. 22. McDonnell, D. P., and Norris, J. D. (1997). Analysis of the molecular pharmacology of estrogen receptor agonists and antagonists provides insights into the mechanism of action of estrogen in bone. Osteoporos. Int. 7 Suppl 1, S29–34. 23. Kousteni, S., Bellido, T., Plotkin, L. I., O’Brien, C. A., Bodenner, D. L., Han, L., Han, K., DiGregorio, G. B., Katzenellenbogen, J. A., Katzenellenbogen, B. S., Roberson, P. K., Weinstein, R. S., Jilka, R. L., and Manolagas, S. C. (2001). Nongenotropic, sex-nonspecific signaling through the estrogen or androgen receptors: dissociation from transcriptional activity. Cell 104(5), 719–730. 24. Kousteni, S., Chen, J. R., Bellido, T., Han, L., Ali, A. A., O’Brien, C. A., Plotkin, L., Fu, Q., Mancino, A. T., Wen, Y., Vertino, A. M., Powers, C. C., Stewart, S. A., Ebert, R., Parfitt, A. M., Weinstein, R. S., Jilka, R. L., and Manolagas, S. C. (2002). Reversal of bone loss in mice by nongenotropic signaling of sex steroids. Science 298(5594), 843–846. 25. Kousteni, S., Han, L., Chen, J. R., Almeida, M., Plotkin, L. I., Bellido, T., and Manolagas, S. C. (2003). Kinase-mediated regulation of common transcription factors accounts for the bone-protective effects of sex steroids. J. Clin. Invest. 111(11), 1651–1664. 26. Parl, F. F. (2000). Estrogen Receptor and Breast Cancer. IOS Press, Amsterdam. 27. Kuiper, G. G., Enmark, E., Pelto-Huikko, M., Nilsson, S., and Gustafsson, J. A. (1996). Cloning of a novel receptor expressed in rat prostate and ovary. Proc. Natl Acad. Sci. USA 93(12), 5925–5930. 28. Arts, J., Kuiper, G. G., Janssen, J. M., Gustafsson, J. A., Lowik, C. W., Pols, H. A., van Leeuwen, J. P. (1997). Differential expression of estrogen receptors alpha and beta mRNA during differentiation of human osteoblast SV-HFO cells. Endocrinology 138(11), 5067–5070. 29. Onoe, Y., Miyaura, C., Ohta, H., Nozawa, S., and Suda, T. (1997). Expression of estrogen receptor beta in rat bone. Endocrinology 138(10), 4509–4512. 30. Sims, N. A., Clement-Lacroix, P., Minet, D., Fraslon-Vanhulle, C., Gaillard-Kelly, M., Resche-Rigon, M., and Baron, R. (2003). A functional androgen receptor is not sufficient to allow estradiol to protect bone after gonadectomy in estradiol receptor-deficient mice. J. Clin. Invest. 111(9), 1319–1327. 31. Sims, N. A., Dupont, S., Krust, A., Clement-Lacroix, P., Minet, D., Resche-Rigon, M., Gaillard-Kelly, M., and Baron, R. (2002). Deletion of

Chapter 19 Sex Steroid Effects on Bone Metabolism

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estrogen receptors reveals a regulatory role for estrogen receptors-beta in bone remodeling in females but not in males. Bone 30(1), 18–25. Onate, S. A., Tsai, S. Y., Tsai, M. J., and O’Malley, B. W. (1995). Sequence and characterization of a coactivator for the steroid hormone receptor superfamily. Science 270(5240), 1354-1357. McKenna, N. J., Lanz, R. B., and O’Malley, B. W. (1999). Nuclear receptor coregulators: cellular and molecular biology. Endocr. Rev. 20(3), 321–344. Voegel, J. J., Heine, M. J., Zechel, C., Chambon, P., and Gronemeyer, H. (1996). TIF2, a 160 kDa transcriptional mediator for the ligand-dependent activation function AF-2 of nuclear receptors. Embo. J. 15(14), 3667–3675. Li, H., Gomes, P. J., and Chen, J. D. (1997). RAC3, a steroid/nuclear receptor-associated coactivator that is related to SRC-1 and TIF2. Proc. Natl Acad. Sci. USA 94(16), 8479–8484. Xu, J., Qiu, Y., DeMayo, F. J., Tsai, S. Y., Tsai, M. J., and O’Malley, B. W. (1998). Partial hormone resistance in mice with disruption of the steroid receptor coactivator-1 (SRC-1) gene. Science 279(5358), 1922–1925. Modder, U. I., Sanyal, A., Kearns, A. E., Sibonga, J. D., Nishihara, E., Xu, J., O’Malley, B. W., Ritman, E. L., Riggs, B. L., Spelsberg, T. C., and Khosla, S. (2004). Effects of loss of steroid receptor coactivator-1 on the skeletal response to estrogen in mice. Endocrinology 145(2), 913–921. Xu, J., Liao, L., Ning, G., Yoshida-Komiya, H., Deng, C., and O’Malley, B. W. (2000). The steroid receptor coactivator SRC-3 (p/CIP/RAC3/AIB1/ACTR/TRAM-1) is required for normal growth, puberty, female reproductive function, and mammary gland development. Proc. Natl Acad. Sci. USA 97(12), 6379–6384. Martini, P. G., and Katzenellenbogen, B. S. (2003). Modulation of estrogen receptor activity by selective coregulators. J. Steroid Biochem. Mol. Biol. 85(2–5), 117–122. Heery, D. M., Kalkhoven, E., Hoare, S., and Parker, M. G. (1997). A signature motif in transcriptional co-activators mediates binding to nuclear receptors. Nature 387(6634), 733–736. Leo, C., and Chen, J. D. (2000). The SRC family of nuclear receptor coactivators. Gene 245(1), 1–11. Monroe, D. G., Johnsen, S. A., Subramaniam, M., Getz, B. J., Khosla, S., Riggs, B. L., and Spelsberg, T. C. (2003). Mutual antagonism of estrogen receptors alpha and beta and their preferred interactions with steroid receptor coactivators in human osteoblastic cell lines. J. Endocrinol. 176(3), 349–357. Riggs, B. L., Khosla, S., and Melton, L. J., III (2002). Sex steroids and the construction and conservation of the adult skeleton. Endocr. Rev. 23(3), 279–302. Riggs, B. L., Wahner, H. W., Melton, L. J., III, Richelson, L. S., Judd, H. L., and Offord, K. P. (1986). Rates of bone loss in the appendicular and axial skeletons of women. Evidence of substantial vertebral bone loss before menopause. J. Clin. Invest. 77(5), 1487–1491. Eriksen, E. F., Colvard, D. S., Berg, N. J., Graham, M. L., Mann, K. G., Spelsberg, T. C., and Riggs, B. L. (1988). Evidence of estrogen receptors in normal human osteoblast-like cells. Science 241(4861), 84–86. Gyda, M., Corisdeo, S., Zaidi, M., and Troen, B. R. (2001). Macrophage colony-stimulating factor suppresses osteoblast formation. Biochem. Biophys. Res. Commun. 285(2), 328–334. Hoyland, J. A., Mee, A. P., Baird, P., Braidman, I. P., Mawer, E. B., and Freemont, A. J. (1997). Demonstration of estrogen receptor mRNA in bone using in situ reverse-transcriptase polymerase chain reaction. Bone 20(2), 87–92. Vidal, O., Kindblom, L. G., and Ohlsson, C. (1999). Expression and localization of estrogen receptor-beta in murine and human bone. J. Bone Miner. Res. 14(6), 923–929. Oursler, M. J., Pederson, L., Fitzpatrick, L., Riggs, B. L., and Spelsberg, T. (1994). Human giant cell tumors of the bone (osteoclastomas) are estrogen target cells. Proc. Natl Acad. Sci. USA 91(12), 5227–5231.

339 50. Riggs, B. L., and Melton, L. J., III (1992). The prevention and treatment of osteoporosis [published erratum appears in N. Engl. J. Med. 1993 Jan 7; 328(1):65]. N. Engl. J. Med. 327, 620–627. 51. Khosla, S. (2001). Minireview: the OPG/RANKL/RANK system. Endocrinology 142(12), 5050–5055. 52. Hofbauer, L. C., Khosla, S., Dunstan, C. R., Lacey, D. L., Boyle, W. J., and Riggs, B. L. (2000). The roles of osteoprotegerin and osteoprotegerin ligand in the paracrine regulation of bone resorption. J. Bone Miner. Res. 15(1), 2–12. 53. Troen, B. R. (2003). Molecular mechanisms underlying osteoclast formation and activation. Exp. Gerontol. 38(6), 605–614. 54. Bell, N. H. (2001). Advances in the treatment of osteoporosis. Curr. Drug Targets Immune Endocr. Metabol. Disord. 1(1), 93–102. 55. Bord, S., Ireland, D. C., Beavan, S. R., and Compston, J. E. (2003). The effects of estrogen on osteoprotegerin, RANKL, and estrogen receptor expression in human osteoblasts. Bone 32(2), 136–141. 56. Hofbauer, L. C., Khosla, S., Dunstan, C. R., Lacey, D. L., Spelsberg, T. C., and Riggs, B. L. (1999). Estrogen stimulates gene expression and protein production of osteoprotegerin in human osteoblastic cells. Endocrinology 140(9), 4367–4370. 57. Liao, E. Y., Luo, X. H., and Su, X. (2002). Comparison of the effects of 17β-E2 and progesterone on the expression of osteoprotegerin in normal human osteoblast-like cells. J. Endocrinol. Invest. 25(9), 785–790. 58. Saika, M., Inoue, D., Kido, S., and Matsumoto, T. (2001). 17β-estradiol stimulates expression of osteoprotegerin by a mouse stromal cell line, ST-2, via estrogen receptor-alpha. Endocrinology 142(6), 2205–2212. 59. Eghbali-Fatourechi, G., Khosla, S., Sanyal, A., Boyle, W. J., Lacey, D. L., and Riggs, B. L. (2003). Role of RANK ligand in mediating increased bone resorption in early postmenopausal women. J. Clin. Invest. 111(8), 1221–1230. 60. Srivastava, S., Toraldo, G., Weitzmann, M. N., Cenci, S., Ross, F. P., and Pacifici, R. (2001). Estrogen decreases osteoclast formation by down-regulating receptor activator of NF-κB ligand (RANKL)induced JNK activation. J. Biol. Chem. 276(12), 8836–8840. 61. Kanatani, M., Sugimoto, T., Takahashi, Y., Kaji, H., Kitazawa, R., and Chihara, K. (1998). Estrogen via the estrogen receptor blocks cAMPmediated parathyroid hormone (PTH)-stimulated osteoclast formation. J. Bone Miner. Res. 13(5), 854–862. 62. Manolagas, S. C., and Jilka, R. L. (1995). Bone marrow, cytokines, and bone remodeling. Emerging insights into the pathophysiology of osteoporosis. N. Engl. J. Med. 332(5), 305–311. 63. Pacifici, R., Brown, C., Puscheck, E., Friedrich, E., Slatopolsky, E., Maggio, D., McCracken, R., and Avioli, L. V. (1991). Effect of surgical menopause and estrogen replacement on cytokine release from human blood mononuclear cells. Proc. Natl Acad. Sci. USA 88(12), 5134–5138. 64. Tanaka, S., Takahashi, N., Udagawa, N., Tamura, T., Akatsu, T., Stanley, E. R., Kurokawa, T., and Suda, T. (1993). Macrophage colonystimulating factor is indispensable for both proliferation and differentiation of osteoclast progenitors. J. Clin. Invest. 91(1), 257–263. 65. Kimble, R. B., Vannice, J. L., Bloedow, D. C., Thompson, R. C., Hopfer, W., Kung, V. T., Brownfield, C., and Pacifici, R. (1994). Interleukin-1 receptor antagonist decreases bone loss and bone resorption in ovariectomized rats. J. Clin. Invest. 93(5), 1959–1967. 66. Ammann, P., Rizzoli, R., Bonjour, J. P., Bourrin, S., Meyer, J. M., Vassalli, P., and Garcia, I. (1997). Transgenic mice expressing soluble tumor necrosis factor-receptor are protected against bone loss caused by estrogen deficiency. J. Clin. Invest. 99(7), 1699–1703. 67. Kimble, R. B., Srivastava, S., Ross, F. P., Matayoshi, A., and Pacifici, R. (1996). Estrogen deficiency increases the ability of stromal cells to support murine osteoclastogenesis via an interleukin-1and tumor necrosis factor-mediated stimulation of macrophage colony-stimulating factor production. J. Biol. Chem. 271(46), 28890–28897. 68. Nanes, M. S. (2003). Tumor necrosis factor-alpha: molecular and cellular mechanisms in skeletal pathology. Gene 321, 1–15.

340 69. Kitazawa, R., Kimble, R. B., Vannice, J. L., Kung, V. T., and Pacifici, R. (1994). Interleukin-1 receptor antagonist and tumor necrosis factor binding protein decrease osteoclast formation and bone resorption in ovariectomized mice. J. Clin. Invest. 94(6), 2397–2406. 70. Manolagas, S. C., Kousteni, S., and Jilka, R. L. (2002). Sex steroids and bone. Recent Prog. Horm. Res. 57, 385–409. 71. Girasole, G., Jilka, R. L., Passeri, G., Boswell, S., Boder, G., Williams, D. C., and Manolagas, S. C. (1992). 17β-estradiol inhibits interleukin-6 production by bone marrow-derived stromal cells and osteoblasts in vitro: a potential mechanism for the antiosteoporotic effect of estrogens. J. Clin. Invest. 89(3), 883–891. 72. Jilka, R. L., Hangoc, G., Girasole, G., Passeri, G., Williams, D. C., Abrams, J. S., Boyce, B., Broxmeyer, H., and Manolagas, S. C. (1992). Increased osteoclast development after estrogen loss: mediation by interleukin-6. Science 257(5066), 88–91. 73. Kassem, M., Harris, S. A., Spelsberg, T. C., and Riggs, B. L. (1996). Estrogen inhibits interleukin-6 production and gene expression in a human osteoblastic cell line with high levels of estrogen receptors. J. Bone Miner. Res. 11(2), 193–199. 74. Poli, V., Balena, R., Fattori, E., Markatos, A., Yamamoto, M., Tanaka, H., Ciliberto, G., Rodan, G. A., and Costantini, F. (1994). Interleukin-6 deficient mice are protected from bone loss caused by estrogen depletion. Embo. J. 13(5), 1189–1196. 75. Ivey, J. L., and Baylink, D. J. (1981). Postmenopausal osteoporosis: proposed roles of defective coupling and estrogen deficiency. Metab. Bone Dis. Relat. Res. 3(1), 3–7. 76. Ernst, M., Heath, J. K., and Rodan, G. A. (1989). Estradiol effects on proliferation, messenger ribonucleic acid for collagen and insulin-like growth factor-I, and parathyroid hormone-stimulated adenylate cyclase activity in osteoblastic cells from calvariae and long bones. Endocrinology 125(2), 825–833. 77. Oursler, M. J., Cortese, C., Keeting, P., Anderson, M. A., Bonde, S. K., Riggs, B. L., and Spelsberg, T. C. (1991). Modulation of transforming growth factor-beta production in normal human osteoblast-like cells by 17β-estradiol and parathyroid hormone. Endocrinology 129(6), 3313–3320. 78. Brincat, M., Moniz, C. F., Kabalan, S., Versi, E., O’Dowd, T., Magos, A. L., Montgomery, J., and Studd, J. W. (1987). Decline in skin collagen content and metacarpal index after the menopause and its prevention with sex hormone replacement. Br. J. Obstet. Gynaecol. 94(2), 126–129. 79. Kawaguchi, H., Pilbeam, C. C., Vargas, S. J., Morse, E. E., Lorenzo, J. A., and Raisz, L. G. (1995). Ovariectomy enhances and estrogen replacement inhibits the activity of bone marrow factors that stimulate prostaglandin production in cultured mouse calvariae. J. Clin. Invest. 96(1), 539–548. 80. Westerlind, K. C., Wakley, G. K., Evans, G. L., and Turner, R. T. (1993). Estrogen does not increase bone formation in growing rats. Endocrinology 133(6), 2924–2934. 81. Gray, C., Colston, K. W., Mackay, A. G., Taylor, M. L., and Arnett, T. R. (1992). Interaction of androgen and 1,25-dihydroxyvitamin D3: effects on normal rat bone cells. J. Bone Miner. Res. 7(1), 41–46. 82. Gray, T. K., Mohan, S., Linkhart, T. A., and Baylink, D. J. (1989). Estradiol stimulates in vitro the secretion of insulin-like growth factors by the clonal osteoblastic cell line, UMR106. Biochem. Biophys. Res. Commun. 158(2), 407–412. 83. Monroe, D. G., Getz, B. J., Johnsen, S. A., Riggs, B. L., Khosla, S., and Spelsberg, T. C. (2003). Estrogen receptor isoform-specific regulation of endogenous gene expression in human osteoblastic cell lines expressing either ERα or ERβ. J. Cell. Biochem. 90(2), 315–326. 84. Kassem, M., Okazaki, R., De Leon, D., Harris, S. A., Robinson, J. A., Spelsberg, T. C., Conover, C. A., and Riggs, B. L. (1996). Potential mechanism of estrogen-mediated decrease in bone formation: estrogen increases production of inhibitory insulin-like growth factor-binding protein-4. Proc. Assoc. Am. Physicians 108(2), 155–164.

DAVID G. MONROE ET AL. 85. Robinson, J. A., Harris, S. A., Riggs, B. L., and Spelsberg, T. C. (1997). Estrogen regulation of human osteoblastic cell proliferation and differentiation. Endocrinology 138(7), 2919–2927. 86. Waters, K. M., Rickard, D. J., Riggs, B. L., Khosla, S., Katzenellenbogen, J. A., Katzenellenbogen, B. S., Moore, J., and Spelsberg, T. C. (2001). Estrogen regulation of human osteoblast function is determined by the stage of differentiation and the estrogen receptor isoform. J. Cell. Biochem. 83(3), 448–462. 87. Cao, L., Bu, R., Oakley, J. I., Kalla, S. E., and Blair, H. C. (2003). Estrogen receptor-β modulates synthesis of bone matrix proteins in human osteoblast-like MG63 cells. J. Cell. Biochem. 89(1), 152–164. 88. Kian Tee, M., Rogatsky, I., Tzagarakis-Foster, C., Cvoro, A., An, J., Christy, R. J., Yamamoto, K. R., and Leitman, D. C. (2004). Estradiol and selective estrogen receptor modulators differentially regulate target genes with estrogen receptors alpha and beta. Mol. Biol. Cell 15(3), 1262–1272. 89. Stossi, F., Barnett, D. H., Frasor, J., Komm, B., Lyttle, C. R., and Katzenellenbogen, B. S. (2004). Transcriptional profiling of estrogen-regulated gene expression via estrogen receptor (alpha) or estrogen receptor (beta) in human osteosarcoma cells: Distinct and common target genes for these receptors. Endocrinology 145(7), 3473–3486. 90. Tomkinson, A., Reeve, J., Shaw, R. W., and Noble, B. S. (1997). The death of osteocytes via apoptosis accompanies estrogen withdrawal in human bone. J. Clin. Endocrinol. Metab. 82(9), 3128–3135. 91. Pitsillides, A. A., Rawlinson, S. C., Suswillo, R. F., Bourrin, S., Zaman, G., and Lanyon, L. E. (1995). Mechanical strain-induced NO production by bone cells: a possible role in adaptive bone (re)modeling? Faseb. J. 9(15), 1614–1622. 92. Bonjour, J. P., Theintz, G., Buchs, B., Slosman, D., and Rizzoli, R. (1991). Critical years and stages of puberty for spinal and femoral bone mass accumulation during adolescence. J. Clin. Endocrinol. Metab. 73(3), 555–563. 93. Gustafsson, P. O., Kasstrom, H., Lindberg, L., and Olsson, S. E. (1975). Growth and mitotic rate of the proximal tibial epiphyseal plate in hypophysectomized rats given estradiol and human growth hormone. Acta Radiol. Suppl. 344, 69–74. 94. Strickland, A. L., and Sprinz, H. (1973). Studies of the influence of estradiol and growth hormone on the hypophysectomized immature rat epiphyseal cartilage growth plate. Am. J. Obstet. Gynecol. 115(4), 471–477. 95. Ho, K. K., and Weissberger, A. J. (1992). Impact of short-term estrogen administration on growth hormone secretion and action: distinct route-dependent effects on connective and bone tissue metabolism. J. Bone Miner. Res. 7(7), 821–827. 96. Delmas, P. D., Bjarnason, N. H., Mitlak, B. H., Ravoux, A. C., Shah, A. S., Huster, W. J., Draper, M., and Christiansen, C. (1997). Effects of raloxifene on bone mineral density, serum cholesterol concentrations, and uterine endometrium in postmenopausal women. N. Engl. J. Med. 337(23), 1641–1647. 97. Brzozowski, A. M., Pike, A. C., Dauter, Z., Hubbard, R. E., Bonn, T., Engstrom, O., Ohman, L., Greene, G. L., Gustafsson, J. A., and Carlquist, M. (1997). Molecular basis of agonism and antagonism in the oestrogen receptor. Nature 389(6652), 753–758. 98. Tamrazi, A., Carlson, K. E., and Katzenellenbogen, J. A. (2003). Molecular sensors of estrogen receptor conformations and dynamics. Mol. Endocrinol. 17(12), 2593–2602. 99. Heldring, N., Nilsson, M., Buehrer, B., Treuter, E., and Gustafsson, J. A. (2004). Identification of tamoxifen-induced coregulator interaction surfaces within the ligand-binding domain of estrogen receptors. Mol. Cell. Biol. 24(8), 3445–3459. 100. Ray, A., Prefontaine, K. E., and Ray, P. (1994). Down-regulation of the interleukin-6 promoter by estrogen is mediated by NF-κB and C/EBP. J. Biol. Chem. 269, 12940–12946.

Chapter 19 Sex Steroid Effects on Bone Metabolism 101. Stein, B., and Yang, M. X. (1995). Repression of the interleukin-6 promoter by estrogen receptor is mediated by NF-kappa B and C/EBP beta. Mol. Cell. Biol. 15(9), 4971–4979. 102. Yang, N. N., Venugopalan, M., Hardikar, S., and Glasebrook, A. (1996). Identification of an estrogen response element activated by metabolites of 17β-estradiol and raloxifene. Science 273(5279), 1222–1225. 103. Yang, N. N., Bryant, H. U., Hardikar, S., Sato, M., Galvin, R. J., Glasebrook, A. L., and Termine, J. D. (1996). Estrogen and raloxifene stimulate transforming growth factor-β3 gene expression in rat bone: a potential mechanism for estrogen- or raloxifene-mediated bone maintenance. Endocrinology 137(5), 2075–2084. 104. Paech, K., Webb, P., Kuiper, G. G., Nilsson, S., Gustafsson, J., Kushner, P. J., and Scanlan, T. S. (1997). Differential ligand activation of estrogen receptors ERα and ERβ at AP1 sites. Science 277(5331), 1508–1510. 105. Kuiper, G. G., and Gustafsson, J. A. (1997). The novel estrogen receptor-beta subtype: potential role in the cell- and promoter-specific actions of estrogens and anti-estrogens. FEBS Lett. 410(1), 87–90. 106. Kuiper, G. G., Carlsson, B., Grandien, K., Enmark, E., Haggblad, J., Nilsson, S., and Gustafsson, J. A. (1997). Comparison of the ligand binding specificity and transcript tissue distribution of estrogen receptors alpha and beta. Endocrinology 138(3), 863–870. 107. MacNamara, P., O’Shaughnessy, C., Manduca, P., and Loughrey, H. C. (1995). Progesterone receptors are expressed in human osteoblast-like cell lines and in primary human osteoblast cultures. Calcif. Tissue Int. 57(6), 436–441. 108. Wei, L. L., Leach, M. W., Miner, R. S., and Demers, L. M. (1993). Evidence for progesterone receptors in human osteoblast-like cells. Biochem. Biophys. Res. Commun. 195(2), 525–532. 109. Flototto, T., Niederacher, D., Hohmann, D., Heimerzheim, T., Dall, P., Djahansouzi, S., Bender, H. G., and Hanstein, B. (2004). Molecular mechanism of estrogen receptor (ER)alpha-specific, estradioldependent expression of the progesterone receptor (PR) B-isoform. J. Steroid Biochem. Mol. Biol. 88(2), 131–142. 110. Rickard, D. J., Waters, K. M., Ruesink, T. J., Khosla, S., Katzenellenbogen, J. A., Katzenellenbogen, B. S., Riggs, B. L., and Spelsberg, T. C. (2002). Estrogen receptor isoform-specific induction of progesterone receptors in human osteoblasts. J. Bone Miner. Res. 17(4), 580–592. 111. Tremollieres, F. A., Strong, D. D., Baylink, D. J., and Mohan, S. (1992). Progesterone and promegestone stimulate human bone cell proliferation and insulin-like growth factor-2 production. Acta Endocrinol. (Copenh.) 126(4), 329–337. 112. Azizi, G., Hansen, A., and Prestwood, K. M. (2003). Effect of micronized progesterone on bone turnover in postmenopausal women on estrogen replacement therapy. Endocr. Res. 29(2), 133–140. 113. Luo, X. H., Liao, E. Y., and Su, X. (2002). Progesterone upregulates TGF-β isoforms (β1, β2, and β3) expression in normal human osteoblast-like cells. Calcif. Tissue Int. 71(4), 329–334. 114. Prior, J. C., Vigna, Y. M., Wark, J. D., Eyre, D. R., Lentle, B. C., Li, D. K., Ebeling, P. R., and Atley, L. (1997). Premenopausal ovariectomy-related bone loss: a randomized, double-blind, one-year trial of conjugated estrogen or medroxyprogesterone acetate. J. Bone Miner. Res. 12(11), 1851–1863. 115. Colvard, D. S., Eriksen, E. F., Keeting, P. E., Wilson, E. M., Lubahn, D. B., French, F. S., Riggs, B. L., and Spelsberg, T. C. (1989). Identification of androgen receptors in normal human osteoblast-like cells. Proc. Natl Acad. Sci. USA 86(3), 854–857. 116. Sanchez-Visconti, G., Herrero, L., Rabadan, M., Pereira, I., and Ruiz-Torres, A. (1995). Ageing and prostate: age-related changes in androgen receptors of epithelial cells from benign hypertrophic glands compared with cancer. Mech. Ageing Dev. 82(1), 19–29. 117. Schweikert, H. U., Rulf, W., Niederle, N., Schafer, H. E., Keck, E., and Kruck, F. (1980). Testosterone metabolism in human bone. Acta Endocrinol. (Copenh.) 95(2), 258–264.

341 118. Nakano, Y., Morimoto, I., Ishida, O., Fujihira, T., Mizokami, A., Tanimoto, A., Yanagihara, N., Izumi, F., and Eto, S. (1994). The receptor, metabolism and effects of androgen in osteoblastic MC3T3-E1 cells. Bone Miner. 26(3), 245–259. 119. Stepan, J. J., Lachman, M., Zverina, J., Pacovsky, V., and Baylink, D. J. (1989). Castrated men exhibit bone loss: effect of calcitonin treatment on biochemical indices of bone remodeling. J. Clin. Endocrinol. Metab. 69(3), 523–527. 120. Tenover, J. S. (1992). Effects of testosterone supplementation in the aging male. J. Clin. Endocrinol. Metab. 75(4), 1092–1098. 121. Anderson, F. H., Francis, R. M., Peaston, R. T., and Wastell, H. J. (1997). Androgen supplementation in eugonadal men with osteoporosis: effects of six months’ treatment on markers of bone formation and resorption. J. Bone Miner. Res. 12(3), 472–478. 122. Riggs, B. L., Jowsey, J., Goldsmith, R. S., Kelly, P. J., Hoffman, D. L., and Arnaud, C. D. (1972). Short- and long-term effects of estrogen and synthetic anabolic hormone in postmenopausal osteoporosis. J. Clin. Invest. 51(7), 1659–1663. 123. Pederson, L., Kremer, M., Judd, J., Pascoe, D., Spelsberg, T. C., Riggs, B. L., and Oursler, M. J. (1999). Androgens regulate bone resorption activity of isolated osteoclasts in vitro. Proc. Natl Acad. Sci. USA 96(2), 505–510. 124. Pilbeam, C. C., and Raisz, L. G. (1990). Effects of androgens on parathyroid hormone and interleukin-1-stimulated prostaglandin production in cultured neonatal mouse calvariae. J. Bone Miner. Res. 5(11), 1183–1188. 125. Bellido, T., Jilka, R. L., Boyce, B. F., Girasole, G., Broxmeyer, H., Dalrymple, S. A., Murray, R., and Manolagas, S. C. (1995). Regulation of interleukin-6, osteoclastogenesis, and bone mass by androgens. The role of the androgen receptor. J. Clin. Invest. 95(6), 2886–2895. 126. Hofbauer, L. C., Ten, R. M., and Khosla, S. (1999). The anti-androgen hydroxyflutamide and androgens inhibit interleukin-6 production by an androgen-responsive human osteoblastic cell line. J. Bone Miner. Res. 14(8), 1330–1337. 127. Raisz, L. G., Wiita, B., Artis, A., Bowen, A., Schwartz, S., Trahiotis, M., Shoukri, K., and Smith, J. (1996). Comparison of the effects of estrogen alone and estrogen plus androgen on biochemical markers of bone formation and resorption in postmenopausal women. J. Clin. Endocrinol. Metab. 81(1), 37–43. 128. Baran, D. T., Bergfeld, M. A., Teitelbaum, S. L., and Avioli, L. V. (1978). Effect of testosterone therapy on bone formation in an osteoporotic hypogonadal male. Calcif. Tissue Res. 26(2), 103–106. 129. Cantatore, F. P., Carrozzo, M., Magli, D., D’Amore, M., and Pipitone, V. (1988). Serum osteocalcin levels in normal humans of different sex and age. An epidemiological study. Panminerva Med. 30(1), 23–25. 130. Morley, J. E., Perry, H. M., III, Kaiser, F. E., Kraenzle, D., Jensen, J., Houston, K., Mattammal, M., and Perry, H. M., Jr. (1993). Effects of testosterone replacement therapy in old hypogonadal males: a preliminary study. J. Am. Geriatr. Soc. 41(2), 149–152. 131. Kasperk, C. H., Wergedal, J. E., Farley, J. R., Linkhart, T. A., Turner, R. T., and Baylink, D. J. (1989). Androgens directly stimulate proliferation of bone cells in vitro. Endocrinology 124(3), 1576–1578. 132. Benz, D. J., Haussler, M. R., Thomas, M. A., Speelman, B., and Komm, B. S. (1991). High-affinity androgen binding and androgenic regulation of alpha 1(I)-procollagen and transforming growth factor-beta steady state messenger ribonucleic acid levels in human osteoblastlike osteosarcoma cells. Endocrinology 128(6), 2723–2730. 133. Hofbauer, L. C., Hicok, K. C., and Khosla, S. (1998). Effects of gonadal and adrenal androgens in a novel androgen-responsive human osteoblastic cell line. J. Cell. Biochem. 71(1), 96–108. 134. Kasperk, C. H., Faehling, K., Borcsok, I., and Ziegler, R. (1996). Effects of androgens on subpopulations of the human osteosarcoma cell line SaOS2. Calcif. Tissue Int. 58(5), 376–382.

342 135. Kasperk, C. H., Wakley, G. K., Hierl, T., and Ziegler, R. (1997). Gonadal and adrenal androgens are potent regulators of human bone cell metabolism in vitro. J. Bone Miner. Res. 12(3), 464–471. 136. Kasperk, C., Fitzsimmons, R., Strong, D., Mohan, S., Jennings, J., Wergedal, J., and Baylink, D. (1990). Studies of the mechanism by which androgens enhance mitogenesis and differentiation in bone cells. J. Clin. Endocrinol. Metab. 71(5), 1322–1329. 137. Gori, F., Hofbauer, L., Conover, C. A., and Khosla, S. (1999). Effects of androgens on the insulin-like growth factor system in an androgen-responsive human osteoblastic cell line. Endocrinology 140, 5579–5586. 138. Durbridge, T. C., Morris, H. A., Parsons, A. M., Parkinson, I. H., Moore, R. J., Porter, S., Need, A. G., Nordin, B. E., and Vernon-Roberts, B. (1990). Progressive cancellous bone loss in rats after adrenalectomy and oophorectomy. Calcif. Tissue Int. 47(6), 383–387. 139. Turner, R. T., Lifrak, E. T., Beckner, M., Wakley, G. K., Hannon, K. S., and Parker, L. N. (1990). Dehydroepiandrosterone reduces cancellous bone osteopenia in ovariectomized rats. Am. J. Physiol. 258(4 Pt 1), E673–677. 140. Labrie, F., Belanger, A., Simard, J., Van, L.-T., and Labrie, C. (1995). DHEA and peripheral androgen and estrogen formation: intracinology. Ann. N. Y. Acad. Sci. 774, 16–28. 141. Meikle, A. W., Dorchuck, R. W., Araneo, B. A., Stringham, J. D., Evans, T. G., Spruance, S. L., and Daynes, R. A. (1992). The presence of a dehydroepiandrosterone-specific receptor binding complex in murine T cells. J. Steroid Biochem. Mol. Biol. 42(3–4), 293–304. 142. Lu, P. W., Briody, J. N., Ogle, G. D., Morley, K., Humphries, I. R., Allen, J., Howman-Giles, R., Sillence, D., and Cowell, C. T. (1994). Bone mineral density of total body, spine, and femoral neck in children and young adults: a cross-sectional and longitudinal study. J. Bone Miner. Res. 9(9), 1451–1458. 143. Riis, B. J., Krabbe, S., Christiansen, C., Catherwood, B. D., and Deftos, L. J. (1985). Bone turnover in male puberty: a longitudinal study. Calcif. Tissue Int. 37(3), 213–217. 144. Parker, L. N. (1991). Adrenarche. Endocrinol. Metab. Clin. North Am. 20(1), 71–83. 145. Slemenda, C. W., Reister, T. K., Hui, S. L., Miller, J. Z., Christian, J. C., and Johnston, C. C., Jr. (1994). Influences on skeletal mineralization in children and adolescents: evidence for varying effects of sexual maturation and physical activity. J. Pediatr. 125(2), 201–207. 146. Fehily, A. M., Coles, R. J., Evans, W. D., and Elwood, P. C. (1992). Factors affecting bone density in young adults. Am. J. Clin. Nutr. 56(3), 579–586. 147. Mauras, N., Haymond, M. W., Darmaun, D., Vieira, N. E., Abrams, S. A., and Yergey, A. L. (1994). Calcium and protein kinetics in prepubertal boys. Positive effects of testosterone. J. Clin. Invest. 93(3), 1014–1019. 148. Hobbs, C. J., Plymate, S. R., Rosen, C. J., and Adler, R. A. (1993). Testosterone administration increases insulin-like growth factor-I levels in normal men. J. Clin. Endocrinol. Metab. 77(3), 776–779. 149. Smith, E. P., Boyd, J., Frank, G. R., Takahashi, H., Cohen, R. M., Specker, B., Williams, T. C., Lubahn, D. B., and Korach, K. S. (1994). Estrogen resistance caused by a mutation in the estrogen-receptor gene in a man. N. Engl. J. Med. 331(16), 1056–1061. 150. Morishima, A., Grumbach, M. M., Simpson, E. R., Fisher, C., and Qin, K. (1995). Aromatase deficiency in male and female siblings caused by a novel mutation and the physiological role of estrogens. J. Clin. Endocrinol. Metab. 80(12), 3689–3698. 151. Carani, C., Qin, K., Simoni, M., Faustini-Fustini, M., Serpente, S., Boyd, J., Korach, K. S., and Simpson, E. R. (1997). Effect of testosterone and estradiol in a man with aromatase deficiency. N. Engl. J. Med. 337(2), 91–95. 152. Morishima, A., Grumbach, M. M., and Bilezikian, J. P. (1997). Estrogen markedly increases bone mass in an estrogen deficient young man with aromatase deficiency. J. Bone Miner. Res. 12(Suppl 1), S126.

DAVID G. MONROE ET AL. 153. Center, J. R., Nguyen, T. V., White, C. P., and Eisman, J. A. (1997). Male osteoporosis predictors: Sex hormones and calcitropic hormones. J. Bone Miner. Res. 12(Suppl 1), S368. 154. Greendale, G. A., Edelstein, S., and Barrett-Connor, E. (1997). Endogenous sex steroids and bone mineral density in older women and men: the Rancho Bernardo Study. J. Bone Miner. Res. 12(11), 1833–1843. 155. Khosla, S., Melton, L. J., 3rd, Atkinson, E. J., O’Fallon, W. M., Klee, G. G., and Riggs, B. L. (1998). Relationship of serum sex steroid levels and bone turnover markers with bone mineral density in men and women: a key role for bioavailable estrogen. J. Clin. Endocrinol. Metab. 83(7), 2266–2274. 156. Slemenda, C. W., Longcope, C., Zhou, L., Hui, S. L., Peacock, M., and Johnston, C. C. (1997). Sex steroids and bone mass in older men. Positive associations with serum estrogens and negative associations with androgens. J. Clin. Invest. 100(7), 1755–1759. 157. Amin, S., Zhang, Y., Sawin, C. T., Evans, S. R., Hannan, M. T., Kiel, D. P., Wilson, P. W., and Felson, D. T. (2000). Association of hypogonadism and estradiol levels with bone mineral density in elderly men from the Framingham study. Ann. Intern. Med. 133(12), 951–963. 158. Khosla, S., Melton, L. J., III, Atkinson, E. J., and O’Fallon, W. M. (2001). Relationship of serum sex steroid levels to longitudinal changes in bone density in young versus elderly men. J. Clin. Endocrinol. Metab. 86(8), 3555–3561. 159. Gennari, L., Merlotti, D., Martini, G., Gonnelli, S., Franci, B., Campagna, S., Lucani, B., Dal Canto, N., Valenti, R., Gennari, C., and Nuti, R. (2003). Longitudinal association between sex hormone levels, bone loss, and bone turnover in elderly men. J. Clin. Endocrinol. Metab. 88(11), 5327–5333. 160. Van Pottelbergh, I., Goemaere, S., and Kaufman, J. M. (2003). Bioavailable estradiol and an aromatase gene polymorphism are determinants of bone mineral density changes in men over 70 years of age. J. Clin. Endocrinol. Metab. 88(7), 3075–3081. 161. Falahati-Nini, A., Riggs, B. L., Atkinson, E. J., O’Fallon, W. M., Eastell, R., and Khosla, S. (2000). Relative contributions of testosterone and estrogen in regulating bone resorption and formation in normal elderly men. J. Clin. Invest. 106(12), 1553–1560. 162. Leder, B. Z., LeBlanc, K. M., Schoenfeld, D. A., Eastell, R., and Finkelstein, J. S. (2003). Differential effects of androgens and estrogens on bone turnover in normal men. J. Clin. Endocrinol. Metab. 88(1), 204–210. 163. Taxel, P., Kennedy, D. G., Fall, P. M., Willard, A. K., Clive, J. M., and Raisz, L. G. (2001). The effect of aromatase inhibition on sex steroids, gonadotropins, and markers of bone turnover in older men. J. Clin. Endocrinol. Metab. 86(6), 2869–2874. 164. Slemenda, C., Hui, S. L., Longcope, C., and Johnston, C. C. (1987). Sex steroids and bone mass. A study of changes about the time of menopause. J. Clin. Invest. 80(5), 1261–1269. 165. Buchanan, J. R., Hospodar, P., Myers, C., Leuenberger, P., and Demers, L. M. (1988). Effect of excess endogenous androgens on bone density in young women. J. Clin. Endocrinol. Metab. 67(5), 937–943. 166. Gallagher, J. C., Riggs, B. L., and DeLuca, H. F. (1980). Effect of estrogen on calcium absorption and serum vitamin D metabolites in postmenopausal osteoporosis. J. Clin. Endocrinol. Metab. 51(6), 1359–1364. 167. Gennari, C., Agnusdei, D., Nardi, P., and Civitelli, R. (1990). Estrogen preserves a normal intestinal responsiveness to 1,25dihydroxyvitamin D3 in oophorectomized women. J. Clin. Endocrinol. Metab. 71(5), 1288–1293. 168. Thomas, M. L., Xu, X., Norfleet, A. M., and Watson, C. S. (1993). The presence of functional estrogen receptors in intestinal epithelial cells. Endocrinology 132(1), 426–430.

Chapter 19 Sex Steroid Effects on Bone Metabolism 169. Hagenfeldt, Y., and Eriksson, H. A. (1988). The estrogen receptor in the rat kidney. Ontogeny, properties and effects of gonadectomy on its concentration. J. Steroid Biochem. 31(1), 49–56. 170. Forte, L. R., Langeluttig, S. G., Biellier, H. V., Poelling, R. E., Magliola, L., and Thomas, M. L. (1983). Upregulation of kidney adenylate cyclase in the egg-laying hen: role of estrogen. Am. J. Physiol. 245(3), E273–280. 171. Stock, J. L., Coderre, J. A., Burke, E. M., Danner, D. B., Chipman, S. D., and Shapiro, J. R. (1992). Identification of estrogen receptor mRNA and the estrogen modulation of parathyroid hormone-stimulated cyclic AMP accumulation in opossum kidney cells. J. Cell. Physiol. 150(3), 517–525. 172. Nordin, B. E., Need, A. G., Morris, H. A., Horowitz, M., and Robertson, W. G. (1991). Evidence for a renal calcium leak in postmenopausal women. J. Clin. Endocrinol. Metab. 72(2), 401–407. 173. McKane, W. R., Khosla, S., Burritt, M. F., Kao, P. C., Wilson, D. M., Ory, S. J., and Riggs, B. L. (1995). Mechanism of renal calcium conservation with estrogen replacement therapy in women in early postmenopause – a clinical research center study. J. Clin. Endocrinol. Metab. 80(12), 3458–3464. 174. Naveh-Many, T., Almogi, G., Livni, N., and Silver, J. (1992). Estrogen receptors and biologic response in rat parathyroid tissue and C cells. J. Clin. Invest. 90(6), 2434–2438. 175. Prince, R. L., MacLaughlin, D. T., Gaz, R. D., and Neer, R. M. (1991). Lack of evidence for estrogen receptors in human and bovine parathyroid tissue. J. Clin. Endocrinol. Metab. 72(6), 1226–1228. 176. Cosman, F., Nieves, J., Horton, J., Shen, V., and Lindsay, R. (1994). Effects of estrogen on response to edetic acid infusion in postmenopausal osteoporotic women. J. Clin. Endocrinol. Metab. 78(4), 939–943. 177. Vincent, A., Riggs, B. L., Atkinson, E. J., Oberg, A. L., and Khosla, S. (2003). Effect of estrogen replacement therapy on parathyroid hormone secretion in elderly postmenopausal women. Menopause 10, 165–171.

343 178. Duarte, B., Hargis, G. K., and Kukreja, S. C. (1988). Effects of estradiol and progesterone on parathyroid hormone secretion from human parathyroid tissue. J. Clin. Endocrinol. Metab. 66(3), 584–587. 179. Hope, W. G., Ibarra, M. J., and Thomas, M. L. (1992). Testosterone alters duodenal calcium transport and longitudinal bone growth rate in parallel in the male rat. Proc. Soc. Exp. Biol. Med. 200(4), 536–541. 180. Lafferty, F. W., Spencer, G. E., Jr., and Pearson, O. H. (1964). Effects of androgens, estrogens and high calcium intakes on bone formation and resorption in osteoporosis. Am. J. Med. 36, 514–528. 181. Need, A. G., Horowitz, M., Morris, H. A., Walker, C. J., and Nordin, B. E. (1987). Effects of nandrolone therapy on forearm bone mineral content in osteoporosis. Clin. Orthop. 225, 273–278. 182. Stefani, S., Aguiari, G. L., Bozza, A., Maestri, I., Magri, E., Cavazzini, P., Piva, R., and del Senno, L. (1994). Androgen responsiveness and androgen receptor gene expression in human kidney cells in continuous culture. Biochem. Mol. Biol. Int. 32(4), 597–604. 183. Goldstone, A. D., Koenig, H., and Lu, C. Y. (1983). Androgenic stimulation of endocytosis, amino acid and hexose transport in mouse kidney cortex involves increased calcium fluxes. Biochim. Biophys. Acta. 762(2), 366–371. 184. Reifenstein, E. C., and Albright, F. (1946). The metabolic effects of steroid hormones in osteoporosis. J. Clin. Invest. 26, 24–56. 185. Chesnut, C. H., III, Ivey, J. L., Gruber, H. E., Matthews, M., Nelp, W. B., Sisom, K., and Baylink, D. J. (1983). Stanozolol in postmenopausal osteoporosis: therapeutic efficacy and possible mechanisms of action. Metabolism 32(6), 571–580. 186. Mason, R. A., and Morris, H. A. (1997). Effects of dihydrotestosterone on bone biochemical markers in sham and oophorectomized rats. J. Bone Miner. Res. 12(9), 1431–1437. 187. Riggs, B. L., Khosla, S., and Melton, L. J., III (1998). A unitary model for involutional osteoporosis: estrogen deficiency causes both type I and type II osteoporosis in postmenopausal women and contributes to bone loss in aging men. J. Bone Miner. Res. 13(5), 763–773. 188. Manolagas, S. C., and Kousteni, S. (2001). Perspective: nonreproductive sites of action of reproductive hormones. Endocrinology 142(6), 2200–2204.

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Chapter 20

Physiology of Calcium and Phosphate Homeostases René Rizzoli, M.D. and Jean-Philippe Bonjour, M.D.

VII. Calcium and Bone Growth VIII. Body Distribution of Phosphorus IX. Determinants of Extracellular Phosphate Concentration X. Homeostatic Responses to Changes in Phosphate Supply or Demand XI. Conclusions References

I. Abstract II. Introduction III. Body Distribution of Calcium IV. Determinants of Extracellular Calcium Concentration V. Relative Importance of the Various Calcium Fluxes in Controlling Extracellular Calcium Homeostasis VI. Homeostatic Responses to Hypocalcemia

I. ABSTRACT

tubular calcium and phosphate transports, or by releasing calcium from intracellular stores, calcium itself plays the role of an effector on homeostatic mechanisms.

Calcium and phosphate homeostases are controlled by bidirectional calcium and phosphate fluxes, occurring at the levels of intestine, bone, and kidney. The latter organ plays a central role in regulating the extracellular concentration of either ion. Sensitive and efficient regulatory mechanisms, involving extracellular calcium sensing, are triggered by changes in calcium demand or supply. Similarly, the renal handling of phosphate can adjust its capacity to meet the need for phosphate of the organism. Not only calciotropic peptides or steroid hormones are capable of modifying the different calcium and phosphate fluxes to various extents, but also a variety of local factors are implicated in the regulation of calcium and phosphate homeostasis, in order to protect the organism against a deficiency or an overload. Finally, by directly influencing renal Dynamics of Bone and Cartilage Metabolism

Division of Bone Diseases, WHO Collaborating Center for Osteoporosis Prevention, Department of Rehabilitation and Geriatrics, University Hospitals, CH - 1211 Geneva 14 (Switzerland)

II. INTRODUCTION Calcium and phosphate play prominent roles in the regulation of cell function. In addition, both are tightly connected to the process of bone mineralization, in which they participate in the formation of hydroxyapatite crystal deposited in specific regions with the collagen fibril networks. The hydroxyapatite crystal ensures the structural rigidity of the skeleton in its function of standing body support, and in the protection and housing of bone marrow. The regulation of calcium and phosphate homeostases is aimed at maintaining extracellular calcium and phosphate 345

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346 concentration and balance as constant as possible, to protect the organism against deficiency or overload of these ions. Extracellular calcium concentrations are maintained remarkably stable, because of the high sensitivity of a variety of cell systems or organs, including the central nervous system, muscle, and exo-/endocrine glands, to small variations of extracellular calcium concentrations. In contrast, to fulfill the requirements of adequate mineral supply for osteoid tissue mineralization, the level of extracellular inorganic phosphate is adjusted to meet the demand of the organism. The homeostasis of both ions and their balance are controlled by a series of hormones and factors tightly interrelated in complex regulatory systems. The production of some of these agents is regulated by the concentration of the solute they are controlling, through negative feedback mechanisms, justifying thereby the term of calciotropic hormones as far as calcium homeostasis control is concerned.

III. BODY DISTRIBUTION OF CALCIUM Calcium is the fifth inorganic element in abundance in the body. For a 70-kg subject, the calcium mass represents about 1300 grams, 99% of which is in bone and teeth, mostly as hydroxyapatite [1]. A small portion of bone calcium (approximately 1%) is rapidly exchangeable with extracellular fluid, contributing to the regulation of the homeostasis of extracellular calcium concentration, by serving as buffer and storage [2]. During puberty, there is nearly a doubling of body mineral stores through an increase in the size of the skeleton, with minor changes in volumetric bone density, i.e. the amount of bone in bone [3, 4]. By the end of the second decade, most of the body mineral capital is accumulated, though a very few percents of bone consolidation have been suggested to occur during the third decade, particularly in males [5]. Approximately 1% of total body calcium is intracellular. At the intracellular level, calcium homeostasis is controlled by an influx following an electrical and chemical gradient, through selective calcium channels [6–8]. As compared with the 1 mmol/L extracellular calcium concentration, a 10 000-fold lower concentration in the cytosol is maintained by the constant extrusion of calcium, through a calcium– magnesium ATPase and sodium–calcium exchange mechanisms [9, 10]. Intracellular calcium homeostasis is also maintained by a dynamic equilibrium with intracellular mitochondrial and microsomal stores [11, 12]. The concentration of cytosolic-free calcium is around 0.1 µmol/L. This concentration is critical in controlling cell membrane permeability, a large variety of enzymatic reactions, endocrine and exocrine hormone secretions,

RENÉ RIZZOLI AND JEAN-PHILIPPE BONJOUR

and in regulating cardiac, skeletal and smooth muscles contraction (Table I). Calcium is also implicated in the control of cell replication and apoptosis [13, 14]. Regulation of intracellular enzymatic functions is achieved through the interaction with various calcium-binding proteins such as, for instance, calmodulin or troponin, or the actin–myosin system, for muscle contraction [2, 8]. About 0.1% of total body calcium is in the extracellular compartment. Extracellular calcium concentration plays a major role in the integrity and stability of cell membrane, in intercellular adhesion, in blood clotting, and it influences neuromuscular excitability. Plasma calcium is tightly regulated in a narrow range, particularly the ionized form, which amounts to approximately 50% of total plasma calcium, and which represents the physiologically active form, whereas 40% is bound to proteins, mostly albumin, and 10% is under the form of ultrafiltrable ion complexes. The binding equilibrium of calcium with albumin is determined by the pH. Indeed, acute acidosis is associated with a decreased binding and, thereby, a higher proportion of the ionized form. In opposite, acute alkalosis decreases ionized calcium by increasing calcium bound to serum albumin.

IV. DETERMINANTS OF EXTRACELLULAR CALCIUM CONCENTRATION Extracellular calcium concentration is maintained in a dynamic equilibrium through fluxes occurring at the level of the intestine, bone, and kidney (Fig. 1). At steady state, when body ions balance is zero, as in nongrowing individuals, the amounts of solutes entering the extracellular space Table I.

Physiological Roles of Calcium and Phosphate Calcium

Structural Hydroxyapatite (99% body constituent calcium) Exchangeable pool (mineral storage) Function

Intracellular signal transduction Cell adhesion Cell proliferation and differentiation Membrane permeability (neuromuscular excitability, muscle contraction, neurotransmission) Cytoskeleton (cell motility) Exo-/endosecretion Coagulation

Phosphate Hydroxyapatite (85% body phosphorus) Nucleic acids Carbohydrates Lipids Energy storage and delivery Intracellular signal transduction Enzyme activity Acid–base homeostasis

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Figure 1

Main fluxes (mg/day) controlling calcium homeostasis.

are matched by the amounts leaving it. By controlling the calcium output, the kidney plays a central role in the maintenance of calcium homeostasis.

A. Intestinal Flux Net intestinal absorption of calcium represents the difference between the amounts of solutes absorbed and secreted into the gut lumen. In humans, under normal conditions, intestinal absorption of calcium constitutes approximately 20% of ingested calcium. Net intestinal absorption of calcium depends on dietary intakes, on the capacity of the intestinal wall to transport calcium, on the bioavailability of calcium present in the intestinal lumen, and on the secretory flux. Under normal conditions, the secretory flux does not appear to vary markedly. However, it could be increased in pathological conditions such as coeliac disease. The intestinal calcium absorptive capacity is mainly controlled by calcitriol, which stimulates the transport through both genomic and nongenomic mechanisms [15–19]. The interaction of calcitriol with its specific nuclear receptor triggers the synthesis of a variety of proteins, including its receptor itself, calbindins, and the calcium–magnesium ATPase pump, which is located in the enterocyte basolateral membrane. Duodenum possesses the highest concentration of calcitriol receptors, and is the site of the intestine most sensitive to the vitamin metabolite in terms of calcium absorptive capacity. However, the short length of this segment and the rapid transit time suggest that it may not play a quantitative major role in the overall net intestinal calcium absorption. Thus, jejunum and ileum could quantitatively absorb more, despite a less efficient

calcium transport capacity. Parathyroid hormone does not exert any direct effect on the intestinal cells [20, 21]. The importance of bioavailability of calcium at absorptive sites is illustrated by the impairment of calcium absorption induced by the formation of complexes with anions, such as phosphate, sulfate, phytate, or oxalate [22, 23]. For instance, the colonic mucosa is equipped with a powerful vitamin D-sensitive mechanism of calcium transport. However, the absorption is quantitatively little, since calcium is in the large intestine lumen under a form not available to the site of absorption [24]. At steady state, a 24-hour urinary excretion of calcium is mainly the reflection of daily net intestinal calcium absorption. The intestinal absorptive capacity can be evaluated by measuring calcium isotope absorption. A deconvolution analysis of calcium isotopes after oral and intravenous simultaneous administration of 2 different tracers, allows one to determine intestinal calcium transport. A simple evaluation is obtained by measuring the increase in urinary calcium after an oral calcium load [25, 26].

B. Bone Fluxes On average, about 1% of total bone calcium exchanges every month, through a mechanism involving bidirectional fluxes. The main regulators of these fluxes are parathyroid hormone and calcitriol, and possibly calcitonin as an inhibitor of osteoclastic bone resorption under certain conditions [18, 19, 27, 28]. A large variety of substances, either circulating or produced locally, or present in the bone matrix, such as prostaglandins, thyroid hormones, glucocorticoids, sex hormones, growth factors, or interleukins,

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components of the RANK-ligand/osteoprotegerin system, produced by the immune or hematopoietic systems, or bone cells, are capable of influencing bone remodeling, and thereby the bidirectional calcium fluxes [29–32]. In fasting urine, calcium excretion related to creatinine is a direct reflection of net bone [33–35]. Indeed, after an overnight fast, provided there is no calcium supplement taken in the previous evening and the patient does not have a postmicturitional urine residue, calcium appearing in the urine originates mostly from bone. The adjustment by creatinine corrects for urine concentration. Multiplying this ratio by serum creatinine provides urinary calcium excretion per glomerular filtration rate unit.

C. Soft Tissues A minute amount of body calcium is intracellular. Thus, any calcium shift from or into the intracellular compartment does not significantly influence extracellular calcium homeostasis, except maybe in response to parathyroid hormone. The transient decrease in plasma calcium observed in the minutes following parathyroid hormone injection has been attributed to a parathyroid hormonemediated acute transfer of calcium into cells [36, 37].

D. Renal Fluxes Approximately 75% of plasma calcium is ultrafiltrable. After filtration, more than 95% of calcium is reabsorbed. Thus, the amount of calcium appearing in the urine

Figure 2

represents the difference between the amounts filtered and reabsorbed. At steady state, this amount is mainly the reflection of net fluxes into the extracellular fluid of calcium originating from intestine and bone. The proximal tubule reabsorbs 60–70% of calcium, this reabsorption is tightly connected to that of sodium [38] (Fig. 2). Then, 20–30% of filtered calcium is reabsorbed along the loop ascending limb, and 10% at the level of the distal tubule. These two latter reabsorptive sites are influenced by parathyroid hormone. The renal tubular capacity to reabsorb calcium is the main determinant of extracellular calcium concentration. Any change of this capacity is able to induce variations of plasma calcium from 1.5–3.8 mmol/L [33, 34]. This concept has been established by the study of the relationship between urinary calcium excretion and plasma calcium in patients suffering from a lack or an excess of parathyroid hormone. Another example of the central control of extracellular calcium by the renal tubule is given by the syndrome of familial hypocalciuric hypercalcemia, the features of which include an increase in renal tubular reabsorption of calcium independent of parathyroid hormone [39–41]. Various situations or pharmacological agents can modulate the renal tubular reabsorption of calcium. Alkalosis stimulates renal tubular reabsorption of calcium, whereas acidosis decreases it. Thiazides and lithium salts increase the reabsorption of calcium, through mechanisms which are independent of parathyroid hormone [39, 40, 42]. Phosphate deficiency, pharmacological doses of calcitonin and loop diuretics are associated with an increase in calcium clearance [38]. Even large variations of the glomerular filtration rate do not cause major changes in calcemia, since the renal tubule can easily maintain the

Central role of the kidney in controlling calcium homeostasis.

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calcium excretion by modulating its reabsorptive capacity. However, the capacity of the kidney to excrete calcium can be overwhelmed under certain circumstances, particularly in case of large calcium loads resulting from extensive bone destruction and/or very high net calcium intestinal absorption, for instance as a consequence of vitamin D excess, which can lead to an increase in extracellular calcium levels.

V. RELATIVE IMPORTANCE OF THE VARIOUS CALCIUM FLUXES IN CONTROLLING EXTRACELLULAR CALCIUM HOMEOSTASIS An important role in the regulation of calcium fluxes and balance between the various body compartments is played by the above-mentioned systemic hormones. Other hormones such as insulin, growth hormone, insulin-like growth factors, parathyroid hormone-related protein [28, 43–45], glucocorticoids, and sex hormones, as well as locally produced and acting interleukins [46], transforming growth factors and colony-stimulating factors, which are capable of directly influencing calcium metabolism, could also modify the target cell sensitivity to parathyroid hormone and/or calcitriol [29–31, 47–49]. However, their secretion does not appear to be directly controlled by variations in extracellular calcium and/or calcium demand. Any disturbance of the above-described fluxes can result in an alteration of extracellular calcium homeostasis. For instance, an excess of vitamin D is associated with an increase in intestinal absorption of calcium and of bone resorption, and leads to hypercalcemia when the renal excretion capacity is overwhelmed [50, 51]. On the other hand, when renal tubular reabsorption of calcium is stimulated, such as by parathyroid hormone or by parathyroid hormone-related protein, plasma calcium levels can rise, despite very minute changes in calcium influx into the extracellular fluid compartment [33, 34]. Then, despite an increase in renal tubular reabsorption of calcium, urinary excretion is elevated as a consequence of a higher filtered load. The relative and quantitative contribution of calcium mobilization from bone, and of renal tubular reabsorption of calcium, to hypercalcemia induced by parathyroid hormone-related protein, can be estimated in studying the model of thyroparathyroidectomized rats chronically infused with parathyroid hormone-related protein [52]. Thyroparathyroidectomy prevents the contraregulation to variations in extracellular calcium concentrations by endogenous hormones. The elevation of plasma calcium is determined by both increased bone resorption and

enhanced renal tubular reabsorption of calcium. However, the complete inhibition of bone resorption by a bisphosphonate, at a dose which fully normalized fasting urinary calcium excretion, taken as a reflection of net bone resorption, is associated with an approximately 30% decrease, but not a correction of plasma calcium. Thus, the residual hypercalcemia can be attributed to a renal tubular reabsorption effect, which accounted for more than two-thirds of the elevated plasma calcium (Fig. 3A). Indeed, it is well established that bisphosphonates are devoid of any direct effect on the renal handling of calcium [53]. To influence the renal tubular reabsorption, the administration of an agent (the free radical scavenger WR-2721), known to impair the tubular reabsorption of calcium through a parathyroid hormone-independent mechanism [54, 55], is able to acutely increase calcium excretion and to further reduce plasma calcium (Fig. 3B). The predominance of stimulated renal tubular reabsorption of calcium or of increased bone resorption in determining an altered extracellular calcium homeostasis can be demonstrated in a variety of clinical disorders associated with hypercalcemia [50] (Table II, Fig. 4).

VI. HOMEOSTATIC RESPONSES TO HYPOCALCEMIA To maintain the extracellular calcium concentration as constant as possible, the response to acute variations implies changes in the various fluxes, without a necessarily major alteration in total body calcium stores. In contrast, when the organism is chronically submitted to calcium deficiency, its capacity to absorb calcium from the gut or to retain it in the kidney cannot match the need; then, to maintain extracellular calcium concentration, the skeletal mineral is mobilized, leading to a progressive decrease in bone mineral mass [1]. This mechanism might be implicated in the pathogenesis of senile osteoporosis. To recognize changes in extracellular calcium concentration, parathyroid cells have a sensitive calcium sensor, capable of transmitting the information to the parathyroid hormonesynthesizing and -releasing machinery.

A. Extracellular Calcium-sensing Receptor A central role in the regulation of calcium homeostasis is played by parathyroid hormone produced by the parathyroid glands, which recognize alterations in plasma calcium [2, 28, 56]. Any change in plasma calcium is detected by a cell membrane-associated calcium sensor/receptor, which can also be activated by other divalent cations [2, 41, 57].

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Figure 3 Relative contribution of net bone resorption and renal tubular reabsorption to parathyroid hormone-related protein-induced hypercalcemia in thyroparathyroidectomized rats. (A) Pair-fed animals were chronically infused by subcutaneous osmotic minipumps with synthetic parathyroid hormonerelated protein (1–34) during 7 days. The bisphosphonate pamidronate was given subcutaneously at a dose which normalized bone resorption, as reflected by the correction of fasting urinary calcium excretion (adapted from Rizzoli et al., J Bone Miner Res 4: 759–765, 1989). (B) The free radical scavenger WR-2721 was acutely administered to parathyroid hormone-related proteininfused and pamidronate-treated rats. The arrows connect the points before and after WR-2721 acute administration. WR-2721 acutely increased the clearance of calcium.

This 1078-amino acid polypeptide belongs to the receptor family of 7 transmembrane domains cell membraneassociated and guanine nucleotide-binding protein-coupled. This structure is present in parathyroid cells, in C cells of the thyroid, in renal epithelial cells, and in the central nervous system. A raise in extracellular calcium concentration stimulates protein kinase C and leads to an increase in

cytosolic free calcium, thereby inhibiting parathyroid hormone synthesis and release. Conversely, a decrease in plasma calcium triggers the exocytosis of PTH within seconds, whereas it takes hours to increase PTH synthesis, and days to stimulate parathyroid cell proliferation [2, 28, 56]. Various mutations have been reported, which account for hyper- or hyposecretion of parathyroid hormone

Chapter 20 Physiology of Calcium and Phosphate Homeostases Table II.

Hypercalcemic Disorders Increased bone resorption

Endocrine disorders Primary hyperparathyroidism Hyperthyroidism Malignancy Granulomatous disorders (with calcitriol production) Disuse Drug-induced Vitamin D poisoning Milk-Alkali syndrome Thiazide diuretics Lithium salts Benign familial hypocalciuric hypercalcemia

Increased renal tubular reabsorption of calcium

+ ++ + or ++ ++

+ − + or − −

++

−

++ − − − −

− + + + +

Figure 4

351

in relation to variations in extracellular calcium concentration [58]. In familial hypocalciuric benign hypercalcemia, circulating levels of parathyroid hormone are insufficiently suppressed for the degree of calcemia, and the renal tubular reabsorption of calcium is increased through a parathyroid hormone-independent mechanism [39, 40, 59]. This disorder appears to be due to mutations associated with hypofunction of the cell membrane calcium-sensing mechanism [58, 60]. Thus, higher plasma calcium levels are necessary to inhibit PTH secretion. A similar syndrome can be reproduced in transgenic mice by the incorporation of transgenes displaying the same mutations [61]. Interestingly, the same biochemical pattern can be encountered in patients treated with lithium salts [62, 63]. Conversely, in certain cases of familial

Relationship between bone resorption, as evaluated by the bone resorption index (BRI) and tubular reabsorption of calcium index (TRCaI) in rehydrated patients with malignant (A) or non malignant (B) hypercalcemia. In parentheses the mean plasma calcium concentration is given. This figure is taken from Buchs et al. (Bone 12: 47–56, 1991), with the permission of the publisher.

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hypoparathyroidism, a mutation-induced hyperactive calcium-sensing receptor has been found [59]. Under these conditions, parathyroid hormone production is not stimulated despite reduced calcium levels. Calcium-sensing receptor activity can be modulated by calcimimetics, which enhance its sensitivity to extracellular calcium [58]. Under these conditions, PTH secretion is inhibited. Conversely, calcilytics are negative modulators of the calcium-sensing receptor, leading thereby to a stimulation of PTH secretion. There are other structures recognizing changes in extracellular calcium concentration [64–67]. In osteoblasts, they may be distinct from those present in parathyroid cells [68–70].

B. Effectors The perturbation of plasma calcium is corrected by the stimulation of the tubular reabsorption of calcium, and the mobilization of calcium from bone mineral (Fig. 5). Parathyroid hormone itself, as well as a decrease in calcium and/or phosphate concentrations, directly stimulates the synthesis of calcitriol at the renal level. The latter hormone contributes to an increase in plasma calcium through a mobilization of calcium from bone and a stimulation of intestinal calcium absorption. Thus, calcitriol plays a central role in the intestinal adaptation to low calcium intake. This adaptative mechanism is blunted in the elderly, as a consequence of a decreased synthesis of, and a lower response to, calcitriol [71, 72]. It appears therefore that the integrated control of extracellular calcium homeostasis is governed by the requirement of maintaining extracellular calcium concentration in a very narrow range. This system

Figure 5

is quite efficient in its capacity to respond to a calcium need. Conversely, prevention of overwhelming by calcium is ensured by the reversal of these systemic mechanisms. Although calcitonin is able to inhibit bone resorption and to increase the renal clearance of calcium, when given in pharmacological doses, it is improbable that this hormone significantly participates in the physiological defense against hypercalcemia. During body growth, or in the third trimester of pregnancy, when calcification of the fetus bone takes place, the need for calcium results in an increase in intestinal calcium absorption. Elevated levels of calcitriol are responsible for stimulating intestinal calcium absorption [16, 18, 19]. On the other hand, with, for instance, estrogen deprivation at menopause, bone bidirectional calcium fluxes are increased, with resorption overcoming accretion, resulting in a net negative skeletal balance. Under these conditions, a reduced intestinal calcium absorption and/or a higher urinary calcium excretion can be viewed as a homeostatic mechanism attempting to prevent the extracellular calcium compartment from being overloaded. A similar reaction can occur with prolonged immobilization and the consequent net calcium loss.

VII. CALCIUM AND BONE GROWTH Several observational studies have suggested that increasing the calcium intake would promote a greater bone mass gain, and thereby a higher peak bone mass [5]. Furthermore, several prospective randomized, doubleblind, placebo-controlled intervention trials indicate that calcium supplementation can increase bone mass gain,

Homeostatic response to a decrease in extracellular (EC) calcium concentration.

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although the magnitude of the calcium effects appears to vary according to the skeletal sites examined, the stage of pubertal maturation at the time of the intervention, and the spontaneous calcium intake [73–78]. Furthermore, these effects could be modulated by an interaction with vitamin D receptor genotype [79]. The positive effects of calcium supplementation have essentially been ascribed to a reduction in bone remodeling. Indeed, in one study the plasma level of osteocalcin, a biochemical marker of bone remodeling in adults, was significantly reduced in the calciumsupplemented children [73]. This interpretation is also in keeping with the currently favored mechanism proposed to account for the inhibitory effect of calcium on age-related bone loss. In the elderly, the calcium effect on bone remodeling is usually ascribed to an inhibition of parathyroid hormone secretion, as the plasma level tends to increase with aging [80–85]. In a double-blind, placebo-controlled study on the effects of calcium supplementation in prepubertal girls, changes in scanned bone area and in standing height suggest that calcium supplementation could affect bone modeling in addition to bone remodeling [76, 77]. Indeed, milk calcium-enriched foods enhanced the gain of both mean scanned bone area of several skeletal sites, and statural height in the group of spontaneously low-calcium consumers, to the level achieved by the spontaneously highcalcium consumers. Morphometric analysis of the changes observed in the lumbar spine and in femoral diaphysis suggests that calcium could enhance both the longitudinal and the cross-sectional growth of the bones [76, 77]. Effects of calcium on the secretion or action of growth factors, including the IGFs-IGF-binding proteins system, could be implicated [86, 87]. Bone mineral density was measured 7.5 years after the end of calcium supplementation. In these young adult girls, it appeared that menarche occurred earlier in the calcium-supplemented group, and that persistent effects of calcium were mostly detectable in those subjects with an earlier puberty [77]. Finally, because of a higher response to calcium supplements preferentially observed in a certain vitamin D receptor genotype [79], it remains to be determined whether the interaction between vitamin D receptor genotype and the influence of calcium supplementation on bone mass accrual affects modeling and/or remodeling.

VIII. BODY DISTRIBUTION OF PHOSPHORUS Phosphorus is the sixth element in abundance in the body, mostly under the form of phosphate. Its mass represents about 1% of body weight, i.e. 700 g for an individual of

70 kg [1]. More than 80% of phosphate is in bone, the remaining in soft tissues, in the cellular and extracellular compartments. Inside the cells, phosphate can be either under an inorganic form, or an organic one, as a constituent of carbohydrates, lipids, or nucleic acids (Table I). It plays a crucial role in the energy storage and delivery, in the regulation of a large variety of different enzymes activity, and in intracellular signal transduction mechanisms. It also contributes to ensure intracellular and plasma membrane stability, and is implicated in the modulation of hemoglobin affinity for oxygen. Phosphate enters the cells through an active transport system, the energy for which is provided by the extra/intracellular sodium gradient [88, 89]. Extracellular phosphate represents 0.1% of body phosphate. In plasma, one-third of phosphate is inorganic, of which 90% is ultrafiltrable. Two-thirds are parts of circulating phospholipids. At physiological pH, the divalent form of inorganic phosphate predominates. This anion contributes to the maintenance of extracellular acid–base homeostasis. Inorganic phosphate concentration varies in relation to age and body growth rate. In adults, its concentration ranges between 0.8 and 1.4 mmol/L, whereas it is 1.4–2.7 and 1.3–2.0 in the neonate and the adolescent, respectively. There is a nycthemeral rhythm in inorganic phosphate concentration, with highest values encountered in the late afternoon.

IX. DETERMINANTS OF EXTRACELLULAR PHOSPHATE CONCENTRATION The levels of extracellular phosphate are determined by the balance between dynamic fluxes from and into extracellular compartment, which occur in intestine, bone, soft tissue, and kidney (Fig. 6).

A. Intestinal Fluxes A balanced normal diet provides sufficient amounts of phosphate in most circumstances, so that phosphate deficiency from dietary origin usually does not occur. In the intestinal lumen, phosphate can be complexed with various cations, including aluminum or calcium, which thereby prevents its absorption. This property is applied in the prevention of phosphate overload in the frame of advanced renal failure and phosphate retention. The jejunum exhibits the highest absorptive capacity. Two mechanisms are involved. The first one, which predominates in the proximal small intestine, is an active and saturable transport system, activated by calcitriol [17, 21]. The second one is a passive

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Figure 6

Main fluxes (mg/day) controlling phosphate homeostasis.

and nonsaturable transport, which plays quantitatively the most important role. Indeed, an increase in phosphate intake is accompanied by a commensurate increment of net intestinal phosphate absorption. The saturable component becomes negligible at high phosphate intake. Under normal conditions, 60–70% of dietary phosphate is absorbed. At steady state, daily urinary phosphate excretion is a direct reflection of phosphate absorbed in the intestine, and thus of the phosphate intake.

B. Bone Fluxes The processes of bone formation and resorption implicate bidirectional fluxes of phosphate from and into bone mineral. Osteoclastic bone resorption releases phosphate into the extracellular fluid. The deposition of phosphate into newly formed osteoid tissue is dependent on its extracellular concentration. In addition, the plasma membrane of osteoblastic cells, as well as extracellular matrix vesicles found in epiphyseal cartilage and in woven bone, are endowed with a saturable, carrier-mediated phosphate transport system, modulated by various hormones and growth factors [90–92]. These extracellular matrix vesicles could play a role in the initiation of the calcification process. The plasma membrane-associated receptor for gibbon ape leukemia virus [93], which functions as a sodium-dependent phosphate transporter, distinct from the renal type I and II sodium-dependent phosphate transporters, has been found in osteoblastic cells and is regulated by IGF-I [94]. Thus, this type III phosphate transporter could participate in the control of the flux of phosphate deposition into bone.

C. Renal Fluxes Approximately 70% of the filtered inorganic phosphate is reabsorbed in the proximal tubule through saturable sodium co-transport mechanisms [95], whereas about 20% of the filtered load is excreted in the urine (Fig. 7). An important step in the transfer of phosphate from the lumen to peritubular capillaries takes place in the epithelial cell brush-border membrane. At least two different sodiumdependent phosphate transporters have been identified in the renal tubule, which share little homology in their amino acid sequence [88, 89]. The type I transporter appears to be constitutive, nonselective, and unregulated. The type II transporter is controlled by dietary phosphate intake or by parathyroid hormone/parathyroid hormone-related protein. Short-term exposure to a low-phosphate diet stimulates the insertion of type II phosphate transporters into the apical membrane, whereas parathyroid hormone decreases it. Prolonged low dietary phosphate intakes increase the expression of type II transporter. Its presence in the brush border membrane is markedly depressed in X-linked hypophosphatemic rachitic mice [88, 89]. However, this defect has been mapped to another gene than that coding for the type II phosphate transporter. The gene mutated in the case of X-linked hypophosphatemia codes for an endopeptidase, for which the substrate is, however, still not clearly elucidated [96–99]. The maximal tubular reabsorption capacity (TmPi/ GFR) is the main determinant of plasma phosphate concentration [95]. Since the tubular reabsorption is a saturable process, fractional reabsorption for a given TmPi/GFR varies according to the filtered load. Therefore, fractional

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Chapter 20 Physiology of Calcium and Phosphate Homeostases

Figure 7

Central role of the kidney in controlling phosphate homeostasis.

excretion or reabsorption cannot be taken as a reliable reflection of the tubular reabsorptive capacity. This capacity is controlled by the phosphate supply and the need for phosphate of the organism [100]. Not only is the low dietary phosphate-mediated stimulation of phosphate transport independent of parathyroid hormone, but it prevents the phosphaturic response to the hormone [101]. During growth, a state of need for phosphate, or during phosphate deficiency, TmPi/GFR is increased [95]. In proximal tubule kidney cells, phosphate transport is activated by IGF-I, whereas it is decreased by parathyroid hormone, or its tumoral analog parathyroid hormonerelated protein [102–104]. Other factors could influence the sodium-dependent phosphate transport [47, 48]; however, their contribution to alterations observed in physiological or pathophysiological situations remains to be established. IGF-I, which plays a prominent role in the longitudinal growth of bone, also increases the renal synthesis of calcitriol [105], which in turn stimulates the intestinal absorption of phosphate [17, 21] (Table III). Table III.

Thus, by acting indirectly on the intestine through calcitriol, and directly on the kidney, IGF-I contributes to make positive the phosphate balance, favoring thereby the mineralization of newly formed bone (Fig. 8). Parathyroid hormone, or parathyroid hormone-related protein, as well as a state of phosphate overload, or elevated extracellular calcium concentration reduce TmPi/GFR. Calcitriol appears to be devoid of any direct and significant effect on the renal tubular reabsorption of phosphate under physiological conditions. Factor(s) other than PTH or PTHrP are known to increase phosphate excretion, through influencing renal tubular reabsorption of phosphate. Although the molecules implicated are not fully elucidated, the name “phosphatonin” has been suggested [106, 107]. Evidence for the existence of such factor(s) has been obtained from pathological conditions such as tumor-associated osteomalacia, X-linked hypophosphatemia, or autosomal dominant hypophosphatemic rickets. Genomic and proteomic procedures allowed one to identify fibroblast

Factors Affecting Phosphate Renal Tubular Reabsorption

Factors that decrease Pi reabsorption Phosphate loading Parathyroid hormone/PTHrP/cAMP Volume expansion Hypercalcemia Carbonic anhydrase inhibitors Fibroblast growth factor 23 Frizzled receptor protein 4

Factors that increase Pi reabsorption Phosphate depletion Parathyroidectomy Volume contraction Hypocalcemia Growth hormone Insulin-like growth factor-I

Figure 8

Effects of IGF-I on bone and on mineral homeostasis.

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growth factor 23 (FGF23), frizzled receptor protein 4 (FRP4), and matrix extracellular phosphoglycoprotein (MEPE) has exhibited phosphaturic activity [107–110] (Fig. 9).

D. Soft Tissues In constrast to calcium, phosphate fluxes from and into the intracellular compartment can alter extracellular phosphate concentration. During fasting, phosphate mobilization from soft tissues, such as the liver, can contribute to an increase in plasma phosphate. Inversely, during the postabsorptive phase, extracellular phosphate is transferred into soft tissues, as a result of incorporation into carbohydrates or lipids under the influence of insulin [111, 112]. Hyperphosphatemia, in the frame of a tumor lysis syndrome following the initiation of cytotoxic therapy of hematologic malignancies, or hypophosphatemia occurring during the treatment of diabetic ketoacidosis with insulin, are examples of the role played by soft tissues in phosphate homeostasis under pathological conditions [113, 114].

X. HOMEOSTATIC RESPONSES TO CHANGES IN PHOSPHATE SUPPLY OR DEMAND Phosphate deficiency stimulates calcitriol production, which increases the transfer of phosphate from the intestine and mobilization from bone mineral (Fig. 10). The renal synthesis and release of calcitriol are related to the phosphate supply and the demand of the organism. On the other hand, low extracellular phosphate concentration is associated with higher calcium concentration through a physicochemical process. In turn, the inhibition of parathyroid hormone release decreases urinary phosphate excretion.

Figure 9

Effects of “phosphatonins” on Pi homeostasis.

Figure 10 Renal Pi handling and serum FGF23 levels in relation to dietary Pi intakes (from Ferrari et al., J Clin Endocrinol Metab 2005, with permission of the publisher).

The most powerful mechanism for phosphate retention is certainly the system, the molecular nature of which is still not elucidated, which allows the kidney to adapt very tightly its capacity of reabsorbing phosphate to dietary supplies and to the need of the organism in phosphate, in a parathyroid hormone-independent manner [88, 89, 95, 96, 100, 101, 115]. FGF23 could contribute to this dietary phosphate-induced modulation of renal tubular transport. Indeed, increasing dietary phosphate intakes was associated with higher FGF23 levels in young healthy males undergoing acute changes in phosphate supply [116] (Fig. 11). The adaptation to low phosphate intakes in terms of calcitriol production and changes in renal tubular phosphate reabsorption is missing in X-linked hypophosphatemic rickets [115]. This indicates that the homeostatic response to meet the phosphate supply and need is altered in this disorder, leading to a renal phosphate leak and inadequate calcitriol levels for the degree of hypophosphatemia, despite the need for phosphate for bone growth and mineralization. Various mutations in the gene to which X-linked hypophosphatemic rickets has been mapped, have been reported, both in humans and in mice [97–99].

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Sensitive and efficient regulatory mechanisms involving local calcium sensing are triggered by changes in calcium demand or supply. Similarly, the renal handling of phosphate can adjust its capacity to meet the need of the organism. The regulation of calcium and phosphate homeostases aims at fighting against a deficiency or an overload. In response to a variation of any regulatory flux, a series of homeostatic responses and adjustments is triggered, leading to a new steady state, in which the initially altered variable has been corrected.

Acknowledgments Figure 11

Homeostatic response to a decrease in extracellular (EC) phosphate concentration.

The role played by this PHEX (phosphate-regulating gene with homology to endopeptidase on the X chromosome) gene product, which is a zinc-binding endopeptidase, in the regulation of phosphate homeostasis, is still not elucidated. As mentioned above, this phosphate homeostatic mechanism could play a role in the increase of renal tubular reabsorption of phosphate observed during growth, possibly in association with IGF-I, but also in disorders characterized by an increased demand, as for instance in extensive osteoblastic metastases [117]. An inhibition of the same system could also be implicated to avoid phosphate overload in the frame of a decreased nephron mass, as for instance in progressive renal failure [118, 119]. There is also evidence that elevated extracellular phosphate concentration could also be able to directly increase parathyroid hormone production [120]. This mechanism is apparently less efficacious than the one triggered by hypocalcemia. Because of the phosphaturic effects of parathyroid hormone, such a regulatory system certainly represents an attempt to maintain the homeostasis of phosphate economy.

XI. CONCLUSIONS The physiology of calcium and phosphate homeostases is regulated by coordinated bidirectional calcium and phosphate fluxes, occurring at the levels of intestine, bone, and kidney. These fluxes are influenced by calciotropic peptides or steroid hormones, and by a variety of locally produced factors. By directly modifying renal tubular calcium and phosphate transport, or by releasing calcium from intracellular stores, calcium itself functions as a regulator. In the control of extracellular concentration of either ion, the kidney tubule reabsorptive capacity plays a central role.

We thank Mrs M. Perez for her expert secretarial assistance.

References 1. Broadus, A.E. (2003). Mineral balance and homeostasis. In “Primer on the Metabolic Bone Diseases and Disorders of Mineral Metabolism” (M. J. Favus, ed.), pp. 105–111. American Society for Bone and Mineral Research, Washington, D.C. 2. Brown, E. M., and MacLeod, R. J. (2001). Extracellular calcium sensing and extracellular calcium signaling. Physiol. Rev. 81, 239–297. 3. Bonjour, J. P., Theintz, G., Buchs, B., Slosman, D., and Rizzoli, R. (1991). Critical years and stages of puberty for spinal and femoral bone mass accumulation during adolescence. J. Clin. Endocrinol. Metab. 73, 555–563. 4. Theintz, G., Buchs, B., Rizzoli, R., Slosman, D., Clavien, H., Sizonenko, P. C., and Bonjour, J. P. (1992). Longitudinal monitoring of bone mass accumulation in healthy adolescents: evidence for a marked reduction after 16 years of age at the levels of lumbar spine and femoral neck in female subjects. J. Clin. Endocrinol. Metab. 75, 1060–1065. 5. Bonjour, J. P., and Rizzoli, R. (2001). Bone acquisition in adolescence. In “Osteoporosis” (R. Marcus, D. Feldman, and J. Kelsey, eds.), pp. 621–638. Academic Press, San Diego, CA. 6. Blair, H. C., Robinson, L. J., and Zaidi, M. (2005). Osteoclast signalling pathways. Biochem. Biophys. Res. Commun. 328, 728–738. 7. Parekh, A. B., and Putney, J. W., Jr. (2005). Store-operated calcium channels. Physiol. Rev. 85, 757–810. 8. Dvorak, M. M., and Riccardi, D. (2004). Ca2+ as an extracellular signal in bone. Cell Calcium 35, 249–255. 9. Szabadkai, G., and Rizzuto, R. (2004). Participation of endoplasmic reticulum and mitochondrial calcium handling in apoptosis: more than just neighborhood? FEBS Lett. 567, 111–115. 10. Reeves, J. P., Condrescu, M., Chernaya, G., and Gardner, J. P. (1994). Na+/Ca2+ antiport in the mammalian heart. J. Exp. Biol. 196, 375–388. 11. Carafoli, E. (2002). Calcium signaling: a tale for all seasons. Proc. Natl Acad. Sci. USA 99, 1115–1122. 12. Carafoli, E. (2004). Calcium-mediated cellular signals: a story of failures. Trends Biochem. Sci. 29, 371–379. 13. Rodland, K. D. (2004). The role of the calcium-sensing receptor in cancer. Cell Calcium 35, 291–295. 14. McConkey, D. J., and Orrenius, S. (1996). The role of calcium in the regulation of apoptosis. J. Leukoc. Biol. 59, 775–783. 15. Rizzoli, R., Fleisch, H., and Bonjour, J. P. (1977). Effect of thyroparathyroidectomy of calcium metabolism in rats: role of 1,25dihydroxyvitamin D3. Am. J. Physiol. 233, E160–164.

358 16. Lopez-Hilker, S., Dusso, A. S., Rapp, N. S., Martin, K. J., and Slatopolsky, E. (1990). Phosphorus restriction reverses hyperparathyroidism in uremia independent of changes in calcium and calcitriol. Am. J. Physiol. 259, F432–437. 17. Hoenderop, J. G., Nilius, B., and Bindels, R. J. (2005). Calcium absorption across epithelia. Physiol. Rev. 85, 373–422. 18. Holick, M. F. (2003). Vitamin D: photobiology, metabolism, mechanism of action, and clinical applications. In “Primer on the Metabolic Bone Diseases and Disorders of Mineral Metabolism” (M. J. Favus, ed.), pp. 129–137. American Society for Bone and Mineral Research, Washington, D.C. 19. Feldman, D., Malloy, P. J., and Gross, C. (2001). Biology, action, and clinical implications. In “Osteoporosis” (R. Marcus, D. Feldman, and J. Kelsey, eds.), pp. 257–303. Academic Press, San Diego, CA. 20. Bonjour, J. P., Fleisch, H., and Trechsel, U. (1977). Calcium absorption in diphosphonate-treated rats: effect of parathyroid function, dietary calcium and phosphorus. J. Physiol. 264, 125–139. 21. Rizzoli, R., Fleisch, H., and Bonjour, J. P. (1977). Role of 1,25dihydroxyvitamin D3 on intestinal phosphate absorption in rats with a normal vitamin D supply. J. Clin. Invest. 60, 639–647. 22. Heaney, R. P., Weaver, C. M., and Recker, R. R. (1988). Calcium absorbability from spinach. Am. J. Clin. Nutr. 47, 707–709. 23. Heaney, R. P., and Weaver, C. M. (1990). Calcium absorption from kale. Am. J. Clin. Nutr. 51, 656–657. 24. Ammann, P., Rizzoli, R., and Fleisch, H. (1986). Calcium absorption in rat large intestine in vivo: availability of dietary calcium. Am. J. Physiol. 251, G14–18. 25. Pak, C. Y., Kaplan, R., Bone, H., Townsend, J., and Waters, O. (1975). A simple test for the diagnosis of absorptive, resorptive and renal hypercalciurias. N. Engl. J. Med. 292, 497–500. 26. Lemann, J., and Favus, M. J. (2003). Intestinal absorption of calcium, magnesium and phosphate. In “Primer on the Metabolic Bone Diseases and Disorders of Mineral Metabolism” (M. J. Favus, ed.), pp. 93–97. American Society for Bone and Mineral Research, Washington, D.C. 27. Hefti, E., Trechsel, U., Fleisch, H., and Bonjour, J. P. (1983). Nature of calcemic effect of 1,25-dihydroxyvitamin D3 in experimental hypoparathyroidism. Am. J. Physiol. 244, E313–316. 28. Jüppner, H., and Kronenberg, H. M. (2003). Parathyroid hormone. In “Primer on the Metabolic Bone Diseases and Disorders of Mineral Metabolism” (M. J. Favus, ed.), pp. 117–124. American Society for Bone and Mineral Research, Washington, D.C. 29. Mundy, G. R., Chen, D., and Oyajobi, B. O. (2003). Bone remodeling. In “Primer on the Metabolic Bone Diseases and Disorders of Mineral Metabolism” (M. J. Favus, ed.), pp. 46–58, American Society for Bone and Mineral Research, Washington, D.C. 30. Manolagas, S. C. (2000). Birth and death of bone cells: basic regulatory mechanisms and implications for the pathogenesis and treatment of osteoporosis. Endocr. Rev. 21, 115–137. 31. Martin, T. J., and Rodan, G. A. (2001). Coupling of bone resorption and formation during bone remodeling. In “Osteoporosis” (R. Marcus, D. Feldman, and J. Kelsey, eds.), pp. 361–371. Academic Press, San Diego, CA. 32. Hofbauer, L. C., and Schoppet, M. (2004). Clinical implications of the osteoprotegerin/RANKL/RANK system for bone and vascular diseases. JAMA 292, 490–495. 33. Peacock, M., Robertson, W. G., and Nordin, B. E. (1969). Relation between serum and urinary calcium with particular reference to parathyroid activity. Lancet 1, 384–386. 34. Nordin, B. E., and Peacock, M. (1969). Role of kidney in regulation of plasma-calcium. Lancet 2, 1280–1283. 35. Bonjour, J. P., Philippe, J., Guelpa, G., Bisetti, A., Rizzoli, R., Jung, A., Rosini, S., and Kanis, J. A. (1988). Bone and renal components in hypercalcemia of malignancy and responses to a single infusion of clodronate. Bone 9, 123–130.

RENÉ RIZZOLI AND JEAN-PHILIPPE BONJOUR 36. Parsons, J. A., and Robinson, C. J. (1971). Calcium shift into bone causing transient hypocalcaemia after injection of parathyroid hormone. Nature 230, 581–582. 37. Parsons, J. A., Neer, R. M., and Potts, J. T., Jr. (1971). Initial fall of plasma calcium after intravenous injection of parathyroid hormone. Endocrinology 89, 735–740. 38. Friedman, P. A. (2000). Mechanisms of renal calcium transport. Exp. Nephrol. 8, 343–350. 39. Marx, S. J., Spiegel, A. M., Levine, M. A., Rizzoli, R. E., Lasker, R. D., Santora, A. C., Downs, R. W., Jr., and Aurbach, G. D. (1982). Familial hypocalciuric hypercalcemia: the relation to primary parathyroid hyperplasia. N. Engl. J. Med. 307, 416–426. 40. Attie, M. F., Gill, J. R., Jr., Stock, J. L., Spiegel, A. M., Downs, R. W., Jr., Levine, M. A., and Marx, S. J. (1983). Urinary calcium excretion in familial hypocalciuric hypercalcemia. Persistence of relative hypocalciuria after induction of hypoparathyroidism. J. Clin. Invest. 72, 667–676. 41. Thakker, R. V. (2004). Diseases associated with the extracellular calcium-sensing receptor. Cell Calcium 35, 275–282. 42. Rizzoli, R., Hugi, K., Fleisch, H., and Bonjour, J. P. (1981). Effect of hydrochlorothiazide on 1,25-dihydroxyvitamin D3-induced changes in calcium metabolism in experimental hypoparathyroidism in rats. Clin. Sci. (Lond.) 60, 101–107. 43. Rizzoli, R., Pizurki, L., and Bonjour, J. P. (1989). Tumoral parathyroid hormone-related protein: a new hormone affecting calcium and phosphate transport. NIPS 4, 195–198. 44. Strewler, G. J., and Nissenson, R. A. (2003). Parathyroid hormonerelated protein. In “Primer on the Metabolic Bone Diseases and Disorders of Mineral Metabolism” (M. J. Favus, ed.), pp. 125–129. American Society for Bone and Mineral Research, Washington, D.C. 45. Bisello, A., Horwitz, M. J., and Stewart, A. F. (2004). Parathyroid hormone-related protein: an essential physiological regulator of adult bone mass. Endocrinology 145, 3551–3553. 46. Caverzasio, J., Rizzoli, R., Vallotton, M. B., Dayer, J. M., and Bonjour, J. P. (1993). Stimulation by interleukin-1 of renal calcium reabsorption in thyroparathyroidectomized rats. J. Bone Miner. Res. 8, 1219–1225. 47. Pizurki, L., Rizzoli, R., Caverzasio, J., and Bonjour, J. P. (1990). Effect of transforming growth factor-alpha and parathyroid hormone-related protein on phosphate transport in renal cells. Am. J. Physiol. 259, F929–935. 48. Law, F., Rizzoli, R., and Bonjour, J. P. (1993). Transforming growth factor-beta inhibits phosphate transport in renal epithelial cells. Am. J. Physiol. 264, F623–628. 49. Law, F., Bonjour, J. P., and Rizzoli, R. (1994). Transforming growth factor-beta: a down-regulator of the parathyroid hormone-related protein receptor in renal epithelial cells. Endocrinology 134, 2037–2043. 50. Buchs, B., Rizzoli, R., and Bonjour, J. P. (1991). Evaluation of bone resorption and renal tubular reabsorption of calcium and phosphate in malignant and nonmalignant hypercalcemia. Bone 12, 47–56. 51. Rizzoli, R., Stoermann, C., Ammann, P., and Bonjour, J. P. (1994). Hypercalcemia and hyperosteolysis in vitamin D intoxication: effects of clodronate therapy. Bone 15, 193–198. 52. Rizzoli, R., Caverzasio, J., Chapuy, M. C., Martin, T. J., and Bonjour, J. P. (1989). Role of bone and kidney in parathyroid hormonerelated peptide-induced hypercalcemia in rats. J. Bone Miner. Res. 4, 759–765. 53. Rizzoli, R., Caverzasio, J., Bauss, F., and Bonjour, J. P. (1992). Inhibition of bone resorption by the bisphosphonate BM 21.0955 is not associated with an alteration of the renal handling of calcium in rats infused with parathyroid hormone-related protein. Bone 13, 321–325. 54. Hirschel-Scholz, S., Caverzasio, J., and Bonjour, J. P. (1985). Inhibition of parathyroid hormone secretion and parathyroid hormoneindependent diminution of tubular calcium reabsorption by WR-2721, a unique hypocalcemic agent. J. Clin. Invest. 76, 1851–1856.

Chapter 20 Physiology of Calcium and Phosphate Homeostases 55. Hirschel-Scholz, S., Caverzasio, J., Rizzoli, R., and Bonjour, J. P. (1986). Normalization of hypercalcemia associated with a decrease in renal calcium reabsorption in Leydig cell tumor-bearing rats treated with WR-2721. J. Clin. Invest. 78, 319–322. 56. Chen, R. A., and Goodman, W. G. (2004). Role of the calcium-sensing receptor in parathyroid gland physiology. Am. J. Physiol. Renal. Physiol. 286, F1005–1011. 57. Brown, E. M., Gamba, G., Riccardi, D., Lombardi, M., Butters, R., Kifor, O., Sun, A., Hediger, M. A., Lytton, J., and Hebert, S. C. (1993). Cloning and characterization of an extracellular Ca(2+)-sensing receptor from bovine parathyroid. Nature 366, 575–580. 58. Hu, J., and Spiegel, A. M. (2003). Naturally occurring mutations of the extracellular Ca2+-sensing receptor: implications for its structure and function. Trends Endocrinol. Metab. 14, 282–288. 59. Brown, E. M., Pollak, M. R., and Hebert, S. C. (1998). The extracellular calcium-sensing receptor: its role in health and disease. Annu. Rev. Med. 49, 15–29. 60. Pollak, M. R., Brown, E. M., Chou, Y. H. W., Hebert, S. C., Marx, S. J., Steinmann, B., Levi, T., Seidman, C. E., and Seidman, J. G. (1993). Mutations in the human Ca2+ sensing receptor gene cause familial hypocalciuric hypercalcemia and neonatal severe hyperparathyroidism. Cell 75, 1297–1303. 61. Ho, C., Conner, D. A., Pollak, M. R., Ladd, D. J., Kifor, O., Warren, H. B., Brown, E. M., Seidman, J. G., and Seidman, C. E. (1995). A mouse model of human familial hypocalciuric hypercalcemia and neonatal severe hyperparathyroidism. Nat. Genet. 11, 389–394. 62. Mallette, L. E., and Eichhorn, E. (1986). Effects of lithium carbonate on human calcium metabolism. Arch. Intern. Med. 146, 770–776. 63. Bendz, H., Sjodin, I., Toss, G., and Berglund, K. (1996). Hyperparathyroidism and long-term lithium therapy – a cross-sectional study and the effect of lithium withdrawal. J. Intern. Med. 240, 357–365. 64. House, M. G., Kohlmeier, L., Chattopadhyay, N., Kifor, O., Yamaguchi, T., Leboff, M. S., Glowacki, J., and Brown, E. M. (1997). Expression of an extracellular calcium-sensing receptor in human and mouse bone marrow cells. J. Bone Miner. Res. 12, 1959–1970. 65. Sands, J. M., Naruse, M., Baum, M., Jo, I., Hebert, S. C., Brown, E. M., and Harris, H. W. (1997). Apical extracellular calcium/polyvalent cation-sensing receptor regulates vasopressin-elicited water permeability in rat kidney inner medullary collecting duct. J. Clin. Invest. 99, 1399–1405. 66. Chattopadhyay, N., Cheng, I., Rogers, K., Riccardi, D., Hall, A., Diaz, R., Hebert, S. C., Soybel, D. I., and Brown, E. M. (1998). Identification and localization of extracellular Ca(2+)-sensing receptor in rat intestine. Am. J. Physiol. 274, G122–130. 67. Buchs, N., Manen, D., Bonjour, J. P., and Rizzoli, R. (2000). Calcium stimulates parathyroid hormone-related protein production in Leydig tumor cells through a putative cation-sensing mechanism. Eur. J. Endocrinol. 142, 500–505. 68. Quarles, L. D., Hartle, J. E., 2nd, Siddhanti, S. R., Guo, R., and Hinson, T. K. (1997). A distinct cation-sensing mechanism in MC3T3-E1 osteoblasts functionally related to the calcium receptor. J. Bone Miner. Res. 12, 393–402. 69. Pi, M., and Quarles, L. D. (2004). A novel cation-sensing mechanism in osteoblasts is a molecular target for strontium. J. Bone Miner. Res. 19, 862–869. 70. Yamaguchi, T., Kifor, O., Chattopadhyay, N., and Brown, E. M. (1998). Expression of extracellular calcium (Ca2 + o)-sensing receptor in the clonal osteoblast-like cell lines, UMR-106 and SAOS-2. Biochem. Biophys. Res. Commun. 243, 753–757. 71. Gallagher, J. C., Riggs, B. L., Eisman, J., Hamstra, A., Arnaud, S. B., and DeLuca, H. F. (1979). Intestinal calcium absorption and serum vitamin D metabolites in normal subjects and osteoporotic patients: effect of age and dietary calcium. J. Clin. Invest. 64, 729–736. 72. Ebeling, P. R., Sandgren, M. E., DiMagno, E. P., Lane, A. W., DeLuca, H. F., and Riggs, B. L. (1992). Evidence of an age-related

73.

74.

75.

76.

77.

78.

79.

80.

81.

82.

83.

84.

85.

86.

87.

359 decrease in intestinal responsiveness to vitamin D: relationship between serum 1,25-dihydroxyvitamin D3 and intestinal vitamin D receptor concentrations in normal women. J. Clin. Endocrinol. Metab. 75, 176–182. Johnston, C. C., Jr., Miller, J. Z., Slemenda, C. W., Reister, T. K., Hui, S., Christian, J. C., and Peacock, M. (1992). Calcium supplementation and increases in bone mineral density in children. N. Engl. J. Med. 327, 82–87. Lloyd, T., Andon, M. B., Rollings, N., Martel, J. K., Landis, J. R., Demers, L. M., Eggli, D. F., Kieselhorst, K., and Kulin, H. E. (1993). Calcium supplementation and bone mineral density in adolescent girls. JAMA 270, 841–844. Lee, W. T., Leung, S. S., Xu, Y. C., Wang, S. H., Zeng, W. P., Lau, J., and Fairweather-Tait, S. J. (1995). Effects of double-blind controlled calcium supplementation on calcium absorption in Chinese children measured with stable isotopes (42Ca and 44Ca). Br. J. Nutr. 73, 311–321. Bonjour, J. P., Carrie, A. L., Ferrari, S., Clavien, H., Slosman, D., Theintz, G., and Rizzoli, R. (1997). Calcium-enriched foods and bone mass growth in prepubertal girls: a randomized, double-blind, placebocontrolled trial. J. Clin. Invest. 99, 1287–1294. Chevalley, T., Rizzoli, R., Hans, D., Ferrari, S., and Bonjour, J. P. (2005). Interaction between calcium intake and menarcheal age on bone mass gain: an eight-year follow-up study from prepuberty to postmenarche. J. Clin. Endocrinol. Metab. 90, 44–51. Chevalley, T., Bonjour, J. P., Ferrari, S., Hans, D., and Rizzoli, R. (2005). Skeletal site selectivity in the effects of calcium supplementation on areal bone mineral density gain: a randomized, double-blind, placebo-controlled trial in prepubertal boys. J. Clin. Endocrinol. Metab. 90, 3342–3349. Ferrari, S. L., Rizzoli, R., Slosman, D. O., and Bonjour, J. P. (1998). Do dietary calcium and age explain the controversy surrounding the relationship between bone mineral density and vitamin D receptor gene polymorphisms? J. Bone Miner. Res. 13, 363–370. Boonen, S., Rizzoli, R., Meunier, P. J., Stone, M., Nuki, G., Syversen, U., Lehtonen-Veromaa, M., Lips, P., Johnell, O., and Reginster, J. Y. (2004). The need for clinical guidance in the use of calcium and vitamin D in the management of osteoporosis: a consensus report. Osteoporos Int. 15, 511–519. Shea, B., Wells, G., Cranney, A., Zytaruk, N., Robinson, V., Griffith, L., Hamel, C., Ortiz, Z., Peterson, J., Adachi, J., Tugwell, P., and Guyatt, G. (2003). Calcium supplementation on bone loss in postmenopausal women. Cochrane Database Syst. Rev. CD004526. Shea, B., Wells, G., Cranney, A., Zytaruk, N., Robinson, V., Griffith, L., Hamel, C., Ortiz, Z., Peterson, J., Adachi, J., Tugwell, P., and Guyatt, G. (2004). Calcium supplementation on bone loss in postmenopausal women. Cochrane Database Syst. Rev. CD004526. Chapuy, M. C., Arlot, M. E., Duboeuf, F., Brun, J., Crouzet, B., Arnaud, S., Delmas, P. D., and Meunier, P. J. (1992). Vitamin D3 and calcium to prevent hip fractures in the elderly women. N. Engl. J. Med. 327, 1637–1642. Chevalley, T., Rizzoli, R., Nydegger, V., Slosman, D., Rapin, C. H., Michel, J. P., Vasey, H., and Bonjour, J. P. (1994). Effects of calcium supplements on femoral bone mineral density and vertebral fracture rate in vitamin-D-replete elderly patients. Osteoporos. Int. 4, 245–252. Dawson-Hughes, B., Harris, S. S., Krall, E. A., and Dallal, G. E. (1997). Effect of calcium and vitamin D supplementation on bone density in men and women 65 years of age or older. N. Engl. J. Med. 337, 670–676. Honda, Y., Fitzsimmons, R. J., Baylink, D. J., and Mohan, S. (1995). Effects of extracellular calcium on insulin-like growth factor II in human bone cells. J. Bone Miner. Res. 10, 1660–1665. Yoo, A., Tanimoto, H., Akesson, K., Baylink, D. J., and Lau, K. H. (1998). Effects of calcium depletion and repletion on serum insulin-like growth factor I and binding protein levels in weanling rats. Bone 22, 225–232.

360 88. Murer, H., Hernando, N., Forster, I., and Biber, J. (2003). Regulation of Na/Pi transporter in the proximal tubule. Annu. Rev. Physiol. 65, 531–542. 89. Bacic, D., Wagner, C. A., Hernando, N., Kaissling, B., Biber, J., and Murer, H. (2004). Novel aspects in regulated expression of the renal type IIa Na/Pi-cotransporter. Kidney Int. Suppl. S5–S12. 90. Pizurki, L., Rizzoli, R., Caverzasio, J., and Bonjour, J. P. (1991). Stimulation by parathyroid hormone-related protein and transforming growth factor-alpha of phosphate transport in osteoblast-like cells. J. Bone Miner. Res. 6, 1235–1241. 91. Montessuit, C., Bonjour, J. P., and Caverzasio, J. (1995). Expression and regulation of Na-dependent P(i) transport in matrix vesicles produced by osteoblast-like cells. J. Bone Miner. Res. 10, 625–631. 92. Caverzasio, J., and Bonjour, J. P. (1996). Characteristics and regulation of Pi transport in osteogenic cells for bone metabolism. Kidney Int. 49, 975–980. 93. Kavanaugh, M. P., Miller, D. G., Zhang, W., Law, W., Kozak, S. L., Kabat, D., and Miller, A. D. (1994). Cell-surface receptors for gibbon ape leukemia virus and amphotropic murine retrovirus are inducible sodium-dependent phosphate symporters. Proc. Natl Acad. Sci. USA 91, 7071–7075. 94. Palmer, G., Bonjour, J. P., and Caverzasio, J. (1997). Expression of a newly identified phosphate transporter/retrovirus receptor in human SaOS-2 osteoblast-like cells and its regulation by insulin-like growth factor I. Endocrinology 138, 5202–5209. 95. Bonjour, J. P., and Caverzasio, J. (1984). Phosphate transport in the kidney. Rev. Physiol. Biochem. Pharmacol. 100, 161–214. 96. Custer, M., Spindler, B., Verrey, F., Murer, H., and Biber, J. (1997). Identification of a new gene product (diphor-1) regulated by dietary phosphate. Am. J. Physiol. 273, F801–806. 97. The HYP Consortium (1995). A gene (PEX) with homologies to endopeptidases is mutated in patients with X-linked hypophosphatemic rickets. The HYP Consortium. Nat. Genet. 11, 130–136. 98. Du, L., Desbarats, M., Viel, J., Glorieux, F. H., Cawthorn, C., and Ecarot, B. (1996). cDNA cloning of the murine Pex gene implicated in X-linked hypophosphatemia and evidence for expression in bone. Genomics 36, 22–28. 99. Econs, M. J., and Francis, F. (1997). Positional cloning of the PEX gene: new insights into the pathophysiology of X-linked hypophosphatemic rickets. Am. J. Physiol. 273, F489–498. 100. Trohler, U., Bonjour, J. P., and Fleisch, H. (1976). Inorganic phosphate homeostasis. Renal adaptation to the dietary intake in intact and thyroparathyroidectomized rats. J. Clin. Invest. 57, 264–273. 101. Trohler, U., Bonjour, J. P., and Fleisch, H. (1976). Renal tubular adaptation to dietary phosphorus. Nature 261, 145–146. 102. Caverzasio, J., Rizzoli, R., and Bonjour, J. P. (1986). Sodium-dependent phosphate transport inhibited by parathyroid hormone and cyclic AMP stimulation in an opossum kidney cell line. J. Biol. Chem. 261, 3233–3237. 103. Caverzasio, J., and Bonjour, J. P. (1989). Insulin-like growth factor I stimulates Na-dependent Pi transport in cultured kidney cells. Am. J. Physiol. 257, F712–717. 104. Pizurki, L., Rizzoli, R., Moseley, J., Martin, T. J., Caverzasio, J., and Bonjour, J. P. (1988). Effect of synthetic tumoral PTH-related peptide

RENÉ RIZZOLI AND JEAN-PHILIPPE BONJOUR

105.

106.

107. 108.

109.

110.

111. 112.

113. 114.

115.

116.

117.

118.

119.

120.

on cAMP production and Na-dependent Pi transport. Am. J. Physiol. 255, F957–961. Caverzasio, J., Montessuit, C., and Bonjour, J. P. (1990). Stimulatory effect of insulin-like growth factor-1 on renal Pi transport and plasma 1,25-dihydroxyvitamin D3. Endocrinology 127, 453–459. Dunstan, C. R., Zhou, H., and Seibel, M. J. (2004). Fibroblast growth factor 23: a phosphatonin regulating phosphate homeostasis? Endocrinology 145, 3084–3086. Schiavi, S. C., and Kumar, R. (2004). The phosphatonin pathway: new insights in phosphate homeostasis. Kidney Int. 65, 1–14. Riminucci, M., Collins, M. T., Fedarko, N. S., Cherman, N., Corsi, A., White, K. E., Waguespack, S., Gupta, A., Hannon, T., Econs, M. J., Bianco, P., and Gehron Robey, P. (2003). FGF-23 in fibrous dysplasia of bone and its relationship to renal phosphate wasting. J. Clin. Invest. 112, 683–692. Berndt, T., Craig, T. A., Bowe, A. E., Vassiliadis, J., Reczek, D., Finnegan, R., Jan De Beur, S. M., Schiavi, S. C., and Kumar, R. (2003). Secreted frizzled-related protein 4 is a potent tumor-derived phosphaturic agent. J. Clin. Invest. 112, 785–794. Quarles, L. D. (2003). FGF23, PHEX, and MEPE regulation of phosphate homeostasis and skeletal mineralization. Am. J. Physiol. Endocrinol. Metab. 285, E1–9. Marinella, M. A. (2005). Refeeding syndrome and hypophosphatemia. J. Intensive Care Med. 20, 155–159. Takeda, E., Yamamoto, H., Nashiki, K., Sato, T., Arai, H., and Taketani, Y. (2004). Inorganic phosphate homeostasis and the role of dietary phosphorus. J. Cell Mol. Med. 8, 191–200. Bohannon, N. J. (1989). Large phosphate shifts with treatment for hyperglycemia. Arch. Intern. Med. 149, 1423–1425. Jones, D. P., Mahmoud, H., and Chesney, R. W. (1995). Tumor lysis syndrome: pathogenesis and management. Pediatr. Nephrol. 9, 206–212. Hruska, K. A., Rifas, L., Cheng, S. L., Gupta, A., Halstead, L., and Avioli, L. (1995). X-linked hypophosphatemic rickets and the murine Hyp homologue. Am. J. Physiol. 268, F357–362. Ferrari, S. L., Bonjour, J. P., and Rizzoli, R. (2005). Fibroblast growth factor-23 relationship to dietary phosphate and renal phosphate handling in healthy young men. J. Clin. Endocrinol. Metab. 90, 1519–1524. Buchs, N., Bonjour, J. P., and Rizzoli, R. (1998). Renal tubular reabsorption of phosphate is positively related to the extent of bone metastatic load in patients with prostate cancer. J. Clin. Endocrinol. Metab. 83, 1535–1541. Caverzasio, J., Gloor, H. J., Fleisch, H., and Bonjour, J. P. (1982). Parathyroid hormone-independent adaptation of the renal handling of phosphate in response to renal mass reduction. Kidney Int. 21, 471–476. Bonjour, J. P., Caverzasio, J., and Rizzoli, R. (1992). Phosphate homeostasis, 1,25-dihydroxyvitamin D3, and hyperparathyroidism in early chronic renal failure. Trends Endocrinol. Metab. 3, 391–395. Slatopolsky, E., Finch, J., Denda, M., Ritter, C., Zhong, M., Dusso, A., MacDonald, P. N., and Brown, A. J. (1996). Phosphorus restriction prevents parathyroid gland growth. High phosphorus directly stimulates PTH secretion in vitro. J. Clin. Invest. 97, 2534–2540.

Chapter 21

The Central Control of Bone Remodeling Paul A. Baldock

Senior Research Officer, Bone Research Program, Garvan Institute of Medical Research, 384 Victoria Street, Sydney NSW 2010, Australia

Susan J. Allison

Postgraduate Scholar, Bone Research Program, Garvan Institute of Medical Research, 384 Victoria Street, Sydney NSW 2010, Australia

Herbert Herzog

Adjunct Professor, Faculty of Medicine, The University of New South Wales, Principal Research Fellow, Head Obesity and Energy Homeostasis Research Group, Director Neurobiology Program, Garvan Institute of Medical Research, 384 Victoria Street, Sydney NSW 2010, Australia

Edith M. Gardiner

Associate Professor, School of Medicine, The University of Queensland, Head, Skeletal Biology Unit, Centre for Diabetes & Endocrine Research, Ground Floor, C Wing, Bldg 1, Princess Alexandra Hospital, Ipswich Road, Brisbane QLD 4102, Australia

V. Interaction between Leptin and Y2-Regulated Bone Antiosteogenic Pathways VI. Concluding Remarks References

I. Introduction II. Actions of Leptin III. Sympathetic Nervous System IV. Neuropeptide Y and the Y Receptors

I. INTRODUCTION

formation, and that both activities require an intact sympathetic nervous system [1, 3, 4]. Leptin pro-resorptive activity appears to act through two distinct mechanisms, one via sympathetic signals that increase RANKL expression in immature osteoblasts and another signaling through the hypothalamic receptor for cocaine amphetamine regulated transcript, CART, which inhibits osteoblastic RANKL expression via an as-yet-undefined mechanism [3]. Importantly, mice lacking the sympathetic system were resistant to ovariectomy-associated bone loss [3]. This finding, together with the potent bone anabolic responses observed with genetic disruption of the leptin and the

Control of bone physiology through the hypothalamopituitary-corticotropic axis has long been recognized and studied, but more recent discoveries have revealed direct central neural mechanisms in addition to the known neuroendocrine pathways. The finding in 2000 that leptin acts through the hypothalamus to inhibit bone formation was rapidly followed by evidence that the hypothalamic neuropeptide Y2 receptor also mediates a distinct but perhaps interacting antiosteogenic pathway [1, 2]. It is now known that leptin regulates bone resorption as well as Dynamics of Bone and Cartilage Metabolism

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Y2-mediated antiosteogenic pathways, emphasizes the potential value of bone-specific therapeutics that target these central control mechanisms for prevention and repair of hypogonadal bone loss. This chapter outlines the current state of understanding and highlights important unresolved issues and future research opportunities in this exciting and rapidly moving area of bone research.

II. ACTIONS OF LEPTIN A. Leptin Role in Energy Homeostasis in Mouse Leptin, the product of the obese gene, is a small polypeptide hormone secreted into the peripheral circulation by adipocytes. Serum levels of leptin, proportional to total body adiposity, are sensed by the leptin receptor in the hypothalamus as part of the energy homeostatic mechanism. Mice with mutations in the obese gene (ob/ob) or its receptor (db/db) display marked obesity accompanied by hyperglycemia, hyperinsulinemia, hypercorticism, and hypogonadism [5–7]. Confirming the potent central action of the hormone, intracerebroventricular (icv) administration of leptin caused a dose-dependent reduction in body weight and food intake in both ob/ob and control mice [6, 8, 9]. Body temperature, locomotor activity, glucose and insulin levels, and body fat mass were also partially corrected after leptin treatment of ob/ob mice [6, 8–10]. Leptin effects in humans and rodents are mediated through receptors in the periphery and the central nervous system. The predominant “short” Ob-Ra form of the leptin receptor is widely expressed in peripheral tissues including kidney, lung, adrenal gland, and choroid plexus [11]. The “long” Ob-Rb receptor variant, generated by alternative splicing [12, 13], is the only isoform that has been shown to mediate signal transduction in vivo [14]. It is most highly expressed in the hypothalamus, particularly in the ventromedial hypothalamic nucleus (VMH) and arcuate nucleus (ARC), which are thought to be the most critical targets for leptin actions [11]. It is the Ob-Rb isoform that is deleted in the db/db obese mouse model [13]. Neuronspecific, but not hepatocyte-specific, disruption of the leptin receptor was associated with obesity, indicating that the nervous system is a primary target for the weight-reducing and neuroendocrine effects of leptin [15].

this was found not to be the case, as cancellous bone volume was markedly increased in both models [1]. This increased bone mass was associated with a marked increase in bone formation, which surpassed the elevation of bone resorption that was also observed. A key role for hypothalamic leptin activity in regulating bone mass was indicated by icv infusion of leptin, which decreased cancellous bone volume in both ob/ob and wild-type mice (Fig. 1) [1]. Consistent with a central mechanism of action, leptin was not detectable in the serum of the icvadministered ob/ob mice. The lack of skeletal effect by circulating leptin was further demonstrated by central icv infusion of leptin to one mouse of a pair of parabiosed ob/ob mice. Although effective cross-circulation was established, correction of the ob/ob skeletal phenotype by a decrease of bone mass occurred only in the recipient, and not the contralateral mouse [4]. In addition, in a separate study using similar duration of leptin treatment, peripheral administration of a 25-fold greater dose of leptin in wild-type mice caused no alteration to cancellous bone volume or bone formation rate [16]. These findings provided strong evidence that, similar to the central control of obesity, the regulation of osteoblast activity on cancellous bone surfaces is mediated by leptin inhibitory actions within the hypothalamus. Subsequent studies implicating

Figure 1

B. Leptin Deficiency and Bone in Mice The hypogonadism and hypercorticism of ob/ob and db/db mice would be expected to result in reduced cancellous bone mass due to elevated bone loss but, unexpectedly,

Central leptin administration reduces cancellous bone mass. Histological sections from vertebrae of 4-month-old ob/ob (A) and wildtype (B) mice following central infusion into the third ventricle with leptin or PBS for 28 days. In both leptin-deficient and wild-type mice, central leptin administration reduced trabecular bone mass and volume. Underlined numbers indicate a statistically significant difference between experimental and control groups of mice (P < 0.05). (Adapted from Ducy et al. Cell 100: 197–207, 2000.)

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the sympathetic nervous system as the downstream mediator of leptin antiosteogenic action are covered later in this chapter. The unfolding story of leptin antiosteogenic control provides exciting new insights into neural mechanisms in skeletal physiology, but there are reports in the literature that suggest additional levels of complexity. For example, leptin-resistant Zucker ( fa/fa) rats, similar to the db/db mouse, carry an inactivating Gln269Pro mutation in the extracellular domain of the Ob-Rb leptin receptor gene [17], but unlike either db/db or ob/ob mice, these rats exhibit reduced femoral bone mineral density (BMD) and calcium content [18], suggesting the possibility of rodent species or strain differences in leptin control of bone. Also, leptin action in cortical bone may be distinct from its antiosteogenic effects in the cancellous compartment, as femoral length and total body BMC are decreased in ob/ob mice and peripheral leptin treatment of immature (4-weekold) ob/ob mice partially restored both parameters [19]. As leptin can stimulate growth plate chondrocyte proliferation and differentiation in vitro [20], it is possible that the ob/ob-associated reductions in bone length and BMC may relate to growth plate rather than osteoblastic effects. A subsequent study also reported lower femur length and bone mineral content, as well as decreased mid-shaft cortical area and thickness in leptin-deficient mice; cortical thickness of lumbar vertebrae was also reduced but, paradoxically, vertebral length and lumbar BMC and BMD were increased [21]. Whether these differences between effects of leptin deficiency on cortical and trabecular compartments or between appendicular and axial skeleton may relate to effects on muscle mass, marrow adipogenesis [21] or growth plate remains to be resolved.

C. Leptin Excess and Bone in Mice The marked increase in bone formation that occurs with ablation of the leptin response in the ob/ob and db/db mice provides important information about the existence of this key inhibitory pathway and potential skeletal consequences of reduced leptin concentration, but does not necessarily predict how bone will respond to elevated serum leptin. In this respect, an initial insight was provided by the lack of change in cancellous bone volume in high-fat-fed wildtype mice despite a 50% increase in body weight and, presumably, a parallel increase in serum leptin [1]. A later study directly testing the effect of raised serum leptin concentration revealed that the magnitude of bone loss achieved with leptin excess was far less than the increase in bone resulting from leptin deficiency [22]. In lypodystrophic A-ZIP/F1 mice, a four-fold reduction in leptin was associated with a doubling of cancellous bone volume associated with an estimated 50% increase in osteoblast activity [1]. By contrast, a transgene-induced increase in serum leptin reduced cancellous bone volume and osteoblast activity by only around 20%, whether the leptin increase was four-fold or 180-fold [22] (Fig. 2). This observation suggested an upper limit to the antiosteogenic potential of leptin, perhaps due to central resistance to the hormone, which is characterized by altered transport of leptin across the blood–brain barrier [23, 24] and reduced signaling from hypothalamic leptin receptors [25]. There are reports that peripheral leptin administration is associated with reduced ovariectomy-induced bone loss in rats [26] and increased bone strength in mice [27]. The clear contrast between these findings and the bone anabolic effects due to leptin reduction in the ob/ob mouse could

Figure 2 Peripheral leptin elevation reduces cancellous bone mass. Vertebrae of 6-month-old models in which transgenic leptin overproduction in liver led to modest (A) and extreme (B) elevation of serum leptin (ng/ml). In addition to severe lipodystrophy (not shown here), cancellous bone volume (BV/TV) was significantly decreased in both models, associated with a reduction in osteoblast activity (BFR/BS) (*P < 0.05). (Adapted from Elefteriou et al. Proc. Nat. Acad. Sci. USA 101: 3258–3263, 2004.)

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be explained if, at elevated levels, positive peripheral leptin effects on bone mass may override those of the central regulatory mechanism. This possibility is discussed further below.

D. Leptin Action in Human Bone 1. Leptin Deficiency and Bone in Humans

The association between leptin deficiency and elevated bone formation has been convincingly demonstrated by genetic studies in mice, but the limited genetic evidence in humans is mixed. There is evidence supporting an osteogenic effect of leptin, as one of four individuals in a family carrying a missense mutation in the leptin gene exhibited very low bone mass despite morbid obesity, but a high degree of consanguinity and multiple endocrine defects cloud interpretation of this observation [28]. In a different study that also supports an osteogenic leptin effect, administration of leptin for one year to a young girl with a leptin mutation was associated with loss of body fat and an increase in whole-body BMD [29]. These findings are inconsistent with the indirect evidence of leptin antiosteogenic function provided by an anecdotal report of advanced bone age in a single leptin-deficient child and more documented evidence of a similar effect in prepubertal patients affected by congenital generalized lipodystrophy [22]. Also supporting leptin antiosteogenic action, a Gln223Arg polymorphism in the human leptin receptor gene, while not associated with altered body weight or serum leptin, was associated with higher lumbar BMD in young men, although the effect was dependent upon genotype at the estrogen receptor α locus and linkage disequilibrium has not been excluded [30]. The encoded amino acid change, which is in the extracellular domain of the leptin receptor, has previously been associated with reduced leptin-binding affinity in carriers of the allele [31], which would be consistent with the observed increase in peak BMD in the study by Koh and colleagues. It is therefore not possible to deduce the physiological role(s) of leptin in human bone biology based on human genetic evidence. Indirect clinical evidence suggests that if there is an association between leptin deficiency and high bone mass in humans, it may not be an overriding factor in some circumstances. BMD was low after self-imposed calorie restriction in anorexia nervosa, which can lead to extremely low levels of the nutritionally dependent hormones leptin and IGF-1 and amenorrhea, suggesting that, unlike the ob/ob mice, leptin deficiency did not compensate for the skeletal consequences of low IGF-1 levels and hypogonadism in these patients [32, 33]. Reinforcing the potential importance of IGF-1 in this context, leptin treatment in

patients with hypothalamic amenorrhea due to strenuous exercise or low body weight resulted in increased levels of both IGF-1 and bone formation markers [34]. Such studies involving reduced nutritional intake, excessive exercise and disrupted menses are not, however, amenable to simple interpretation in terms of direct leptin effects on bone vs. indirect effects acting through altered reproductive and other neuroendocrine function. 2. Normal to Excess Leptin and Bone in Humans

Initial correlations between bone mass and body weight or fat mass [35] were attributed either to variations in mechanical load or estrogen production related to fat tissue mass, but after the discovery of leptin, this additional factor was also investigated. In cross-sectional studies, serum leptin levels and BMD are directly correlated in pre- and postmenopausal females by some investigators [36] but not others [37]. Furthermore, leptin and whole body bone area, as well as change in bone area over time, were correlated in one study of pubertal girls [38], a positive association between leptin level and bone mineral content was observed in a group of healthy nonobese women [39], and a weak correlation was observed in postmenopausal osteoporotic women but not the controls [40]. This uncertain relationship between leptin and bone may be genderspecific, as studies in males have reported either no association [36] or a negative association [41] between leptin and BMD. The possibility that some of these associations (or lack thereof) may relate to measurement artifacts associated with the particular technologies for measuring bone density employed in each study is a subject of ongoing discussion [35]. In this context, it is interesting that at least one study that did detect a positive association between leptin and BMD found no association between leptin and biochemical markers of bone resorption or formation, and concluded that leptin is not likely to play a significant direct role in regulating bone cell activity [42]. This interpretation is challenged, however, by direct examinations of bone cell responses to leptin in vitro. 3. Studies of Bone Cell Responses to Leptin IN VITRO

Evidence for presence of the signal-transducing Ob-Rb leptin receptor or leptin-binding sites has been detected on ossifying fetal cartilage [43], immortalized marrow stromal cells [44], chondrocytes [19, 20, 27, 45], primary osteoblasts [19, 27, 46–48], and some but not all osteosarcoma cell lines [47]. Adding further to the possibility of peripheral actions of leptin is evidence of leptin production by primary human and rat osteoblast cultures [49, 50].

Chapter 21 The Central Control of Bone Remodeling

A number of in vitro functional studies support direct leptin effects on chondrocytes and osteoblasts. Leptin has been observed to stimulate proliferation and differentiation of cultured growth plate chondrocytes [20] and to induce osteoblast differentiation and suppress adipogenesis in a human stromal cell line [44]. The hormone also has direct stimulatory effects on cell proliferation, differentiation, and function in human and rat osteoblastic cultures [27, 51]. In addition, leptin can inhibit osteoclastogenesis in vitro [26, 52]. One notable exception to these studies detected no osteoblastic expression of leptin or the leptin receptor long form, and no effect of leptin treatment on formation of mineralized nodules in primary mouse osteoblastic cultures [1]. The majority of the in vitro evidence, however, supports peripheral actions of leptin in bone biology. It therefore remains possible that, particularly at high leptin concentrations, competing interactions of opposing peripheral osteogenic and central antiosteogenic actions of leptin may in part explain the inconsistencies in the literature.

III. SYMPATHETIC NERVOUS SYSTEM A. Evidence for Sympathetic Control of Bone Remodeling by Leptin The sympathetic nervous system (SNS) is known to mediate some of the effects of leptin on body composition, with decreased sympathetic tone noted in leptin-deficient ob/ob mice [53]. Based on those observations, Takeda and colleagues investigated whether the central antiosteogenic actions of leptin might also be mediated via the sympathetic nervous system and found that dopamine β-hydroxylase (DBH)-deficient mice, which are unable to produce epinephrine and norepinephrine, the catecholamine ligands for adrenergic receptors, also have high bone mass [4]. Importantly, icv infusion of leptin failed to reduce the bone mass of DBH mice, indicating a requirement for a functional autonomic nervous system for leptin antiosteogenic actions, and identifying the sympathetic system as a neuronal mediator of leptin effects on bone [4]. This conclusion is supported by studies using β-adrenergic receptor agonists and antagonists. Administration of the β-agonist isoproterenol restored sympathetic activity in ob/ob mice and reduced bone mass in both wild-type and ob/ob mice without affecting body weight (Fig. 3) [4]. Conversely, administration of the nonselective β-adrenergic receptor antagonist propranolol increased bone mass in wild-type mice and prevented bone loss following ovariectomy, demonstrating that modulation of SNS activity affects bone remodeling (Fig. 4) [4]. Mice lacking

365 functional β2-adrenergic receptors (Adrb2-/-) have normal fat mass and fertility, accompanied by an even more marked high bone mass phenotype than ob/ob mice or wild-type mice receiving β-receptor antagonists [3]. As would be expected, icv leptin infusion into Adrb2-/- mice was unable to reduce bone mass, further demonstrating that intact and functional sympathetic nervous system signaling is required for the antiosteogenic actions of leptin [3]. Furthermore, transplantation of nonadherent bone marrow cells from wild-type mice normalized bone formation in Adrb2-/- mice, whereas the reciprocal transplant led to increased bone formation, consistent with sympathetic signaling at least in part via cells of the osteoblast lineage to control bone mass (Fig. 5) [3]. Involvement of β-adrenergic signaling in bone formation has been addressed with mixed results in rodent surgical models. In one study, treatment with propanolol increased bone strength in intact rats and enhanced bone formation in the repair of surgically introduced bone defects in rats [54]. Not all reports have been supportive of an autonomic antiosteogenic circuit, however, as administration of the specific β2-AR agonist clenbuterol prevented a reduction in mineralization of bone caused by sectioning of the sciatic nerve in rats [55]. However, in this study, the anabolic effect of clenbuterol in skeletal muscle was proportional to the inhibition of bone loss, implying an indirect effect through maintenance of loading by skeletal muscle. Consistent with the in vivo evidence for sympathetic control of bone formation, β-adrenergic receptors (β-AR) have been identified on osteoblasts and in osteoblast-like cell lines. β1- and β2-adrenergic receptors are expressed at different levels in various human osteoblast-like cell lines [56] and β2-adrenergic receptors have been identified on rat osteoblast-like cells, and in mouse primary osteoblast cultures [4, 57], suggesting that sympathetic activity might control bone formation through direct modulation of osteoblast function. Supporting this, administration of the specific β2-AR agonist, formoterol, induced cAMP levels and expression of the immediate early gene c-fos in the SaOS-2 human osteosarcoma cell line [56]. Likewise, c-fos expression was inhibited by administration of a specific β2-AR antagonist, demonstrating that β2-adrenergic receptors in osteoblasts are indeed functionally coupled to intracellular signaling pathways. Furthermore, in vitro studies have confirmed the ability of adrenergic signaling to alter bone remodeling. Administration of norepinephrine stimulated bone resorption in neonatal mouse calvariae in organ culture [57] and treatment of the MC3T3-E1 preosteoblastic mouse cell line with epinephrine stimulated expression of osteoclast differentiation factor [58], suggesting that β-adrenergic stimulation of resorption may occur via an indirect osteoblast-mediated pathway.

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Pharmacological β-adrenergic agonism reduces cancellous bone mass. Proximal tibia and vertebrae from 1-month-old ob/ob (A) and wild-type (C) mice following 6 week treatment with the β-adrenergic agonist isoproterenol (3 mg/kg/day for ob/ob and 30 mg/kg/day for wild-type). Cancellous bone volume was significantly reduced in both groups (B, D) following β-agonism associated with a reduction in osteoblast activity (BFR/BS) and number (N.Ob./B.Pm.) (*P < 0.01). (Adapted from Takeda et al. Cell 111: 305–317, 2002.)

Figure 3

Figure 4 Pharmacological β-adrenergic antagonism increases cancellous bone mass. Tibia and vertebrae from 1-month-old wild-type mice following 5-week treatment with the β-adrenergic antagonist propranolol (orally, 0.4 mg/day). Cancellous bone volume (A, BV/TV) was significantly increased following β-antagonism associated with (B) an increase in osteoblast activity (BFR/BS) and number (N.Ob./B.Pm.) (*P < 0.01). (Adapted from Takeda et al. Cell 111: 305–317, 2002.)

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Figure 5 Reciprocal bone marrow transplants confirm sympathetic control of bone mass. Vertebrae from 6-month-old recipient mice, collected 4 months after lethal irradiation followed by transplantation with nonadherent marrow cells from donor mice. Transplantation of bone marrow cells from wild-type mice into Adrb2-/mice, which lack β2 adrenergic receptor, reduced bone volume (BV/TV) and osteoblast activity (BFR) and surface (Ob.S/BS) to normal levels and increased urinary elimination of deoxypryidinoline (Dpd/creat.), an indicator of osteoclast function. Conversely, transplantation of Adrb2-/- marrow cells increased bone formation and decreased bone resorption in wild-type recipients (*P < 0.05). (Adapted from Elefteriou et al. Nature 434: 514–520, 2005.)

In addition to increased bone formation, Adrb2-/- mice also exhibited significantly decreased bone resorption and, importantly, this conferred a resistance to gonadectomyinduced bone loss. Furthermore, as mentioned in the previous section, icv leptin treatment failed to normalize the baseline Adrb2-/- phenotype [3]. Together with the in vitro evidence from the previous section, these observations indicate that leptin also regulates bone resorption via sympathetic signaling. Transplantation of wild-type bone marrow cells restored normal levels of bone resorption in Adrb2-/- mice, and the reciprocal transplant reduced resorption parameters, consistent with sympathetic signaling via cells of the osteoblast lineage to control bone resorption [3]. In addition, the β-agonist isoproterenol did not increase osteoclast formation in co-cultures of wild-type bone marrow macrophages with Adrb2-/- osteoblasts, although an increase did occur in co-cultures with wild-type osteoblasts. Also, isoproterenol increased expression of the osteoclast differentiation factor RANKL in cultured wild-type but not Adrb2-/- osteoblasts, via a mechanism involving the osteoblast-specific transcription factor ATF4. Further evidence indicated that sympathetic and PTH signaling target different subsets of osteoblasts, with sympathetic control acting through undifferentiated osteoblasts expressing β2-AR and ATF-4, and PTH acting

by an ATF-4-independent mechanism later in osteoblastic differentiation [3].

B. Clinical Relevance of b-adrenergic Control of Bone Beta-blocker usage to treat cardiovascular diseases is prevalent, but effects on skeletal mass and strength had not been widely investigated prior to the evidence from mouse studies cited above. The effects of β-adrenergic antagonists on bone turnover, bone mineral density (BMD), and fracture risk have now, however, been assessed in human population-based studies. β-blocker use was associated with a reduction in fracture risk and increased BMD at the hip and forearm in women over 50 years of age [59], and reduced fracture risk in women and men between 30 and 79 years of age [60], consistent with the rodent model data. However, a third observational study contradicted these findings, with β-blocker usage associated with a three-fold increase in fracture risk and reduced serum osteocalcin, an osteoblastic marker of bone formation, in perimenopausal women [61]. The difference in this latter finding may relate to variations in study design. Thus, there is some epidemiological evidence to support the hypothesis

368 that β-adrenergic signaling is involved in bone metabolism; however, placebo-controlled randomized clinical trials are necessary to more effectively assess potential therapeutic benefit from bone-specific β-adrenergic blockade in hypogonadal and age-related osteoporosis and other conditions that could benefit from stimulation of bone formation.

C. Role of CART and Melanocortin Receptors in Bone Remodeling The finding that Adrb2-/- mice were resistant to gonadectomy-induced bone loss due to lack of a bone resorption response was somewhat surprising, as this was a fundamental physiological difference from the elevated bone resorption observed in the hypogonadic ob/ob mouse model, for which gonadectomized Adrb2-/- mice had until then been considered a phenocopy [3]. This discrepancy between the two models appears to relate to another neuropeptide, CART (cocaine amphetamine-regulated transcript), a known hypothalamic target of positive leptin regulation that is low in ob/ob mice [1] but normal in Adrb2-/- [3]. Cart-/- mice displayed a low bone mass phenotype associated with increased bone resorption (Fig. 6), and also exhibited an enhanced resorptive response to icv leptin infusion, demonstrating both that

Figure 6 Cart gene deletion associated with decreased cancellous bone mass. Vertebrae from 6-month-old Cart-/- mice, displaying reduced cancellous bone volume (BV/TV) associated with increased osteoclast perimeter (Ob.Nb/BPm and Oc.S/BS) and bone resorption (Dpd/creat.), with no change in bone formation (BFR) (*P < 0.05). (Adapted from Elefteriou et al. Nature 434: 514–520, 2005.)

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leptin-mediated sympathetic regulation of bone mass functions in the absence of CART, and that CART is an inhibitor of bone resorption [3]. CART transcripts were not detected in bone cells and in vitro analyses of Cart-/bone cells yielded no evidence of cell-autonomous CART effects, but RANKL expression was up-regulated in Cart-/- bones, implicating this osteoblastic regulator of osteoclastogenesis as a mechanistic element of the Cart-/phenotype. Earlier work from this group using chemical ablation of different regions of the hypothalamus indicated that the ventromedial hypothalamic nucleus (VMH) mediates the leptin antiosteogenic actions, with the arcuate nucleus (ARC) not involved in that regulatory circuit [4]. These more recent data implicating CART in central control of bone resorption that is independent of the sympathetic nervous system are consistent with those findings, as CART neurons are enriched in the ARC, as are neurons that produce POMC (pro-opiomelanocortin), the precursor of α-melanocyte-stimulating hormone (α-MSH) [62, 63]. After release from ARC neurons, α-MSH binds to the melanocortin-4 and -5 receptors (MC4R, MC5-R) on target neurons, thereby activating them and leading to anorexic effects on energy homeostasis [64, 65]. The Ay yellow agouti protein is a high-affinity competitive antagonist for MC4R [66], and Ay/a mice are sensitive to leptin antiosteogenic function, indicating that MC4R is not directly involved [4]. Mc4r-/- mice exhibit increased hypothalamic Cart expression and late-onset high bone mass that was associated with markedly reduced osteoclast number but normal bone formation parameters [3]. Thus, the Cart and Mc4r expression changes in Mc4r-/mice are mutually consistent and compatible with the reduction in bone resorption. In this context, it is interesting to note that markers of bone resorption are decreased in MC4R-deficient patients, lending explanation to the increased bone density previously observed in these patients [67]. Together, these observations support a model in which neural control of bone resorption may be regulated by two distinct neural mediators. Tonic inhibition by leptinsensitive VMH neurons of sympathetic signaling to immature osteoblasts via β2-adrenergic receptors induces osteoclastic differentiation, whereas CART and MC4R on arcuate neurons inhibit resorptive activity. Both these pathways appear to act on the osteoblastic cell lineage by modifying expression of the osteoclast differentiation factor, RANKL. However, the precise mechanism(s) by which CART and MC4R act is as yet unknown. Another unknown in this scenario is how the CART and MC4Rmediated circuits may relate to a tonic antiosteogenic pathway that involves neuropeptide Y responsive neurons

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in the ARC. What is known about the skeletal roles of these Y receptor neurons is detailed in the following section.

IV. NEUROPEPTIDE Y AND THE Y RECEPTORS A. NPY System Ligands The neuropeptide Y (NPY) system involves the actions of three ligands, neuropeptide Y, peptide YY, and pancreatic polypeptide. These ligands are produced in specific distributions. NPY is produced by neurons of the central and peripheral nervous systems, present in many sympathetic nerve fibers, especially around blood vessels, and also in some parasympathetic nerves. NPY is a wellcharacterized vasoconstrictor in these neurons, in addition to enhancing the action of other pressor agents [68–70]. Furthermore, NPY-ergic neurons are abundant in the brain, with high levels in the hypothalamic arcuate nucleus (ARC) and ventromedial hypothalamus (VMH) [71]. Central NPY action is associated with the regulation of food intake, cardiac and respiratory activity, and the release of pituitary hormones [71, 72]. Peptide YY (PYY) is produced primarily from the endocrine L cells of the gastrointestinal tract with some expression in the pancreatic islets, and its function is related to satiety control and gastrointestinal regulation [73]. PYY expression has also been reported in brainstem neurons, although the functional significance of this localization is unknown [74]. Pancreatic polypeptide (PP) is produced by endocrine islet cells of the pancreas and regulates endocrine functions of the pancreas as well as satiety centrally [75]; to date, production of this neuropeptide has not been detected in peripheral or central neural tissue [74].

cAMP synthesis. Y1 receptors can also couple to phospholipase C to provoke release of Ca2+ from intracellular stores [79, 80]. Because of the multiplicity of Y receptors and the range of their physiological targets, receptorspecific antagonists have been developed with the goal of dissociating the various effects of NPY, with reasonable success for Y1 and Y5, but limited success for Y2 and none for Y4 [81]. To date, problems with solubility, toxicity, and inability to cross the blood–brain barrier have limited the potential utility of even the most promising nonpeptidic ligands for in vivo research and possible clinical application. Fortunately, the recent development of Y receptor knockout mouse models has enabled significant research progress, as will be evident below. Y1 and Y2 receptors are located at the post- and presynaptic terminals of the neuroeffector junction, respectively. They are both abundant in central tissue, with hypothalamic expression of Y1 predominantly in the paraventricular nucleus (PVN) and Y2 in the arcuate nucleus (ARC). Importantly, Y2 and leptin receptors are co-expressed in the ARC, which lies outside the blood–brain barrier and is therefore exposed to the peripheral circulation [69]. The Y2 receptor is predominantly presynaptic and inhibitory, and in some instances thought to act as an autoreceptor [82]. Activation of Y2 receptors modulates neurotransmitter synthesis and release, as well as production of neuropeptides. In the context of this chapter, it is particularly important that Y2 activation inhibits NPY production by Y2-expressing (and leptin receptor-expressing) NPY-ergic neurons [82]. Y4 receptor is widely distributed in central and peripheral tissues, with abundant expression in specific brain stem nuclei like the area postrema, also located outside the blood–brain barrier [69].

C. NPY in Bone Tissue B. The Y Receptors The neuropeptide Y system mediates its effects through activation of at least five different receptors: Y1, Y2, Y4, and Y5 receptors, which are present in humans and rodents, and y6, which is a pseudogene in humans but generates a functional receptor in the mouse [76, 77]. While each individual Y receptor binds NPY and PYY with equal affinities, the outright binding affinity characteristic of each receptor varies, with stronger binding for Y2 and Y1 compared to the other receptors. In contrast, the Y4 receptor has a low affinity for NPY and PYY, but the highest affinity of all the Y receptors for PP [78]. The Y receptors are seven-transmembrane receptors coupled to inhibitory G proteins, thereby mediating inhibition of

There is evidence that NPY-positive autonomic nerves are present in healthy bone tissue, particularly in association with blood vessel walls, suggestive of a role in modulation of vascular tone rather than direct effects on bone cells [83–86]. However, NPY immunoreactive nerve fibers were also detected in the synovium and bone marrow of ankle joints in arthritic rats [87]. In addition, sympathetic denervation has been observed to reduce NPY immunoreactive nerves in the periosteum. A functional study indicated the presence of NPY-responsive receptors on osteoblastic cells, with NPY treatment inhibiting the cAMP response in osteoblastic cell lines that had also been treated with PTH or norepinephrine, although NPY treatment alone did not alter basal cAMP levels [88, 89]. These findings are consistent with the known inhibitory

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action of NPY on the cAMP response in other systems. There have been a few contradictory reports about NPY receptor transcripts in bone, with positive results reported for human periosteum-derived osteoblastic cells and human osteosarcoma cell lines [90], and transcripts encoding an isoform of the Y1 receptor detected in bone marrow cells and hematopoietic cell lines [91]. Others, however, have failed to detect Y receptor transcripts in primary osteoblastic cultures using RT-PCR detection [2]. It is essential that this question be resolved, given the accumulating evidence that Y receptors are involved in the regulation of bone formation, as described below.

D. Y2 Receptor Deletion Effects on Bone Because of the presence of both Y2 and leptin receptors on neurons of the arcuate nucleus [92, 93] and the emerging evidence that central leptin signaling inhibits bone

formation, a potential role for Y2 in controlling bone mass was investigated. Cancellous bone volume in Y2 receptor germline knockout mice (Y2-/-) of both genders was doubled, with increases in trabecular number and thickness [2]. Central Y2 receptors were critical for this phenotype, as a doubling of trabecular bone volume was observed only 5 weeks after conditional deletion of hypothalamic Y2 receptors from adult mice (Fig. 7). This increase in bone volume was primarily the result of greater osteoblastic activity, with the only detectable change in bone resorption a modest elevation in osteoclast number. Changes in circulating levels of insulin-like growth factor-1 (IGF-1), free T4, and testosterone were unchanged, ruling out these hypothalamo-pituitary effectors as potential mediators of the Y2-dependent bone regulatory signal [2]. Corticosterone levels were modestly but not significantly reduced [2], plasma calcium and leptin levels were unchanged, and there was no difference in Y2-/- body weight despite evidence that Y2 receptors on leptin-responsive arcuate neurons do

Figure 7 Hypothalamic Y2 receptor deletion increased cancellous bone formation. Histological sections from distal femurs of 4-month-old mice after conditional deletion of hypothalamic Y2 receptor at 3 months of age (A). Y2 floxed mice (Y2lox/lox) were injected with CRE (CRE-Y2lox/lox) or GFP (GFP-Y2lox/lox) expressing adeno-associated virus in the hypothalamus at 11 weeks of age. Five weeks later, trabecular bone volume was significantly increased (B), associated with greater bone formation rate (C) (**P < 0.001, *P < 0.05). (Adapted from Baldock et al. J Clin Invest 109: 915–921, 2002.)

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modify whole-body energy homeostasis [94]. Much of the evidence therefore strongly suggested that the hypothalamic Y2-dependent antiosteogenic circuit may act through a neural mechanism.

E. Pancreatic Polypeptide and Y4 Receptor Two additional changes in Y2-/- mice that were of interest were an elevation of plasma PP concentration and increased hypothalamic NPY expression [95]. Elevation of PP in a transgenic mouse model had no effect on cortical or cancellous bone structure, effectively ruling out the increase in PP as a sole source of the Y2-/- -associated increase in bone formation [95]. This did not, however, exclude the possibility that Y4 activation in response to significantly elevated hypothalamic NPY levels in the Y2 KO mice might have stimulated osteoblast activity. As with Y2 deletion, germline Y4 receptor knockout led to increased plasma PP concentration, accompanied by a lean phenotype with decreased food intake [96], but, as for the PP transgenic mice, there was no discernible bone phenotype in Y4-/- mice [2]. Importantly, Y4 deletion did not alter hypothalamic expression of neurotransmitters involved in energy homeostasis such as NPY [95], thus providing no insights into the skeletal importance of the elevated hypothalamic NPY seen in the Y2-/- model. To address this, the bones of Y2Y4 double-mutant mice were examined, in which hypothalamic NPY expression was markedly elevated [95].

F. Y2Y4 Receptor Double Deletion Remarkably, coordinate germline deletion of both Y2 and Y4 receptors resulted in a synergistic anabolic effect, with a threefold increase in trabecular bone volume, compared with the twofold increase in Y2-/- mice (Fig. 8) [95]. This anabolic response was associated with increased bone formation rate and mineral apposition rate, as in the Y2-/- model. Importantly, this synergism between Y2 and Y4 receptors in the control of bone was only evident in male mice, although both male and female Y2-/-Y4-/mice were very lean, with body weight and white adipose tissue mass reduced compared to either Y2-/- or Y4-/- mice. Notably, plasma leptin was significantly reduced by about 60% in male mice, but not in female mice [95], which suggested that the added increase in cancellous bone volume in the Y2Y4 KO male mice might be an additive effect of both Y2 and leptin-mediated pathways. Consistent with this interpretation were the reduced, mid-diaphyseal cortical bone area and thickness in the Y2Y4 double

Figure 8

Deletion of Y2 and Y4 receptors associated with synergistic increase in cancellous bone volume. Histological sections from distal femoral metaphysis of male 4-month-old wild-type (A), Y2-/- (B), Y4-/- (C), and Y2-/-Y4-/- (D) mice. Greater trabecular bone volume was evident in Y2-/-Y4-/- compared to Y2-/-, with Y4-/- not different from wild-type (E). (*P < 0.05 vs. wild-type, # P < 0.05 vs. Y2-/-). (Adapted from Sainsbury et al. Mol Cell Biol 23: 5225–5233, 2003.)

mutants and the more marked increase in NPY transcript levels in the ARC of Y2-/-Y4-/- mice [95]. These data were consistent with a Y4-mediated pathway that is separate from but may interact with the proposed Y2 neural pathway and may act via a humoral, perhaps leptin-mediated, mechanism. This possibility is examined in greater detail in the next section.

V. INTERACTION BETWEEN LEPTIN AND Y2-REGULATED BONE ANTIOSTEOGENIC PATHWAYS The actions of leptin and the Y receptor ligand NPY are intimately linked in the hypothalamus. Deficiency of leptin or its receptor in the ob/ob and db/db mouse mutant strains, respectively, results in strongly elevated hypothalamic NPY expression. This is due to lack of leptin-induced inhibition of NPY expression and secretion, and has been shown to contribute to the massive obesity, hypercorticism, and reproductive defects characteristic of ob/ob mice [97]. NPY ablation reverses these defects, supporting a role for

372 NPY as a downstream mediator of leptin in energy homeostatic control [98]. As mentioned previously, Y2 and leptin receptors are co-expressed on NPY-ergic neurons in the arcuate nucleus, and Y2 receptor is thought to act as an autoreceptor, modulating the expression and secretion of NPY [82]. As deficiency in either the leptin or Y2 receptor results in increased NPY expression, the two receptors appear to share a common signaling pathway to regulate energy homeostasis, and could therefore also share a common centrally mediated pathway to regulate bone formation. The similarity between the high bone mass and elevated bone formation phenotypes of ob/ob and Y2-/- models, and the elevation of hypothalamic NPY expression in both models [2, 97], suggests that the high bone mass phenotype may be due to the increased NPY expression. However, continuous icv administration of NPY into wild-type mice for 28 days actually decreased bone volume, suggesting that leptin and NPY might in fact use different pathways to control bone mass and body weight [1]. One issue that was not addressed in that study, however, is that icv NPY infusion would cause an increase in total fat mass, even in pair-fed animals. This increased adiposity would consequently lead to increased leptin levels, which could have caused the antiosteogenic effects that were observed. The chemical lesion studies that support the model in which neural regulation of bone resorption may be by two distinct mediators, as mentioned earlier in this chapter, also demonstrated that distinct neuron populations are responsible for the hypothalamic control of energy homeostasis and bone mass by leptin [4]. Administration of monosodium glutamate (MSG) to ablate NPY-synthesizing neurons and arcuate nucleus (ARC) structures did not affect bone mass or leptin antiosteogenic function. Intracerebroventricular infusion of leptin into MSG-ablated mice did, however, fail to reduce body weight, indicating that, although NPY-synthesizing neurons are dispensable for leptin action on bone, they are required for leptin action on body weight. In contrast, gold thioglucose-sensitive neurons in the ventromedial hypothalamic nuclei (VMH) were required for leptin antiosteogenic function, as destruction of these neurons led to increased bone formation and was not reversed by icv infusion of leptin [4]. Therefore, it appears that the control of antiosteogenic and anorexigenic networks by leptin differs, as the ARC neuronal population responsible for regulating anorexic function is not necessary for leptin effects on bone, which instead appear to act via the neuronal populations present in the VMH [4]. Evidence for interaction between these distinct leptinand Y2-sensitive antiosteogenic pathways was sought in Y2-/-ob-/- double knockout mice [99]. Interestingly, bone

PAUL A. BALDOCK ET AL .

formation of the ob/ob model was not further enhanced by Y2 receptor deletion, although cancellous bone volume in both the Y2-/-ob-/- and ob/ob models was lower than that of Y2-/- mice due to increased bone resorption associated with the hypogonadal status of the leptin-deficient models (Fig. 9). Importantly, the levels of osteoblast activity were comparably elevated in all three models, indicating that Y2 receptor deletion did not convey additional osteoblastic stimulation in the absence of leptin, and thus suggesting that these models may share a common pathway to regulate osteoblast activity. In the presence of elevated hypothalamic NPY, however, a different result was obtained. Recombinant adeno-associated viral vector expressing NPY under the control of a neuron-specific promoter (AAV-NPY) was bilaterally injected into the hypothalamus of wild-type and Y2-/- mice, dramatically increasing hypothalamic NPY [99]. The resultant stimulation of the orexigenic pathways resulted in pronounced feeding, which doubled body weight and markedly raised plasma leptin levels in both genotypes in 3 weeks. Consistent with previous observations, NPY overexpression and leptin overproduction was associated with significantly reduced osteoblast activity in both Y2-/- and wild-type groups (Fig. 10). Importantly, however, the Y2-/- osteoblast activity remained twice that of wild-type mice, despite the elevated circulating leptin level and the overall reduction in bone formation in both mouse genotypes, indicating that action of the Y2-mediated pathway was independent of the leptin/NPY antiosteogenic pathway. The fact that the

Figure 9

Presence of leptin mutation associated with lower cancellous bone formation and higher resorption compared to Y2-deficient mice. Histomorphometric assessment of male 4-month-old wild-type, Y2-/-, ob/ob, and Y2-/-/ob mice. Cancellous bone volume (BV/TV) in ob/ob and Y2-/-/ob mice was greater than wild-type but less than Y2-/- (A). This difference was associated with greater osteoclast surface in the hypogonadal leptin-deficient ob/ob and Y2-/-/ob mice (C), rather than a difference in mineral apposition rate, which was comparably elevated relative to wild-type in all three mutant models (B). aP < 0.05 vs. wild-type, bP < 0.05 vs. ob/ob, cP < 0.05 vs. Y2/ob. (Adapted from Baldock et al. J Bone Miner Res 2005, 20, 1851–1857.)

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Figure 10

Y2-associated anabolic activity maintained despite reduced bone formation accompanying hypothalamic NPY overproduction. Injection of either control or NPY-expressing adeno-associated virus (empty or NPY) into the hypothalamus resulted in a rapid increase in body weight (A) and serum leptin (B) in both wild-type (wt) and Y2-/- mice. NPY overproduction in wild-type mice led to a marked reduction in osteoblast activity, evident as decreases in osteoid width and volume (C, D), with no change in osteoblast number (E). Osteoblast activity in Y2-/- mice was also reduced in association with NPY overproduction, but osteoid parameters remained significantly elevated compared to the wild-type NPY-overproducing mice, as in empty controls. aP < 0.05 vs. wt-empty, bP < 0.05 vs. Y2-/--empty, cP < 0.05 wt-NPY vs. Y2-/−-NPY. (Adapted from Baldock et al. J. Bone. Miner. Res. 2005, in press.)

Y2-associated osteoblastic advantage was not evident in the Y2-/-/ob bones suggests the possibility that a permissive level of leptin may be necessary for detection of the Y2 pathway activity, or else that the hypothalamic Y2- and leptin-mediated antiosteogenic pathways may share a common feedback mechanism.

VI. CONCLUDING REMARKS The identification of nerve fibers within bone first led to the proposal that the regulation of bone remodeling may be influenced by neuronal mechanisms. The presence of neurotransmitters and neuropeptides and their receptors was subsequently revealed by immunohistochemical techniques, indicating the the neuro-osteogenic control of bone was likely to occur by direct mechanisms [100]. Indeed, many of the neurotransmitters and other neuropeptides present in bone, including vasoactive intestinal peptide (VIP), calcitonin gene-related protein (CGRP), glutamate, and substance P (SP) have been demonstrated to have direct effects on osteoblast and osteoclast cells in culture [101]. However, the first clues about the central control mechanisms came with the discoveries that leptin acts through a hypothalamic relay to inhibit bone formation via the sympathetic nervous system, and hypothalamic Y receptors mediate a distinct but possibly interacting antiosteogenic pathway. Experimental evidence since then has implicated other key hypothalamic neuropeptides in the regulation of bone growth, formation and resorption, which include NPY, CART, and α-MSH. There is no doubt that this list will expand rapidly as investigators strive to gain greater understanding of this recently discovered

aspect of bone physiology, but other necessary breakthroughs must come with better knowledge of the neural circuits between brain and bone, the bone cell responses to local neurotransmitter and neuropeptide release, and the extent to which interactions occur between neural, endocrine, paracrine, and autocrine regulatory mechanisms to integrate the skeletal response to physiological demands. In practical terms, greater understanding of the actions and controls on these potent central antiosteogenic and proresorptive pathways will be invaluable in the development of anabolic therapies to be used in the treatment of postmenopausal osteoporosis, other osteopenic, and orthopaedic conditions.

References 1. Ducy, P., Amling, M., Takeda, S., Priemel, M., Schilling, A. F., Beil, F. T., Shen, J., Vinson, C., Rueger, J. M., and Karsenty, G. (2000). Leptin inhibits bone formation through a hypothalamic relay: a central control of bone mass. Cell 100, 197–207. 2. Baldock, P. A., Sainsbury, A., Couzens, M., Enriquez, R. F., Thomas, G. P., Gardiner, E. M., and Herzog, H. (2002). Hypothalamic Y2 receptors regulate bone formation. J. Clin. Invest. 109, 915–921. 3. Elefteriou, F., Ahn, J. D., Takeda, S., Starbuck, M., Yang, X., Liu, X., Kondo, H., Richards, W. G., Bannon, T. W., Noda, M., Clement, K., Vaisse, C., and Karsenty, G. (2005). Leptin regulation of bone resorption by the sympathetic nervous system and CART. Nature 434, 514–520. 4. Takeda, S., Elefteriou, F., Levasseur, R., Liu, X., Zhao, L., Parker, K. L., Armstrong, D., Ducy, P., and Karsenty, G. (2002). Leptin regulates bone formation via the sympathetic nervous system. Cell 111, 305–317. 5. Tartaglia, L. A., Dembski, M., Weng, X., Deng, N., Culpepper, J., Devos, R., Richards, G. J., Campfield, L. A., Clark, F. T., Deeds, J., Muir, C., Sanker, S., Moriarty, A., Moore, K. J., Smutko, J. S., Mays, G. G., Wool, E. A., Monroe, C. A., and Tepper, R. I. (1995).

374

6.

7. 8.

9.

10.

11.

12.

13.

14.

15.

16.

17.

18.

19.

20. 21.

22.

23.

Identification and expression cloning of a leptin receptor, OB-R. Cell 83, 1263–1271. Halaas, J. L., Gajiwala, K. S., Maffei, M., Cohen, S. L., Chait, B. T., Rabinowitz, D., Lallone, R. L., Burley, S. K., and Friedman, J. M. (1995). Weight-reducing effects of the plasma protein encoded by the obese gene. Science 269, 543–546. Friedman, J. M., and Halaas, J. L. (1998). Leptin and the regulation of body weight in mammals. Nature 395, 763–770. Campfield, L. A., Smith, F. J., Guisez, Y., Devos, R., and Burn, P. (1995). Recombinant mouse OB protein: evidence for a peripheral signal linking adiposity and central neural networks. Science 269, 546–549. Pelleymounter, M. A., Cullen, M. J., Baker, M. B., Hecht, R., Winters, D., Boone, T., and Collins, F. (1995). Effects of the obese gene product on body weight regulation in ob/ob mice. Science 269, 540–543. Chehab, F. F., Lim, M. E., and Lu, R. (1996). Correction of the sterility defect in homozygous obese female mice by treatment with the human recombinant leptin. Nat. Genet. 12, 318–320. Fei, H., Okano, H. J., Li, C., Lee, G. H., Zhao, C., Darnell, R., and Friedman, J. M. (1997). Anatomic localization of alternatively spliced leptin receptors (Ob-R) in mouse brain and other tissues. Proc. Natl Acad. Sci. USA 94, 7001–7005. Lee, G., Proenca, R., Montez, J. M., Carroll, K. M., Darvishzadeh, J. G., Lee, J. I., and Friedman, J. M. (1996). Abnormal splicing of the leptin receptor in diabetic mice. Nature 379, 632–635. Chen, H., Charlat, O., Tartaglia, L. A., Woolf, E. A., Weng, X., Ellis, S. J., Lakey, N. D., Culpepper, J., Moore, K. J., Breitbart, R. E., Duyk, G. M., Tepper, R. I., and Morgenstern, J. P. (1996). Evidence that the diabetes gene encodes the leptin receptor: identification of a mutation in the leptin receptor gene in db/db mice. Cell 84, 491–495. Vaisse, C., Halaas, J. L., Horvath, C. M., Darnell, J., Stoffel, M., and Friedman, J. M. (1996). Leptin activation of Stat3 in the hypothalamus of wild-type and ob/ob mice but not db/db mice. Nat. Genet. 14, 95–97. Cohen, P., Zhao, C., Cai, X., Montez, J. M., Rohani, S. C., Feinstein, P., Mombaerts, P., and Friedman, J. M. (2001). Selective deletion of leptin receptor in neurons leads to obesity. J. Clin. Invest. 108, 1113–1121. Kveiborg, M., Chiusaroli, R., Sims, N. A., Wu, M., Sabatakos, G., Horne, W. C., and Baron, R. (2002). The increased bone mass in deltaFosB transgenic mice is independent of circulating leptin levels. Endocrinology 143, 4304–4309. Takaya, K., Ogawa, Y., Isse, N., Okazaki, T., Satoh, N., Masuzaki, H., Mori, K., Tamura, N., Hosoda, K., and Nakao, K. (1996). Molecular cloning of rat leptin receptor isoform complementary DNAs– identification of a missense mutation in Zucker fatty (fa/fa) rats. Biochem. Biophys. Res. Commun. 225, 75–83. Mathey, J., Horcajada-Molteni, M. N., Chanteranne, B., Picherit, C., Puel, C., Lebecque, P., Cubizoles, C., Davicco, M. J., Coxam, V., and Barlet, J. P. (2002). Bone mass in obese diabetic Zucker rats: influence of treadmill running. Calcif. Tissue Int. 70, 305–311. Steppan, C. M., Crawford, D. T., Chidsey-Frink, K. L., Ke, H., and Swick, A. G. (2000). Leptin is a potent stimulator of bone growth in ob/ob mice. Regul. Pept. 92, 73–78. Nakajima, R., Inada, H., Koike, T., and Yamano, T. (2003). Effects of leptin to cultured growth plate chondrocytes. Horm. Res. 60, 91–98. Hamrick, M. W., Pennington, C., Newton, D., Xie, D., and Isales, C. (2004). Leptin deficiency produces contrasting phenotypes in bones of the limb and spine. Bone 34, 376–383. Elefteriou, F., Takeda, S., Ebihara, K., Magre, J., Patano, N., Kim, C. A., Ogawa, Y., Liu, X., Ware, S. M., Craigen, W. J., Robert, J. J., Vinson, C., Nakao, K., Capeau, J., and Karsenty, G. (2004). Serum leptin level is a regulator of bone mass. Proc. Natl Acad. Sci. USA 101, 3258–3263. Oh-I, S., Shimizu, H., Sato, T., Uehara, Y., Okada, S., and Mori, M. (2005). Molecular mechanisms associated with leptin

PAUL A. BALDOCK ET AL .

24.

25.

26.

27.

28.

29.

30.

31.

32.

33.

34.

35. 36.

37.

38.

39.

resistance: n-3 polyunsaturated fatty acids induce alterations in the tight junction of the brain. Cell Metab. 1, 331–341. Caro, J. F., Kolaczynski, J. W., Nyce, M. R., Ohannesian, J. P., Opentanova, I., Goldman, W. H., Lynn, R. B., Zhang, P. L., Sinha, M. K., and Considine, R. V. (1996). Decreased cerebrospinal-fluid/serum leptin ratio in obesity: a possible mechanism for leptin resistance. Lancet 348, 159–161. Munzberg, H., Flier, J. S., and Bjorbaek, C. (2004). Region-specific leptin resistance within the hypothalamus of diet-induced obese mice. Endocrinology 145, 4880–4889. Burguera, B., Hofbauer, L. C., Thomas, T., Gori, F., Evans, G. L., Khosla, S., Riggs, B. L., and Turner, R. T. (2001). Leptin reduces ovariectomy-induced bone loss in rats. Endocrinology 142, 3546–3553. Cornish, J., Callon, K. E., Bava, U., Lin, C., Naot, D., Hill, B. L., Grey, A. B., Broom, N., Myers, D. E., Nicholson, G. C., and Reid, I. R. (2002). Leptin directly regulates bone cell function in vitro and reduces bone fragility in vivo. J. Endocrin. 175, 405–415. Ozata, M., Ozdemir, I. C., and Licinio, J. (1999). Human leptin deficiency caused by a missense mutation: multiple endocrine defects, decreased sympathetic tone, and immune system dysfunction indicate new targets for leptin action, greater central than peripheral resistance to the effects of leptin, and spontaneous correction of leptin-mediated defects. J. Clin. Endocrinol. Metab. 84, 3686–3695. Farooqi, I. S., Jebb, S. A., Langmack, G., Lawrence, E., Cheetham, C. H., Prentice, A. M., Hughes, I. A., McCamish, M. A., and O’Rahilly, S. (1999). Effects of recombinant leptin therapy in a child with congenital leptin deficiency. New Engl. J. Med. 341, 879–884. Koh, J. M., Kim, D. J., Hong, J. S., Park, J. Y., Lee, K. U., Kim, S. Y., and Kim, G. S. (2002). Estrogen receptor alpha gene polymorphisms Pvu II and Xba I influence association between leptin receptor gene polymorphism (Gln223Arg) and bone mineral density in young men., Eur. J. Endocrinol. 147, 777–783. Quinton, N. D., Lee, A. J., Ross, R. J., Eastell, R., and Blakemore, A. I. (2001). A single nucleotide polymorphism (SNP) in the leptin receptor is associated with BMI, fat mass and leptin levels in postmenopausal Caucasian women. Hum. Genet. 108, 233–236. Miller, K. K., Grinspoon, S., Gleysteen, S., Grieco, K. A., Ciampa, J., Breu, J., Herzog, D. B., and Klibanski, A. (2004). Preservation of neuroendocrine control of reproductive function despite severe undernutrition. J. Clin. Endocrinol. Metab. 89, 4434–4438. Soyka, L. A., Grinspoon, S., Levitsky, L. L., Herzog, D. B., and Klibanski, A. (1999). The effects of anorexia nervosa on bone metabolism in female adolescents. J. Clin. Endocrinol. Metab. 84, 4489–4496. Welt, C. K., Chan, J. L., Bullen, J., Murphy, R., Smith, P., DePaoli, A. M., Karalis, A., and Mantzoros, C. S. (2004). Recombinant human leptin in women with hypothalamic amenorrhea. New Engl. J. Med. 351, 987–997. Reid, I. R. (2002). Relationships among body mass, its components, and bone. Bone 31, 547–555. Thomas, T., Burguera, B., Melton, L. J., Atkinson, E. J., O’Fallon, W. M., Riggs, B. L., and Khosla, S. (2001). Role of serum leptin, insulin, and estrogen levels as potential mediators of the relationship between fat mass and bone mineral density in men versus women. Bone 29, 114–120. Rauch, F., Blum, W. F., Klein, K., Allolio, B., and Schonau, E. (1998). Does leptin have an effect on bone in adult women? Calcif. Tissue Int. 63, 453–455. Matkovic, V., Ilich, J. Z., Skugor, M., Badenhop, N. E., Goel, P., Clairmont, A., Klisovic, D., Nahhas, R. W., and Landoll, J. D. (1997). Leptin is inversely related to age at menarche in human females. J. Clin. Endocrinol. Metab. 82, 3239–3245. Pasco, J. A., Henry, M. J., Kotowicz, M. A., Collier, G. R., Ball, M. J., Ugoni, A. M., and Nicholson, G. C. (2001). Serum leptin levels are associated with bone mass in nonobese women. J. Clin. Endocrinol. Metab. 86, 1884–1887.

Chapter 21 The Central Control of Bone Remodeling 40. Odabasi, E., Ozata, M., Turan, M., Bingol, N., Yonem, A., Cakir, B., Kutlu, M., and Ozdemir, I. C. (2000). Plasma leptin concentrations in postmenopausal women with osteoporosis. Eur. J. Endocrinol. 142, 170–173. 41. Sato, M., Takeda, N., Sarui, H., Takami, R., Takami, K., Hayashi, M., Sasaki, A., Kawachi, S., Yoshino, K., and Yasuda, K. (2001). Association between serum leptin concentrations and bone mineral density, and biochemical markers of bone turnover in adult men. J. Clin. Endocrinol. Metab. 86, 5273–5276. 42. Goulding, A., and Taylor, R. W. (1998). Plasma leptin values in relation to bone mass and density and to dynamic biochemical markers of bone resorption and formation in postmenopausal women. Calcif. Tissue Int. 63, 456–458. 43. Hoggard, N., Hunter, L., Duncan, J. S., Williams, L. M., Trayhurn, P., and Mercer, J. G. (1997). Leptin and leptin receptor mRNA and protein expression in the murine fetus and placenta. Proc. Natl Acad. Sci. USA 94, 11073–11078. 44. Thomas, T., Gori, F., Khosla, S., Jensen, M. D., Burguera, B., and Riggs, B. L. (1999). Leptin acts on human marrow stromal cells to enhance differentiation to osteoblasts and to inhibit differentiation to adipocytes. Endocrinology 140, 1360–1368. 45. Maor, G., Rochwerger, M., Segev, Y., and Phillip, M. (2002). Leptin acts as a growth factor on the chondrocytes of skeletal growth centers. J. Bone Miner. Res. 17, 1034–1043. 46. Enjuanes, A., Supervia, A., Nogues, X., and Diez-Perez, A. (2002). Leptin receptor (OB-R) gene expression in human primary osteoblasts: confirmation. J. Bone Miner. Res. 17, 1135. 47. Reseland, J. E., Syversen, U., Bakke, I., Qvigstad, G., Eide, L. G., Hjertner, O., Gordeladze, J. O., and Drevon, C. A. (2001). Leptin is expressed in and secreted from primary cultures of human osteoblasts and promotes bone mineralization. J. Bone Miner. Res. 16, 1426–1433. 48. Lee, Y. J., Park, J. H., Ju, S. K., You, K. H., Ko, J. S. and Kim, H. M. (2002). Leptin receptor isoform expression in rat osteoblasts and their functional analysis. FEBS Lett. 528, 43–47. 49. Morroni, M., De Matteis, R., Palumbo, C., Ferretti, M., Villa, I., Rubinacci, A., Cinti, S., and Marotti, G. (2004). In vivo leptin expression in cartilage and bone cells of growing rats and adult humans. J. Anat. 205, 291–296. 50. Reseland, J. E., Syversen, U., Bakke, I., Qvigstad, G., Eide, L. G., Hjertner, O., Gordeladze, J. O., and Drevon, C. A. (2002). Leptin is expressed in and secreted from primary cultures of human osteoblasts and promotes bone mineralization. J. Bone Miner. Res. 16, 1426–1433. 51. Gordeladze, J. O., Drevon, C. A., Syversen, U., and Reseland, J. E. (2002). Leptin stimulates human osteoblastic cell proliferation, de novo collagen synthesis, and mineralization: Impact on differentiation markers, apoptosis, and osteoclastic signaling. J. Cell Biochem. 85, 825–836. 52. Holloway, W. R., Collier, F. M., Aitken, C. J., Myers, D. E., Hodge, J. M., Malakellis, M., Gough, T. J., Collier, G. R., and Nicholson, G. C. (2002). Leptin inhibits osteoclast generation. J. Bone Miner. Res. 17, 200–209. 53. Satoh, N., Ogawa, Y., Katsuura, G., Numata, Y., Tsuji, T., Hayase, M., Ebihara, K., Masuzaki, H., Hosoda, K., Yoshimasa, Y., and Nakao, K. (1999). Sympathetic activation of leptin via the ventromedial hypothalamus: leptin-induced increase in catecholamine secretion. Diabetes 48, 1787–1793. 54. Minkowitz, B., Boskey, A. L., Lane, J. M., Pearlman, H. S., and Vigorita, V. J. (1991). Effects of propranolol on bone metabolism in the rat. J. Orthop. Res. 9, 869–875. 55. Zeman, R. J., Hirschman, A., Hirschman, M. L., Guo, G., and Etlinger, J. D. (1991). Clenbuterol, a beta 2-receptor agonist, reduces net bone loss in denervated hindlimbs. Am. J. Physiol. 261, E285–289. 56. Kellenberger, S., Muller, K., Richener, H., and Bilbe, G. (1998). Formoterol and isoproterenol induce c-fos gene expression in osteoblastlike cells by activating beta2-adrenergic receptors. Bone 22, 471–478.

375 57. Moore, R. E., Smith, C. K., Bailey, C. S., Voelkel, E. F., and Tashjian, A. H. (1993). Characterization of beta-adrenergic receptors on rat and human osteoblast-like cells and demonstration that betareceptor agonists can stimulate bone resorption in organ culture. Bone Miner. 23, 301–315. 58. Takeuchi, T., Tsuboi, T., Arai, M., and Togari, A. (2000). Adrenergic stimulation of osteoclastogenesis mediated by expression of osteoclast differentiation factor in MC3T3-E1 osteoblast-like cells. Biochem. Pharmacol. 61, 5. 59. Pasco, J. A., Henry, M. J., Sanders, K. M., Kotowicz, M. A., Seeman, E., and Nicholson, G. C. (2004). Beta-adrenergic blockers reduce the risk of fracture partly by increasing bone mineral density: Geelong Osteoporosis Study. J. Bone Miner. Res. 19, 19–24. 60. Schlienger, R. G., Kraenzlin, M. E., Jick, S. S., and Meier, C. R. (2004). Use of beta-blockers and risk of fractures. JAMA 292, 1326–1332. 61. Rejnmark, L., Vestergaard, P., Kassem, M., Christoffersen, B. R., Kolthoff, N., Brixen, K., and Mosekilde, L. (2004). Fracture risk in perimenopausal women treated with beta-blockers. Calcif. Tissue Int. 75, 365–372. 62. Bertagna, X. (1994). Proopiomelanocortin-derived peptides. Endocrinol. Metab. Clin. North Am. 23, 467–485. 63. Solomon, S. (1999). POMC-derived peptides and their biological action. Ann. N Y Acad. Sci. 885, 22–40. 64. Seeley, R. J., Drazen, D. L., and Clegg, D. J. (2004). The critical role of the melanocortin system in the control of energy balance. Annu. Rev. Nutr. 24, 133–149. 65. Mountjoy, K. G., Wu, C. S., Cornish, J., and Callon, K. E. (2003). alpha-MSH and desacetyl-alpha-MSH signaling through melanocortin receptors. Ann. N Y Acad. Sci. 994, 58–64. 66. Shimizu, H., Shargill, N. S., Bray, G. A., Yen, T. T., and Gesellchen, P. D. (1989). Effects of MSH on food intake, body weight and coat color of the yellow obese mouse. Life Sci. 45, 543–552. 67. Farooqi, I. S., Yeo, G. S., Keogh, J. M., Aminian, S., Jebb, S. A., Butler, G., Cheetham, T., and O’Rahilly, S. (2000). Dominant and recessive inheritance of morbid obesity associated with melanocortin 4 receptor deficiency. J. Clin. Invest. 106, 271–279. 68. Pernow, J., Ohlen, A., Hokfelt, T., Nilsson, O., and Lundberg, J. M. (1987). Neuropeptide Y: presence in perivascular noradrenergic neurons and vasoconstrictor effects on skeletal muscle blood vessels in experimental animals and man. Regul. Pept. 19, 313–324. 69. Parker, R. M., and Herzog, H. (1999). Regional distribution of Y-receptor subtype mRNAs in rat brain. Eur. J. Neurosci. 11, 1431–1448. 70. Morris, J. L. (1994). Selective constriction of small cutaneous arteries by NPY matches distribution of NPY in sympathetic axons. Regul. Pept. 49, 225–236. 71. Hokfelt, T., Broberger, C., Zhang, X., Diez, M., Kopp, J., Xu, Z., Landry, M., Bao, L., Schalling, M., Koistinaho, J., DeArmond, S. J., Prusiner, S., Gong, J., and Walsh, J. H. (1998). Neuropeptide Y: some viewpoints on a multifaceted peptide in the normal and diseased nervous system. Brain Res. Brain Res. Rev. 26, 154–166. 72. Wettstein, J. G., Earley, B., and Junien, J. L. (1995). Central nervous system pharmacology of neuropeptide Y. Pharmacol. Ther. 65, 397–414. 73. Lundberg, J. M., Terenius, L., Hokfelt, T., and Tatemoto, K. (1984). Comparative immunohistochemical and biochemical analysis of pancreatic polypeptide-like peptides with special reference to presence of neuropeptide Y in central and peripheral neurons. J. Neurosci. 4, 2376–2386. 74. Ekblad, E., and Sundler, F. (2002). Distribution of pancreatic polypeptide and peptide YY. Peptides 23, 251–261. 75. Ueno, N., Inui, A., Iwamoto, M., Kaga, T., Asakawa, A., Okita, M., Fujimiya, M., Nakajima, Y., Ohmoto, Y., Ohnaka, M., Nakaya, Y., Miyazaki, J. I., and Kasuga, M. (1999). Decreased food intake and body weight in pancreatic polypeptide-overexpressing mice. Gastroenterology 117, 1427–1432.

376 76. Starback, P., Wraith, A., Eriksson, H., and Larhammar, D. (2000). Neuropeptide Y receptor gene y6: multiple deaths or resurrections? Biochem. Biophys. Res. Commun. 277, 264–269. 77. Lin, S., Boey, D., Couzens, M., Lee, N., Sainsbury, A., and Herzog, H. (2005). Compensatory changes in [125I]-PYY binding in Y receptor knockout mice suggest the potential existence of further Y receptor(s). Neuropeptides 39, 21–28. 78. Bard, J. A., Walker, M. W., Branchek, T. A., and Weinshank, R. L. (1995). Cloning and functional expression of a human Y4 subtype receptor for pancreatic polypeptide, neuropeptide Y, and peptide YY. J. Biol. Chem. 270, 26762–26765. 79. Selbie, L. A., Darby, K., Schmitz-Peiffer, C., Browne, C. L., Herzog, H., Shine, J., and Biden, T. J. (1995). Synergistic interaction of Y1-neuropeptide Y and alpha 1b-adrenergic receptors in the regulation of phospholipase C, protein kinase C, and arachidonic acid production. J. Biol. Chem. 270, 11789–11796. 80. Herzog, H., Hort, Y. J., Ball, H. J., Hayes, G., Shine, J., and Selbie, L. A. (1992). Cloned human neuropeptide Y receptor couples to two different second messenger systems. Proc. Natl Acad. Sci. USA 89, 5794–5798. 81. Balasubramaniam, A. (2003). Neuropeptide Y (NPY) family of hormones: progress in the development of receptor selective agonists and antagonists. Curr. Pharm. Des. 9, 1165–1175. 82. King, P. J., Williams, G., Doods, H., and Widdowson, P. S. (2000). Effect of a selective neuropeptide Y Y(2) receptor antagonist, BIIE0246 on neuropeptide Y release. Eur. J. Pharmacol. 396, R1–R3. 83. Bjurholm, A., Kreicbergs, A., Terenius, L., Goldstein, M., and Schultzberg, M. (1988). Neuropeptide Y-, tyrosine hydroxylase- and vasoactive intestinal polypeptide-immunoreactive nerves in bone and surrounding tissues. J. Auton. Nerv Syst. 25, 119–125. 84. Ahmed, M., Bjurholm, A., Kreicbergs, A., and Schultzberg, M. (1993). Neuropeptide Y, tyrosine hydroxylase and vasoactive intestinal polypeptide-immunoreactive nerve fibers in the vertebral bodies, discs, dura mater, and spinal ligaments of the rat lumbar spine. Spine 18, 268–273. 85. Lindblad, B. E., Nielsen, L. B., Jespersen, S. M., Bjurholm, A., Bunger, C., and Hansen, E. S. (1994). Vasoconstrictive action of neuropeptide Y in bone. The porcine tibia perfused in vivo. Acta. Orthop. Scand. 65, 629–634. 86. Sisask, G., Bjurholm, A., Ahmed, M., and Kreicbergs, A. (1996). The development of autonomic innervation in bone and joints of the rat. J. Auton. Nerv. Syst. 59, 27–33. 87. Ahmed, M., Bjurholm, A., Theodorsson, E., Schultzberg, M., and Kreicbergs, A. (1995). Neuropeptide Y- and vasoactive intestinal polypeptide-like immunoreactivity in adjuvant arthritis: effects of capsaicin treatment. Neuropeptides 29, 33–43. 88. Bjurholm, A., Kreicbergs, A., Schultzberg, M., and Lerner, U. H. (1992). Neuroendocrine regulation of cyclic AMP formation in osteoblastic cell lines (UMR-106–01, ROS 17/2.8, MC3T3-E1, and Saos-2) and primary bone cells. J. Bone Miner. Res. 7, 1011–1019.

PAUL A. BALDOCK ET AL . 89. Bjurholm, A. (1991). Neuroendocrine peptides in bone. Int. Orthop. 15, 325–329. 90. Togari, A., Arai, M., Mizutani, S., Mizutani, S., Koshihara, Y., and Nagatsu, T. (1997). Expression of mRNAs for neuropeptide receptors and beta-adrenergic receptors in human osteoblasts and human osteogenic sarcoma cells. Neurosci. Lett. 233, 125–128. 91. Nakamura, M., Sakanaka, C., Aoki, Y., Ogasawara, H., Tsuji, T., Kodama, H., Matsumoto, T., Shimizu, T., and Noma, M. (1995). Identification of two isoforms of mouse neuropeptide Y-Y1 receptor generated by alternative splicing. Isolation, genomic structure, and functional expression of the receptors. J. Biol. Chem. 270, 30102–30110. 92. Baskin, D. G., Breininger, J. F., and Schwartz, M. W. (1999). Leptin receptor mRNA identifies a subpopulation of neuropeptide Y neurons activated by fasting in rat hypothalamus. Diabetes 48, 828–833. 93. Broberger, C., Landry, M., Wong, H., Walsh, J. N., and Hokfelt, T. (1997). Subtypes Y1 and Y2 of the neuropeptide Y receptor are respectively expressed in pro-opiomelanocortin- and neuropeptide-Ycontaining neurons of the rat hypothalamic arcuate nucleus. Neuroendocrinology 66, 393–408. 94. Sainsbury, A., Schwarzer, C., Couzens, M., Fetissov, S., Furtinger, S., Jenkins, A., Cox, H. M., Sperk, G., Hokfelt, T., and Herzog, H. (2002). Important role of hypothalamic Y2 receptors in body weight regulation revealed in conditional knockout mice. Proc. Natl Acad. Sci. USA 99, 8938–8943. 95. Sainsbury, A., Baldock, P. A., Schwarzer, C., Ueno, N., Enriquez, R. F., Couzens, M., Inui, A., Herzog, H., and Gardiner, E. M. (2003). Synergistic effects of Y2 and Y4 receptors on adiposity and bone mass revealed in double knockout mice. Mol. Cell Biol. 23, 5225–5233. 96. Sainsbury, A., Schwarzer, C., Couzens, M., Jenkins, A., Oakes, S. R., Ormandy, C. J., and Herzog, H. (2002). Y4 receptor knockout rescues fertility in ob/ob mice. Genes Dev. 16, 1077–1088. 97. Stephens, T. W., Basinski, M., Bristow, P. K., Bue-Valleskey, J. M., Burgett, S. G., Craft, L., Hale, J., Hoffmann, J., Hsiung, H. M., Kriauciunas, A., MacKellar, W., Rosteck, P. R. Jr., Schoner, B., Smith, D., Tinsley, F. C., Zhang, X., and Heiman, M. (1995). The role of neuropeptide Y in the antiobesity action of the obese gene product. Nature 377, 530–532. 98. Erickson, J. C., Hollopeter, G., and Palmiter, R. D. (1996). Attenuation of the obesity syndrome of ob/ob mice by the loss of neuropeptide Y. Science 274, 1704–1707. 99. Baldock, P. A., Sainsbury, A., Allison, S., Lin, D., Couzens, M., Boey, D., Enriquez, R. F., During, M., Herzog, H., and Gardiner, E. M. (2005). Hypothalamic control of bone formation: Distinct actions of leptin and Y2 receptor pathways. J. Bone Miner. Res. 20, 1851–1857. 100. Konttinen, Y., Imai, S., and Suda, A. (1996). Neuropeptides and the puzzle of bone remodeling. State of the art. Acta Orthop. Scand. 67, 632–639. 101. Lerner, U.H., and Lundberg, P. (2002). Kinins and Neuro-osteogenic factors. In Principles of Bone Biology, G. A. Rodan (ed.). Academic Press, San Diego, 1, 773–799.

Chapter 22

New Concepts in Bone Remodeling David W. Dempster and Hua Zhou

I. II. III. IV.

V. Possible Mechanisms whereby a Reduction in Activation Frequency may Protect against Fracture References

Introduction An Overview of the Remodeling Cycle Functions of Bone Remodeling The Role of Apoptosis in Regulating Bone Balance

II. AN OVERVIEW OF THE REMODELING CYCLE

I. INTRODUCTION One of the many remarkable features of the mammalian skeleton is its ability to constantly renew itself throughout life. This is achieved by means of the concerted efforts of a diverse group of cells, collectively referred to as the bone remodeling unit (BRU), which includes osteoclasts, osteoblasts, osteocytes, and other accessory cells whose identity and functions are still obscure. Our current knowledge of the mechanics of bone remodeling has been reviewed in detail elsewhere [1–5] and will be considered only briefly here. The purpose of this chapter is to review some new concepts pertaining to the activation of BRUs, intercellular communication within the BRU, and the role of apoptosis in regulating the bone balance within each BRU and, ultimately, bone mass. We shall also consider why a reduction in remodeling activation frequency in osteoporotic patients may reduce the risk of fracture without substantial increments in bone mass. Dynamics of Bone and Cartilage Metabolism

Regional Bone Center, Helen Hayes Hospital, West Haverstraw, New York, USA

There are four distinct phases in the remodeling cycle: activation, resorption, reversal, and formation (Figs 1 and 2). Activation refers to the initiating event that converts a previously quiescent bone surface into a remodeling one. The main processes involved are recruitment of mononucleated osteoclast precursors from cells in the monocyte– macrophage lineage in the circulation [6], penetration of the bone lining cell layer, and fusion of the mononuclear cells to form multinucleated preosteoclasts. The extent to which the activation process is deterministic (purposeful) and the extent to which it is stochastic (random) is unclear. However, as will be discussed below, there is evidence that at least some remodeling is targeted towards the repair of fatigue damage, but most remodeling is probably stochastic and regulated by hormones [7, 8]. The formation, activation, and activity of osteoclasts are all regulated by local 377

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378

Figure 1

Cross-sectional diagrams of evolving BRU in cancellous bone (upper) and cortical bone (lower). The arrow indicates the direction of movement through space. Note that the cancellous BRU is essentially one half of the cortical BRU.

Figure 2

Scanning electron micrograph of a BRU in human cancellous bone. Note the eroded surface (ES) underlying an area of new bone formation (BF), and the forming osteocyte lacunae (arrowheads) in the area of new bone formation.

DAVID W. DEMPSTER AND HUA ZHOU

cytokines such as receptor activator of nuclear factor-κB ligand (RANKL), interleukins-1 and -6 (IL-1 and IL-6), colony-stimulating factors (CSFs), and systemic hormones such as parathyroid hormone, 1,25-dihydroxyvitamin D3 and calcitonin [6, 9–12]. Osteoclasts are ultimately activated by membrane signaling through RANK, the cognate receptor for RANKL, a member of the family of tumor necrosis factors expressed on marrow stromal cells and osteoblasts. The RANKL/RANK interaction promotes the differentiation of osteoclast precursors and results in the activation and prolonged survival of mature osteoclasts. Osteoprotegerin (OPG), a member of the superfamily of tumor necrosis factor receptors, is a decoy receptor for RANKL. The binding of RANKL to OPG results in inhibition of the differentiation and activity of osteoclasts. The preosteoclasts bind tightly to the bone matrix via interaction between integrin receptors in their membranes and RGD-containing peptides in the organic matrix, creating an annular sealing zone. The resorption phase of the cycle begins when fully differentiated osteoclasts secrete protons and proteolytic enzymes into the hemivacuole created by the sealing zone (Fig. 3). The proteolytic enzymes are primarily cathepsins, the most important one being cathepsin K [13]. Initially, resorption is accomplished by multinucleated osteoclasts, but there is some evidence to suggest that the job is completed by mononucleated cells [14]. Recent in vitro studies have confirmed that mononuclear cells derived from human peripheral monocytes are capable of extensive resorption [15]. The ending of the resorption phase, signaled by apoptosis of osteoclasts [16], is followed by reversal. The mechanism of osteoclast apoptosis remains unclear. Studies indicate that TGF-β, sex steroids such as estradiol and testosterone promote osteoclast apoptosis [6], and Fas/Fas ligandmediated osteoclast apoptosis [17]. During reversal, resorption lacunae are occupied by mononuclear cells, including monocytes, osteocytes that have been released from bone, and osteoblasts that are moving in to begin the formation phase of the cycle [18]. It is during the reversal phase that coupling signals, derived both from the resorbed organic matrix and osteoblast lineage cells, summon osteoblast progenitors into the resorption site and promote the proliferation and differentiation of osteoblast progenitors. One attractive feature of the hypothesis that bone matrix serves as a source of osteoblast growth factors is that it provides a mechanism whereby the number of osteoblast precursors recruited could be quantitatively related to the amount of bone removed and, therefore, the amount that has to be replaced. The nature of the signals is not completely known, but osteoblast growth factors, such as transforming growth factor (TGF)-β, insulin-like growth factors I and II (IGF-I and -II), bone morphogenetic proteins (BMPs),

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Chapter 22 New Concepts in Bone Remodeling

Figure 3

Human osteoclast (OC) fixed in the process of excavating a resorption lacuna (RL) in vitro. Note the sharpness of the edge (arrowheads) between unresorbed and resorbed areas indicating how tightly fitting the sealing zone must be. Reproduced with permission from Reference 107.

platelet-derived growth factor (PDGF) and fibroblast growth factor (FGF), are plausible candidates [19–24]. A crucial transcription factor for osteoblast development, the core binding factor 1 (Cbfa1 or Runx2), also plays an important role in the recruitment of osteoblasts to the remodeling site by directing the commitment of multipotential mesenchymal stem cells to osteoprogenitor cells and promoting the proliferation and differentiation of osteoprogenitors to mature osteoblasts [25]. TGF-β has also recently been shown to prolong osteoblast life-span in vitro by inhibiting apoptosis (see below). That TGF-β plays an important role in bone remodeling is supported by the demonstration that the concentration of TGF-β is positively correlated with histomorphometric indices of bone resorption and bone formation and with serum levels of osteocalcin and bone-specific alkaline phosphatase [26]. Formation is a two-step process in which the osteoblasts initially synthesize the organic matrix and then preside over its mineralization. As bone formation ensues, osteoblasts become incarcerated in the matrix as osteocytes. These cells, however, are not in “solitary confinement”. They maintain close contact with each other via gap junctions between cytoplasmic processes that course through the canaliculae. They also make contact with lining cells on the inactive bone surface and with osteoblasts and osteoclasts on the remodeling bone surface via these processes.

These cell–cell connections form a complex, threedimensional web (Fig. 4), which puts the osteocytes in an ideal position to “sense” a change in the mechanical properties of the surrounding bone and to communicate this to the cells on the surface to initiate or regulate bone remodeling if it is necessary [27–29]. This ultimately results in bone structure adapting to changes in mechanical usage. The intercellular link could occur by wiring transmission (WT) and/or volume transmission (VT), these being two mechanisms of communication that have been defined in the central nervous system [30]. WT refers to signaling through synapses, gap junctions, and cell–cell adhesion molecules, such as connexin43 [31]. VT occurs by diffusion of soluble factors (hormones, cytokines, and growth factors) through the canalicular network. When formation is completed, the inactive osteoblasts become flattened and elongated and are referred to as lining cells. These cells have long been thought to regulate the flow of ions in and out of the bone extracellular fluid. Furthermore, it has recently been suggested that under certain conditions, for example stimulation by PTH or mechanical force, bone-lining cells can revert to boneforming osteoblasts [32, 33]. The newly formed piece of bone is referred to as a “basic structural unit” (BSU) of bone (Fig. 5). In cancellous bone, it is generally called a “packet” or a “trabecular osteon”; in cortical bone it is an “Haversian system” or a “cortical osteon”. Note that

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Figure 5

Basic structural units in cancellous bone (A) and cortical bone (B). The arrowheads delineate reversal lines. Reproduced with permission from Reference 4.

Figure 4 Transmission (A) and scanning (B) electron micrographs showing osteocyte processes communicating with cells on the bone surface. Reproduced with permission from Reference 30.

III. FUNCTIONS OF BONE REMODELING

the process of bone remodeling is essentially identical in cancellous and cortical bone. The BRU in cortical bone is equivalent to two apposed cancellous BRUs (Fig. 1) [34]. The difference between the volume of bone removed by the osteoclasts and that replaced by the osteoblasts is referred to as the “bone balance”. BRUs on the periosteal surface of cortical bone produce a slightly positive bone balance so that, with aging, the periosteal circumference increases as the effect of the small positive balance in each BRU accumulates. Remodeling units on the endosteal surface of cortical bone are in negative balance so that the marrow cavity enlarges with age. Furthermore, the balance is more negative on the endosteal surface than it is on the periosteal surface and, as a result, cortical thickness decreases with age. The bone balance on cancellous surfaces is also negative, resulting in a gradual thinning of the trabecular plates with the passage of time [1].

Two principal functions of bone remodeling are known, although some have argued convincingly that there must be other functions, of which we are as yet unaware, or other reasons why the human skeleton undergoes such extensive remodeling [35]. The main functions of bone remodeling are presumed to be the preventive maintenance of mechanical strength by continuously replacing fatigued bone by new, mechanically sound bone, and, secondly, an important role in mineral homeostasis by providing access to the skeletal stores of calcium and phosphorus. While the turnover rate of 2–3% per year in cortical bone is consistent with maintenance of mechanical properties, the turnover rate in cancellous bone is much higher than would be required for this purpose, suggesting that this is driven more by its role in mineral metabolism [5]. In extreme situations, however, cortical bone can participate in mineral homeostasis. A striking example of this is the increase in cortical remodeling that occurs in Rocky Mountain deer during the antler-forming

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season [36]. Cortical porosity increases dramatically, but temporarily, during this time to provide the mineral necessary for antler growth. In less extreme situations, it has been proposed [5] that activation of new remodeling units is not required for short-term mineral homeostasis, but that this may only require the presence of a minimum number of osteoclasts whose activity can be regulated. Continual remodeling activity therefore ensures the presence of this contingent of osteoclasts. Remodeling also ensures the continuous supply of young bone of low mineral density, which is necessary for ionic exchange at quiescent surfaces [5]. There are also a number of other postulated functions related to the skeleton’s role as a depository. One of these is the skeleton’s role in acid–base balance with bone serving as a source of buffer in the form of bicarbonate [37]. The skeleton is also a depot for growth factors and cytokines, some of which, for example transforming growth factor β, are activated by the acidic microenvironment created by the osteoclast. Remodeling may therefore function to supply hematopoietic marrow with these factors. This may account for some of the excess remodeling that occurs at cancellous sites where marrow and bone co-exist in a symbiotic manner [5]. The best experimental evidence for a role of bone remodeling in the maintenance of the mechanical integrity of the skeleton has come from studies in dogs [38, 39]. When long bones were loaded to induce fatigue damage and then studied histologically, a statistical association between microcracks and resorption spaces was demonstrated, with as much as six times as many cracks associated with resorption spaces as predicted by chance alone (Fig. 6) [40]. This could be explained by a propensity of microcracks to form close to resorption spaces. However, further experiments showed that the activation of remodeling was in response to the appearance of microcracks [39]. This was demonstrated by loading the left forelimbs of dogs 8 days prior to sacrifice and loading the right forelimbs immediately prior to sacrifice. If cracks simply localize at sites of resorption, then the number of cracks associated with resorption spaces should have been the same in each limb. The data showed that there was the same number of microcracks in each limb, but that the limb that was loaded first contained more resorption spaces and a greater association between microcracks and resorption than the limb that was loaded immediately prior to sacrifice. The conclusion from this observation is that microcracks cause the appearance of resorption spaces. One proposed mechanism for this deterministic remodeling is that microcracks may result in the debonding of a cortical osteon (Fig. 7) [41]. Consequently, there is a reduction in stress and strain in that aspect of the osteon, which could be “sensed” by the osteocytes and communicated to the

Figure 6 Images of a resorption space (Rc) associated with a microcrack (arrows) and another resorption space (R) that has no associated microcrack. A is a secondary electron image and B is backscattered electron image of the same sample. Reproduced with permission from Reference 38.

surface-lining cells by means of the lacuno-canalicular network. The lining cells could then initiate activation of a bone remodeling unit from the Haversian canal and this would move in the direction of the damage. Once the crack was reached the bone remodeling unit would turn to move in a longitudinal direction, thus repairing the crack.

IV. THE ROLE OF APOPTOSIS IN REGULATING BONE BALANCE One of the most interesting concepts to be applied to the study of bone biology in recent years is that of apoptosis or programmed cell death. The biological significance of programmed cell death was first recognized by Kerr, Wyllie, and Currie, who also coined the term apoptosis to describe it [42, 43]. In contrast to necrosis, cells dying by apoptosis do so in a purposeful and orderly manner;

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Figure 7 Possible mechanism whereby microcracks may initiate their own repair. Reproduced with permission from Reference 41.

they shrink rather than swell, and do not spill their cytoplasmic contents, thus avoiding a local inflammatory response. Apoptosis plays a crucial role in the size and shape of organs and limbs during embryonic development and a failure of apoptosis is responsible for the unbridled growth of solid tumors. Similarly, apoptosis of bone cells is integral to skeletal development and physiological bone turnover [44–47]. One of the earliest descriptions of apoptosis in bone was in osteogenic cells involved in the formation of calvarial sutures [48]. The leading osteoblasts appeared

to die by apoptosis in regions where there was a failure of the normal pattern of overlap between two approaching segments of bone. In the postnatal and adult skeletons, apoptosis ultimately regulates bone volume and architecture by controlling the balance among proliferation, differentiation, and death of osteoblasts and osteoclasts [49, 50]. After the termination of bone formation, a proportion of osteoblasts undergo apoptosis and the remainder differentiate into either osteocytes or bone-lining cells. Apoptosis has also been observed in osteoclasts (Fig. 8) and osteocytes.

Figure 8 A series of transmission electron micrographs illustrating different stages of apoptosis in cultured neonatal rat osteoclasts. (A) Normal osteoclast. (B) Two osteoclasts in an early stage of apoptosis. Note cell shrinkage and compaction of nuclear chromatin against the nuclear membrane. (C) A later stage; the chromatin is condensed and fragmented and there is blebbing of the plasma membrane. These blebs contain recognizable cytoplasmic structures and are termed “apoptotic bodies”. (D) Final stage; the cell has assumed a small spherical shape and chromatin is highly condensed. Original magnification: A = ×2925; B = ×5010; C = ×5000; D = ×11 000. Reproduced with permission from Reference 46.

Chapter 22 New Concepts in Bone Remodeling

Because of its low frequency and transient nature, apoptosis is rarely observed in bone under normal conditions [44]. However, it is readily observed in the presence of agents that perturb the remodeling cycle. For example, osteoclast apoptosis is enhanced in the presence of estrogen [51, 52], selective estrogen receptor modulators (SERMs) [51, 53–55], bisphosphonates [56], and glucocorticoids [46], and is suppressed in the presence of calcitonin [57], and macrophage colony-stimulating factor [58]. Many factors regulate apoptosis in osteoblasts, including FGF, FGF2, IGF-I, IGF-II, and PDGF. Apoptosis of osteoblasts is induced by glucocorticoids [59, 60–62], human T-cell leukemia virus type I tax protein [63] and tumor necrosis factor [59, 63, 64], and is inhibited by various factors, such as transforming growth factor-β and interleukin-6-type cytokines, that are present in the bone/bone marrow environment [59]. Enhanced apoptosis of osteoclasts in response to estrogen, SERMs, and bisphosphonates may account for part of the therapeutic efficacy of these agents in the prevention and treatment of osteoporosis. With regard to estrogen, it has been suggested that a reduction in osteoclast apoptosis after menopause may contribute to the enhanced resorption and increased erosion depth that occurs at this time [44, 51]. Reducing apoptosis would extend the lifespan of individual osteoclasts, perhaps allowing them to penetrate deeper into the trabecular plates and thus shifting bone balance in a negative direction. A negative shift in bone balance would also occur in situations in which apoptosis is enhanced in osteoblasts, e.g. in the presence of glucocorticoid excess [65]. This could be an important component of the mechanism underlying the reduction in the wall thickness of trabecular packets that is observed in glucocorticoid-induced osteoporosis [66]. Osteocyte apoptosis is also enhanced by glucocorticoids [65], which may be a contributing factor to the aseptic or avascular osteonecrosis that is sometimes seen following high-dose glucocorticoid therapy. The finding that glucocorticoids enhanced apoptosis in rat osteoclasts [46] initially appeared to be at odds with their well-known action to enhance bone loss. However, several studies have revealed that, in the rat, glucocorticoid excess actually inhibits bone resorption, increases bone mass, and protects against bone loss induced by ovariectomy, dietary calcium deficiency, and immobilization [67–69]. It has been shown that apoptosis in osteocytes appears to be influenced by microdamage induced by mechanical loading [70–72]. Noble et al. and Verborgt et al. found that large-scale osteocyte apoptosis was associated with cortical remodeling induced by mechanical loading in a rat model of fatigue fracture [70–74]. Furthermore, Bronckers et al. [75] observed enhanced osteocyte apoptosis in deeper

383 layers of bone, i.e. in older cells. This was mainly seen in locations where intense bone resorption was occurring. However, mechanical loading to produce strains within the physiological range appeared to reduce osteocyte apoptosis [70]. Consequently, it has been proposed that, while undergoing apoptosis, osteocytes may transmit signals that are conveyed through the lacuno-canalicular system to enhance osteoclast recruitment or activation. Whatever the exact mechanism, it seems clear that osteocyte apoptosis plays an important role in the targeting of remodeling to replace microdamaged bone tissue with new, healthy bone.

V. POSSIBLE MECHANISMS WHEREBY A REDUCTION IN ACTIVATION FREQUENCY MAY PROTECT AGAINST FRACTURE At first glance, the postulate that a reduction in remodeling rate may reduce fracture risk seems counterintuitive if, as discussed above, we believe that an important function of remodeling is to maintain bone strength. In fact, there are relatively few data showing that reducing bone turnover results in an increase in fractures. The work that is most often cited is Flora et al.’s study [76] of the effects of etidronate in dogs, which showed an increase in spontaneous fractures at doses which severely depressed bone turnover. It should be noted, however, that this was a very dramatic inhibition of turnover, and, secondly, at these high doses, etidronate may have adversely affected mineralization, which could also increase skeletal fragility. More recently, treatment of dogs with alendronate or risedronate was shown to increase the density and length of microcracks in bone and this was associated with a decrease in the intrinsic toughness of the bone tissue [77]. However, again the doses of bisphosphonates were five times those used clinically and, despite the increase in microdamage, whole bone strength was not affected, presumably because changes in the mechanical properties of the tissue itself were compensated for by an increase in bone mass [77]. In contrast to this finding there is a growing body of data to support the view that high turnover rates are associated with increased fracture risk. Some of these data have come from two large prospective studies. The EPIDOS study [78] followed 7500 women for 22 months, and the Rotterdam study [79] followed 8000 men and women for almost 30 months. The EPIDOS study found an odds ratio (or risk of fracture) of 2.7 for patients with low bone mass and an odds ratio (2.2) that was almost as high for patients with normal bone mass who had high levels of urinary c-terminal telopeptide (CTx), a bone resorption marker.

384 Patients who had both low bone mass and high urinary CTx were at the highest risk of fracture (odds ratio of 4.8). Similar results were obtained in the Rotterdam study with a significant association between a resorption marker level and fractures in women, even when adjusted for bone mineral density at the femur [79]. It has been known for many years that the degree of reduction in bone mass is a good predictor of fracture risk but it is also true that the magnitude of the increase in bone mass upon treatment is not always a good predictor of the degree of fracture protection. One obvious example is sodium fluoride, which has been shown to produce significant increases in bone mass without affording fracture protection [80, 81]. This is due to the fact that the matrix formed in the presence of sodium fluoride is mechanically inferior [82]. From an analysis of the data from the Fracture Intervention Trial, Cummings et al. [83] found that the observed reduction in fracture incidence following treatment with alendronate was much greater than that predicted from the change in bone density alone. This was also the case for a number of other antiresorptive treatments in which the proportion of fracture protection that could be explained by the observed increase in bone mass ranged from about 70% in the case of alendronate to as low as 5% in the case of calcitonin (Fig. 9). Thus, there are now compelling data to support the hypothesis that a reduction in remodeling activation frequency reduces fracture risk by mechanisms other than those associated with an increase in bone mass. What are some of the possible mechanisms for this phenomenon? First, reducing activation frequency has the effect of reducing the size of the remodeling space, which results in only a modest increase in bone mass [2]. Perhaps more important, however, it also reduces the number of resorptive sites that are active at any one time. In a trabecular network

Figure 9

The increments in bone mass achieved by antiresorptive, or, more correctly, antiremodeling, agents account for only a part, and in some cases a small part, of the degree of fracture protection observed. Drawn from data presented in Reference 83.

DAVID W. DEMPSTER AND HUA ZHOU

that is already compromised by bone loss, the resorptive phase of the remodeling cycle may weaken crucial, but unsupported, surviving trabeculae to the point of failure without causing a significant decrease in bone mass [84]. Furthermore, high turnover rates increase the probability that resorption will occur simultaneously at more than one point along the length of such trabeculae, rendering them vulnerable to buckling and failure (Figs. 10 and 11). By analogy, a v-cut in the shaft of a thin walking stick has little effect on its mass but greatly reduces its ability to support a load, and several v-cuts have an even greater deleterious effect. In high turnover situations, the probability of trabecular plates being perforated is also increased.

Figure 10 Scanning electron micrographs of human cancellous bone. In (A) the entire circumference of a thinned trabecular rod in an osteoporotic patient rod is covered in resorption bays. Note that the rod has cracked at three points along its length. While this most likely occurred ex vivo it nevertheless indicates how fragile the rod was. In (B) a trabecular plate in a postmenopausal woman has been subjected to extensive osteoclastic resorption and has been perforated in the area between the arrowheads. A few millimeters to the left the plate has been breached by osteoclasts working from the other side (single arrowhead). Reproduced with permission from References 108 and 4, respectively.

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Figure 11 Theoretical explanation for the increase in bone fragility associated with enhanced remodeling and why treatment with an agent that reduces bone turnover may be beneficial without changing bone mass significantly. In a normal subject (N) the two resorption cavities do not represent a threat to the stability of the trabecula because it is buttressed by the two horizontal trabeculae. In an osteoporotic individual (OP) the same two cavities significantly increase the chance of failure because the horizontal trabeculae have disappeared and the trabecula is thinner. Reducing the number of resorption cavities by treatment (Rx) decreases the risk of failure. Reproduced with permission from Reference 84.

Loss of vital trabeculae in this manner could dramatically reduce bone strength, again with relatively little effect on bone mass. This is supported by histomorphometric studies [85–88] showing that, when biopsies from fracture and nonfracture cases are matched for cancellous bone volume, the former show lower values for trabecular number. The importance of architecture in determining bone fragility has recently been confirmed by microcomputed tomography of the proximal femurs from hip fracture patients and controls [89]. Cancellous bone in specimens from fracture victims was found to have significantly greater anisotropy than in controls, even when the two groups were matched for bone mass. The greater anisotropy in the fracture cases was the result of preferential loss of trabeculae perpendicular to the principal loading axis. Several recent studies have shown that agents that lower fracture risk either maintain [90–92] or improve [93, 94] the microarchitecture of human cancellous bone. Secondly, higher turnover rates are accompanied by a shorter bone formation period in each remodeling cycle and also a lower mean age of the matrix. Consequently, mineralization density of the matrix is decreased. This has recently been demonstrated by microradiography of bone samples from baboons that had been ovariectomized and either left untreated or treated with alendronate (Fig. 12) [95]. The degree of mineralization was reduced in the ovariectomized animals and this was partially reversed by alendronate treatment. Furthermore, two recent studies have shown differences in the mineralization density in fracture patients compared to nonfractured controls [96, 97].

Figure 12

Microradiographs of cortical bone from control (CTRL), ovariectomized (OVX) and alendronate-treated (ALN) baboons. The darker osteons are of lower mineral density. Reproduced with permission from Reference 95.

It should be noted that the relationship between the degree of mineralization and the mechanical properties of bone is complex and that the optimal mineralization density value for bone strength has not yet been determined [98]. Increasing mineralization density up to an ash content of 60% by weight increases the bending strength of cortical bone [99], but increasing it beyond that point decreases the ability of the bone to withstand impacts. Bone that is too highly mineralized is brittle and more prone to failure because microfractures propagate more easily through the highly mineralized matrix [100]. In contrast to the studies mentioned above in ovariectomized animals, Boyde et al. [98] found that estrogen withdrawal in young women treated with gonadotrophin-releasing hormone analogs was accompanied by an increase in mineralization density, which the authors felt could contribute to increased skeletal fragility. While it remains to be determined exactly how changes in mineralization density affect bone quality in humans, one can hypothesize that, if the initial mineralization density is low, an increase due to a reduction in remodeling activation could have a beneficial effect on

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bone strength. In support of this concept, treatment with antiresorptive drugs has been shown to reverse the decrease in mineralization density seen in postmenopausal women with osteoporosis [101–103]. A third potential mechanism for a beneficial effect of decreasing bone turnover is a reduction of the probability of a positive feedback loop being established between damage and the porosity associated with repair. Excessive repair of any structure has the potential to be self-defeating. This is particularly true if, as in bone, the repair process initially involves removal of material. As noted above, there is now good evidence that fatigue damage stimulates a remodeling response. With this in mind, it has been hypothesized [104, 105] that the possibility exists for a positive feedback loop between damage and porosity. In this postulate it is suggested that each time bone repairs a microcrack, it creates additional porosity and reduces bone mass. This reduces bone strength, resulting in more microdamage, which in turn initiates another wave of remodeling, which will result in further microdamage and so on and so forth. Martin has used a computer model to simulate this kind of scenario [104]. The model predicts that an increase in loading results in an increase in remodeling activation frequency to repair the increased damage. If loading is below a certain threshold the system is stable and reaches a new steady state. However, if the threshold is exceeded the porosity associated with increased remodeling causes the system to become unstable with damage, porosity, and strain all increasing at a very high rate and without limit. This instability has been suggested to be the theoretical equivalent of a stress fracture. One could propose, therefore, that a reduction in remodeling activation frequency would decrease the probability of this type of vicious circle being set up. In conclusion, although the study of bone remodeling began over 300 years ago with the microscopical observations of Antonie van Leeuwenhoek [106], we still have much to learn about the intricacies and regulation of this fascinating process. In recent years, clinical studies have tended to focus on bone mass, partly because it can be measured noninvasively and with relative ease. It is clear, however, that bone mass gives us only part of the picture. In order to understand how drugs, both those currently available and those under development, prevent fractures we must turn again to techniques that provide information on the remodeling cycle.

References

Acknowledgment This work was supported in part by NIH grants AR 39101. 39191 and 41331.

1. Frost, H. M. (1986). Intermediary Organization Of The Skeleton. CRC Press, Boca Raton, FL. 2. Parfitt, A. M. (2002). Physiologic and pathogenetic significance of bone histomorphometric data. In: “Disorders of Bone and Mineral Metabolism,” F. L. Coe and M. J. Favus (eds), Second Edition, pp. 469–485, Lippincott Williams and Wilkins, Philadelphia. 3. Dempster, D. W. (2002). Bone remodeling. In: “Disorders of Bone and Mineral Metabolism” F. L. Coe and M. J. Favus (eds), Second Edition, Lippincott Williams and Wilkins, Philadelphia, pp. 315–343. 4. Dempster, D. W. (1995). Bone remodeling. In: Osteoporosis: Etiology, Diagnosis, and Management, B. L. Riggs, L. J. Melton III (eds), Second Edition, Raven Press, New York, pp. 67–91. 5. Parfitt, A. M. (1995). Problems in the application of in vitro systems to the study of human bone remodeling. Calcified Tissue Int. 56 (Suppl. 1), S5–7. 6. Roodman, G. D. (1999). Cell biology of the osteoclast. Exp. Hematol. 27, 1229–1241. 7. Burr, D. B. (2002). Targeted and nontargeted remodeling. Bone 30, 2–4. 8. Parfitt, A. M. (2002). Targeted and nontargeted bone remodeling: Relationship to basic multicellular unit origination and progression. Bone 30, 5–7. 9. Roodman, G. D. (2004). Mechanisms of bone metastasis. N. Engl. J. Med. 350, 1655–1664. 10. Khosla, S. (2001). Minireview: The OPG/RANKL/RANK system. Endocrinology 142, 5050–5055. 11. Aubin, J. E., and Bonnelye, E. (2000). Osteoprotegerin and its ligand: a new paradigm for regulation of osteoclastogenesis and bone resorption. Osteoporos. Int. 11, 905–913. 12. Blair, H. C., and Athanasou, N. A. (2004). Recent advances in osteoclast biology and pathological bone resorption. Histol. Histopathol. 19, 189–199. 13. Littlewood-Evans, A., Kokubo, T., Ishibashi, O., Inaoka, T., Wlodarski, B., Gallagher, J. A., and Bilbe, G. (1997). Localization of cathepsin K in human osteoclasts by in situ hybridization and immunohistochemistry. Bone 20, 81–86. 14. Eriksen, E. F. (1986). Normal and pathological remodeling of human trabecular bone: three-dimensional reconstruction of the remodeling sequence in normals and in metabolic bone disease. Endocrine Rev. 7, 379–408. 15. Dempster, D. W., Hughes-Begos, C., Plavetic-Chee, K., Brandao-Burch, A., Cosman, F., Nieves, J., Neubort, S., Lu, S., Iida-Klein, A., Arnett, T., and Lindsay, R. (2005). Normal human osteoclasts formed from peripheral blood monocytes express PTH type 1 receptors and are stimulated by PTH in the absence of osteoblasts. J. Cell. Biochem. 95, 139–148. 16. Wright, K. R., McMillan, P. J., Hughes, D. F., and Boyce, B. F. (1995). Calcium deficiency/repletion in rats as an in vivo model for the study of osteoclast apoptosis. J. Bone Miner. Res. 10, S328. 17. Wu, X., Mckenna, M. A., Feng, X., Nagy, T. R., and Mcdonald, J. M. (2003). Osteoclast apoptosis: The role of Fas in vivo and in vitro. Endocrinology 144, 5545–5555. 18. Baron, R., Vignery, A., and Tran Van, P. (1980). The significance of lacunar erosion without osteoclasts: studies on the reversal phase of the remodeling sequence. Metab. Bone Dis. Rel. Res. 2S, 35–40. 19. Bonewald, L. F., and Mundy, G. R. (1990). Role of transforming growth factor beta in bone remodeling. Clin. Orthop. Rel. Res. 250, 261–276. 20. Mohan, S., and Baylink, D. (1991). Bone growth factors. Clin. Orthop. Rel. Res. 263, 30–48. 21. Hock, J. M., Centrella, M., and Canalis, E. (1998). Insulin-like growth factor I (IGF-I) has independent effects on bone matrix formation and cell replication. Endocrinology 122, 254–260. 22. Fiedler, J., Roderer, G., Gunther, K. P., and Brenner, R. E. (2002). BMP-2, BMP-4, and PDGF-bb stimulate chemotactic migration of

Chapter 22 New Concepts in Bone Remodeling

23.

24.

25.

26.

27. 28.

29.

30. 31.

32.

33.

34.

35. 36.

37.

38.

39. 40.

41. 42.

43.

primary human mesenchymal progenitor cells. J. Cell. Biochem. 87, 306–312. Tanaka, H., Wakisaka, A., Ogasa, H., Kawai, S., and Liang, C. T. (2003). Effects of basic fibroblast growth factor on osteoblast-related gene expression in the process of medullary bone formation induced in rat femur. J. Bone Miner. Metab. 21, 74–79. Locklin, R. M., Oreffo, R. O., and Triffitt, J. T. (1999). Effects of TGFbeta and bFGF on the differentiation of human bone marrow stromal fibroblasts. Cell Biol. Int. 23, 185–194. Lian, J. B., Javed, A., Zaidi, S. K., Lengner, C., Montecino, M., van Wijnen, A. J., Stein, J. L., and Stein, G. S. (2004). Regulatory controls for osteoblast growth and differentiation: role of Runx/Cbfa/AML factors. Crit. Rev. Eukaryot. Gene. Expr. 14, 1–41. Pfeilschifter, J., Diel, I., Scheppach, B., Bretz, A., Krempien, R., Erdmann, J., Schmidd, G., Reske, N., Bismar, H., Seck, T., Krempien, B., and Ziegler, R. (1998). Concentration of transforming growth factor beta in human bone tissue: relationship to age, menopause, bone turnover, and bone volume. J. Bone Miner. Res. 13, 716–730. Smit, T. H., and Burger, E. H. (2000). Is BMU-coupling a strain-regulated phenomenon? A finite element analysis. J. Bone Miner. Res. 15, 301–307. Huiskes, R., Ruimerman, R., van Lenthe, G. H., and Janssen, J. D. (2000). Effects of mechanical forces on maintenance and adaptation of form in trabecular bone. Nature 405, 704–706. Burger, E. H., Klein-Nulend, J., and Smit, T. H. (2003). Strain-derived canalicular fluid flow regulates osteoclast activity in a remodeling osteon – a proposal. J. Biomech. 36, 1452–1459. Marotti, G. (1996). The structure of bone tissues and the cellular control of their deposition. Ital. J. Anat. Embryol. 101, 25–79. Civitelli, R., Beyer, E. C., Warlow, P. M., Roberstson, A. J., Geist, S. T., and Steinberg, T. H. (1993). Connexin 43 mediates direct intercellular communication in human osteoblastic cell networks. J. Clin. Invest. 91, 1888–1896. Dobnig, H., and Turner, R. T. (1995). Evidence that intermittent treatment with parathyroid hormone increases bone formation in adult rats by activation of bone lining cells. Endocrinology 136, 3632–3638. Chow, J. W., Wilson, A. J., Chambers, T. J., and Fox, S. W. (1998). Mechanical loading stimulates bone formation by reactivation of bone lining cells in 13-week-old rats. J. Bone Miner Res. 13, 1760–1767. Parfitt, A. M. (1994). Osteonal and hemiosteonal remodeling: the spatial and temporal framework for signal traffic in adult bone. J. Cell. Biochem. 55, 273–276. Currey, J. D. (2002). Bones: Structure and Mechanics. Princeton University Press, Princeton, NJ. Banks, W. J., Epling, G. P., Kainer, R. A., and Davio, R. W. (1968). Antler growth and osteoporosis. I. Morphological and morphometric changes in the costal compacta during the antler growth cycle. Anat. Rec. 162, 387–398. Barzel, U. S. (1995). The skeleton as an ion exchange system: implications for the role of acid-base imbalance in the genesis of osteoporosis. J. Bone Miner. Res. 10, 1431–1436. Burr, D.B., Martin, R.B., Schaffler, M.B., and Radin, E.L. (1985). Bone remodeling in response to in vivo fatigue microdamage. J. Biomech. 18, 189–200. Mori, S., Burr, D. B. (1993). Increased intracortical remodeling following fatigue damage. Bone 14, 103–109. Burr, D. B., Forwood, M. R., Fyhrie, D. P., Martin, R. B., Schaffler, M. B., and Turner, C. H. (1997). Perspective: Bone microdamage and skeletal fragility in osteoporotic and stress fractures. J. Bone Miner. Res. 12, 6–15. Martin, R. B., and Burr, D. B. (1989). Structure, Function, and Adaptation of Compact Bone. Raven Press, New York. Kerr, J. F. R., Wyllie, A. H., and Currie, A. R. (1972). Apoptosis: a basic biological phenomenon with wide-ranging implications in tissue kinetics. Brit. J. Cancer 26, 239–257. Wyllie, A. H., Kerr, J. F. R., and Currie, A. R. (1980). Cell death: the significance of apoptosis. Int. Rev. Cytol. 68, 258–306.

387 44. Parfitt, A. M., Mundy, G. R., Roodman, G. D., Hughes, D. E., and Boyce, B. F. (1996). Theoretical perspective: a new model for the regulation of bone resorption, with particular reference to the effects of bisphosphonates. J. Bone Miner. Res. 11, 150–159. 45. Boyce, B. F., Hughes, D. E., and Wright, K. R. (1997). Methods for studying cell death in bone. In: “Methods in Bone Biology,” T. R. Arnett, B. Henderson (eds), pp. 127–148, Chapman and Hall, London. 46. Dempster, D. W., Moonga, B. S., Stein, L. S., Horbert, W. R., and Antakly, T. (1997). Glucocorticoids inhibit bone resorption by isolated rat osteoclasts by enhancing apoptosis. J. Endocrinol. 154, 397–406. 47. Hughes, D. E., and Boyce, B. F. (1997). Apoptosis in bone physiology and disease. J. Clin. Pathol. Mol. Pathol. 50, 132–137. 48. Furtwangler, J. A., Hall, S. H., and Koskine-Moffett, L. K. (1985). Sutural morphogenesis in the mouse calvaria: the role of apoptosis. Acta. Anat. 124, 74–80. 49. Hock, J. M., Krishnan, V., Onyia, J. E., Bidwell, J. P., Milas, J., and Stanislaus, D. (2001). Perspective: Osteoblast apoptosis and bone turnover. J. Bone Miner. Res. 16, 975–984. 50. Weinstein, R. S., and Manolagas, S. C. (2000). Apoptosis and osteoporosis. Am. J. Med. 108, 153–164. 51. Hughes, D. E., Dai, A., Tiffee, J. C., Li, H. H., Mundy, G. R., and Boyce, B. F. (1996). Estrogen promotes apoptosis of murine osteoclasts mediated by TGF-β. Nat. Med. 2, 1132–1136. 52. Kameda, T., Mano, H., Yuasa, T., Mori, Y., Miyazawa, K., Shiokawa, M., Nakamura, Y., Hiroi, E., Hiura, K., Kameda, A., Yang, N. N., Hakeda, Y., and Kumegawa, M. (1997). Estrogen inhibits bone resorption by directly inducing apoptosis of the bone resorbing osteoclasts. J. Exp. Med. 186, 489–495. 53. Arnett, T., Lindsay, R., Kilb, L., Moonga, B. S., Spowage, M., and Dempster, D. W. (1996). Selective toxic effects of tamoxifen on osteoclasts: comparison with the effects of oestrogen. J. Endocrinol. 149, 503–508. 54. Lutton, J. D., Moonga, B. S., and Dempster, D. W. (1996). Osteoclast demise: physiological versus degenerative cell death. Experimental Physiology 81, 251–260. 55. Grasser, W. A., Pan, L. C., Thompson, D. D., and Paralkar, V. M. (1997). Common mechanism for the estrogen agonist and antagonist activities of droloxifene. J. Cell. Biochem. 65, 159–171. 56. Hughes, D. E., Wright, K. R., Uy, H. L., Sasaki, A., Yoneda, T., Roodman, G. D., Mundy, G. R., and Boyce, B. F. (1995). Bisphosphonates promote apoptosis in murine osteoclasts in vitro and in vivo. J. Bone Miner. Res. 10, 1478–1487. 57. Selander, K. S., Harkonen, P. L., Valve, E., Mönkkönen, J., Hannuniemi, R., and Väänänen, H. K. (1996). Calcitonin promotes osteoclast survival in vitro. Mol. Cell. Endocrinol. 122, 119–129. 58. Fuller, K., Owens, J. M., Jagger, C. J., Wilson, A., Moss, R., Chambers, T. J. (1993). Macrophage colony-stimulating factor stimulates survival and chemotactic behavior in isolated osteoclasts. J. Exp. Med. 178, 1733–1744. 59. Jilka, R. L., Weinstein, R. S., Bellido, T., Parfitt, A. M., Manolagas, S. C. (1998). Osteoblast programmed cell death (apoptosis): modulation by growth factors and cytokines. J. Bone Miner. Res. 13, 793–802. 60. Manusukhani, A., Bellosta, P., Sahni, M., and Basilico, C. (2000). Signaling by fibroblast growth factors (FGF) and fibroblast growth factor 2 (FGFR2)-activating mutations blocks mineralization and induces apoptosis in osteoblasts. J. Cell. Biol. 149, 1297–1308. 61. Hill, P., Tumber, A., and Meikle, M. (1997). Multiple extracellular signals promote osteoblast survival and apoptosis. Endocrinology 138, 3849–3858. 62. Gronowicz, G. A., McCarthy, M. B., Zhang, H., and Zhang, W. (2004). Insulin-like growth factor II induces apoptosis inosteoblasts. Bone 35, 621–628. 63. Kitajima, I., Nakajima, T., Imamura, T., Takasaki, I., Kawahara, K., Okano, T., Tokioka, T., Soejima, Y., Abeyama, K., and Murayama, I. (1996). Induction of apoptosis in murine clonal osteoblasts expressed

388

64.

65.

66. 67.

68.

69.

70.

71.

72.

73.

74.

75.

76.

77.

78.

79.

80.

by human T-cell leukemia virus type I tax by NF-κB and TNF-α. J. Bone Miner. Res. 11, 200–210. Tsuboi, M., Kawakami, A., Nakashima, T., Matsuoka, N., Urayama, S., Kawabe, Y., Fujiyama, K., Kiriyama, T., Aoyagi, T., Maeda, K., and Euguchi, K. (1999). Tumor necrosis factor-alpha and interleukin-1beta increase the Fas-mediated apoptosis of human osteoblasts. J. Lab. Clin. Med. 134, 222–231. Weinstein, R. S., Jilka, R. L., Parfitt, A. M., and Manolagas, S. C. (1998). Inhibition of osteoblastogenesis and promotion of apoptosis of osteoblasts and osteocytes by glucocorticoids. Potential mechanisms of their deleterious effects on bone. J. Clin. Invest. 102, 274–282. Dempster, D.W. (1989). Bone histomorphometry in glucocorticoidinduced osteoporosis. J. Bone Min. Res. 4, 137–141. King, C. S., Weir, C. E., Gundberg, C. W., Fox, J., and Insogna, K. L. (1996). Effects of continuous glucocorticoid infusion on bone metabolism in the rat. Calcif. Tissue Int. 59, 184–191. Li, M., Shen, Y., Halloran, B. P., Baumann, B. D., Miller, K., and Wronski, T. J. (1996). Skeleton response to corticosteroid deficiency and excess in growing male rats. Bone 19, 81–88. Shen, V., Birchman, R., Liang, X. G., Wu, D. D., Lindsay, R., and Dempster, D. W. (1997). Prednisolone alone, or in combination with estrogen or dietary calcium deficiency or immobilization, inhibits bone formation but does not induce bone loss in mature rats. Bone 21, 345–351. Noble, B. S., Peet, N., Stevens, H. Y., Brabbs, A., Mosley, J. R., Reilly, G. C., Reeve, J., Skerry, T. M., and Lanyon, L. E. (2003). Mechanical loading: biphasic osteocyte survival and targeting of osteoclasts for bone destruction in rat cortical bone. Am. J. Physiol. Cell Physiol. 284, C934–C943. Verborgt, O., Tatton, N. A., Majeska, R. J., and Schaffler, M. B. (2002). Spatial distribution of Bax and Bcl-2 in osteocytes after bone fatigue: complementary roles in bone remodeling regulation? J. Bone Miner. Res. 17, 907–914. Verborgt, O., Gibson, G. J., and Schaffler, M. B. (2000). Loss of osteocyte integrity in associated with microdamage and bone remodeling after fatigue in vivo. J. Bone Miner. Res. 15, 60–67. Noble, B. S., Stevens, H., Loveridge, N., and Reeve, J. (1997). Identification of apoptotic changes in osteocytes in normal and pathological bone. Bone 20, 273–282. Ehrlich, P. J., Noble, B. S., Jessop, H. L., Stevens, H. Y., Mosley, J. R., Lanyon, L. E. The effect of in vivo mechanical loading on estrogen receptor alpha expression in rat ulnar osteocytes. J. Bone Miner. Res. 2002, 17:1646–1655. Bronckers, A. L. J. J., Goei, W., Luo, G., Karsenty, G., D’Sousa, R. N., Lyaruu, D. M., and Burger, E. H. (1996). DNA fragmentation during bone formation in neonatal rodents assessed by transferase-mediated end labeling. J. Bone Miner. Res. 11, 1281–1291. Flora, L., Hassing, G. S., Cloyd, G. G., Bevan, J. A., Parfitt, A. M., and Villanueva, A. R. (1981). The long term skeletal effects of EHDP in dogs. Metab. Bone Dis. Relat. Res. 3, 289–300. Mashiba, T., Hirano, T., Turner, C. H., Forwood, M. R., Johnston, C. C., and Burr, D. B. (2000). Suppressed bone turnover by bisphosphonates increases microdamage accumulation and reduces some biomechanical properties in dog rib. J. Bone Miner. Res. 15, 613–620. Garnero, P., Hausherr, E., Chapuy, M-C., Marcelli, C., Grandjean, H., Muller, C., Cormier, C., Breart, G., Meunier, P. J., and Delmas, P. D. (1996). Markers of bone resorption predict hip fracture in elderly women: the EPIDOS prospective study. J. Bone Miner. Res. 11, 1531–1538. van Daele, P. L. A., Seibel, M. J., Burger, H., Hofman, A., Grobbee, D. E., van Leeuwen, J. P. T. M., Birkenhäger, J. C., and Pols, H. A. P. (1996). Case-control analysis of bone resorption markers, disability and hip fracture risk: the Rotterdam study. Br. Med. J. 312, 482–483. Riggs, B. L., Hodgson, S. F., O’Fallon, W. M., Chao, E. Y. S., Wahner, H. W., Muhs, J. M., Cedel, S. L., and Melton, L. J. III (1990).

DAVID W. DEMPSTER AND HUA ZHOU

81.

82.

83.

84. 85.

86.

87.

88.

89.

90.

91.

92.

93.

94.

95.

96.

97.

Effect of fluoride treatment on the fracture rate in postmenopausal women with osteoporosis. N. Engl. J. Med. 322, 802–809. Kleerekoper, M., Peterson, E. L., Nelson, D. A., Phillips, E, Schork, M. A., Tilley, B. C., and Parfitt, A. M. (1991). A randomized trial of sodium fluoride as a treatment for postmenopausal osteoporosis. Osteoporosis Int. 1, 155–161. Sogaard, C. H., Mosekilde, L, Richards, A, and Mosekilde, L. (1994). Marked decrease in trabecular bone quality after five years of sodium fluoride therapy; assessed by biomechanical testing of iliac crest bone biopsies in osteoporotic patients. Bone 15, 393–399. Cummings, S. R., Karpf, D. B., Harris, F., Genant, H. K., Ensrud, K., LaCroix, A. Z., and Black, D. M. (2002). Improvement in spine bone density and reduction in risk of vertebral fractures during treatment with antiresorptive drugs. Am. J. Med. 112, 281–289. Parfitt, A. M. (1991). Use of bisphosphonates in the prevention of bone loss and fractures. Am. J. Med. 91, (Suppl 5B), 42S–46S. Kleerekoper, M., Villanueva, A. R., Stanciu, J., Rao, D. S., and Parfitt, A. M. (1985). The role of three dimensional trabecular microstructure in the pathogenesis of vertebral compression fractures. Calcified Tissue Int. 37, 594–597. Legrand, E., Chappard, D., Pascaretti, C., Duquenne, M., Krebs, S., Rohmer, V., Basle, M. F., and Audran, M. (2000). Trabecular bone microarchitecture, bone mineral density, and vertebral fractures in male osteoporosis. J. Bone Miner. Res. 15, 13–19. Aaron, J. E., Shore, P. A., Shore, R. C., Beneton, M., and Kanis, J. A. (2000). Trabecular architecture in women and men of similar bone mass with and without vertebral fracture: II. Three-dimensional histology. Bone 27, 277–282. Oleksik, A., Ott, S. M., Vedi, S., Bravenboer, N., Compston, J., and Lips, P. (2000). Bone structure in patients with low bone mineral density with or without vertebral fractures. J. Bone Miner. Res. 15, 1368–1375. Ciarelli, T. E., Fyhrie, D. P., Schaffler, M. B., and Goldstein, S. A. (2000). Variations in three-dimensional cancellous bone architecture of the proximal femur in female hip fractures and controls. J. Bone Miner. Res. 15, 32–40. Borah, B., Dufresne, T. E., Chmielewski, P. A., Johnson, T. D., Chines, A., Manhart, M. D. (2004). Risedronate preserves bone architecture in postmenopausal women with osteoporosis as measured by threedimensional microcomputed tomography. Bone 34, 736–746. Dufresne, T. E., Chmielewski, P. A., Manhart, M. D., Johnson, T. D., and Borah, B. (2003). Risedronate preserves bone architecture in early postmenopausal women in 1 year as measured by three-dimensional microcomputed tomography. Calcif. Tissue Int. 73, 423–432. Recker, R., Masarachia, P., Santora, A., Howard, T., Chavassieux, P., Arlot, M., Rodan, G., Wehren, L., Kimmel, D. (2005). Trabecular bone microarchitecture after alendronate treatment of osteoporotic women. Curr. Med. Res. Opin. 21, 185–194. Dempster, D. W., Cosman, F., Kurland, E. S., Zhou, H., Nieves, J., Woelfert, L., Shane, E., Plavetic, K., Müller, R., Bilezikian, J. P., and Lindsay, R. (2001). Effects of daily treatment with parathyroid hormone on bone microarchitecture and turnover in patients with osteoporosis: a paired biopsy study. J. Bone Miner. Res. 16, 1846–1853. Jiang, Y., Zhao, J. J., Mitlak, B. H., Wang, O., Genant, H. K., and Eriksen, E. F. (2003). Recombinant human parathyroid hormone (1–34) [teriparatide] improves both cortical and cancellous bone structure. J. Bone Miner. Res. 18, 1932–1941. Meunier, P. J., and Boivin, G. (1997). Bone mineral density reflects bone mass but also the degree of mineralization of bone: therapeutic implications. Bone 21, 373–377. Ciarelli, T. E., Fyhrie, D. P., and Parfitt, A. M. (2003). Effects of vertebral bone fragility and bone formation rate on the mineralization levels of cancellous bone from white females. Bone 32, 311–315. Loveridge, N., Power, J., Reeve, J., and Boyde, A. (2004). Bone mineralization density and femoral neck fragility. Bone 35, 929–941.

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Chapter 22 New Concepts in Bone Remodeling 98. Boyde, A., Compston, J. E., Reeve, J., Bell, K. L., Noble, B. S., Jones, S. J., and Loveridge, N. (1998). Effect of estrogen suppression on the mineralization density of iliac crest biopsies in young women as assessed by backscattered electron imaging. Bone 22, 241–250. 99. Currey, J. D. (1969). The mechanical consequences of variation in the mineral content of bone. J. Biomech. 2, 1–11. 100. Currey, J. D., Brear, K., and Zioupos, P. (1995). The effects of aging and changes in the mineral content in degrading the toughness of human femora. J. Biomech. 29, 257–260. 101. Roschger, P., Rinnerthaler, S., Yates, J., Rodan, G. A., Fratzl, P., and Klaushofer, K. (2001). Alendronate increases degree and uniformity of mineralization in cancellous bone and decreases the porosity in cortical bone of osteoporotic women. Bone 29, 185–191. 102. Boivin, G., Vedi, S., Purdie, D. W., Compston, J. E., and Meunier, P. J. (2005). Influence of estrogen therapy at conventional and high doses on the degree of mineralization of iliac bone tissue: a quantitative microradiographic analysis in postmenopausal women. Bone 36, 562–567. 103. Boivin, G., Lips, P., Ott, S. M., Harper, K. D., Sarkar, S., Pinette, K. V., and Meunier, P. J. (2003). Contribution of raloxifene and calcium and

104. 105.

106.

107.

108.

vitamin D3 supplementation to the increase of the degree of mineralization of bone in postmenopausal women. J. Clin. Endocrinol. Metab. 88, 4199–4205. Martin, B. (1995). Mathematical model for repair of fatigue damage and stress fracture in osteonal bone. J. Orthop. Res. 13, 309–316. Burr, D. B., Forwood, M. R., Fyhrie, D. P., Martin, R. B., Schaffler, M. B., and Turner, C. H. (1997). Perspective: Bone microdamage and skeletal fragility in osteoporosis and stress fractures. J. Bone Miner. Res. 12, 6–15. Leeuwenhoek, A. (1678). Microscopical observations on the structure of teeth and other bones. Philos. Trans. R. Soc. Lond. 12, 1002–1003. Murrills, R. J., Shane, E., Lindsay, R., and Dempster, D. W. (1989). Bone resorption by isolated human osteoclasts in vitro: effects of calcitonin. J. Bone Miner. Res. 4, 259–268. Dempster, D. W., Shane, E. S., Horbert, W., and Lindsay, R. (1986). A simple method for correlative light and scanning electron microscopy of human iliac crest bone biopsies: qualitative observations in normal and osteoporotic subjects. J. Bone Miner. Res. 1, 15–21.

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Chapter 23

Products of Bone Collagen Metabolism Juha Risteli and Leila Risteli

Department of Clinical Chemistry, FI-90014 University of Oulu, Oulu, Finland

I. Introduction II. Products of Bone Collagen Synthesis, the Procollagen Propeptides

III. Degradation Products of Type I Collagen IV. Closing Remarks References

I. INTRODUCTION

on chromosome 7. The basic pathways of collagen synthesis and degradation are shared by all tissues, but there are certain features that are more pronounced, either in the hard or the soft tissues, allowing some relative distinction to be made between these two sources of type I collagen metabolites. The main cells synthesizing the type I collagen in soft tissues are fibroblasts, which also produce significant amounts of type III collagen. Bone collagen is synthesized by osteoblasts, which normally do not express type III collagen [6]. However, certain established cell lines with an apparent osteoblastic nature (e.g., the human osteosarcoma line MG-63) produce large amounts of type III collagen [7]. Several other cell types (e.g., smooth muscle cells) also synthesize both type I and III collagens. They also produce several other collagens, the amounts of which are far less than those of the type I and III collagens. The assembly of type I collagen molecules into fibrils differs somewhat between bones and skin and this arrangement affects the cross-linking of the collagen molecules in collagen fibers, in particular at their carboxytermini [8]. However, the fibrillar collagens of many soft tissues, such as tendons, fasciae, vessel walls, and internal organs, contain cross-links similar to those present in bone.

Most of the collagen in the organic matrix of bone is type I collagen, which provides a well-organized and insoluble scaffold for the deposition of the mineral. Although the mineral is even more abundant than collagen, it is difficult and tedious to reliably assess its metabolism. This leaves type I collagen as the best target for quantification of the metabolic turnover in the skeleton [1–3]. Τhe synthesis of type I collagen involves the production of specific by-products that, among other things, provide the possibility of elegantly assessing the rate of the synthesis of this collagen. Another set of metabolic products is related to the degradation of this collagen, which can occur either together with the dissolution of the mineral phase or independently, e.g., in situations like rickets, when there is increased breakdown of a nonmineralized matrix [4–5]. Although most of the type I collagen in the human body is located in the skeleton, this protein is also the most abundant collagen in soft tissues. Unfortunately, there are no good quantitative estimates for the distribution of type I collagen between these two pools, either in health or in disease. In all locations the protein is encoded by the same genes, the COL1A1 on chromosome 17 and the COL1A2 Dynamics of Bone and Cartilage Metabolism

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The mineralization of bone also affects the maturation of bone collagen cross-links; thus many of them remain immature, divalent in nature, and a significant part of the α1-chains in the bone collagen molecules that contain potential cross-linking sites even remain uncross-linked [9]. Another difference between the soft tissues and bones is the fact that bone metabolites can be directly released into the circulation, whereas most soft tissues are first drained via lymphatics into larger vessels. Since there is much microheterogeneity in type I collagen, it is not possible using only biochemical means to predict whether a certain metabolic product, present either in the blood or in the urine, will predominantly reflect bone metabolism. More well-focused studies are still needed, e.g., experimental work on the structure and physiology of the collagens of various tissues, as well as clinical studies of different diseases. Detection of most of the metabolic products of collagen metabolism depends on the use of well-characterized immunological reagents, and proper quantification, in principle, can be achieved with a number of techniques, including RIAs, ELISAs, IRMAs, etc. However, proper immunochemical and biological validation is always necessary; it is particularly relevant if the assay is just based on a linear synthetic peptide, as the natural collagen metabolites are relatively complex structures. Conclusions based on assays not properly validated may introduce confusion in medical literature.

II. PRODUCTS OF BONE COLLAGEN SYNTHESIS, THE PROCOLLAGEN PROPEPTIDES A. Biochemical Basis The biosynthesis of fibrillar collagens is a complex chain of events that includes the release of two additional proteins, known as the propeptides of the respective procollagen. These proteins seem to be largely removed from their sites of origin without further degradation and thus offer the potential of directly measuring the rate of collagen synthesis in a manner analogous to the use of the C-peptide of proinsulin as an indicator of endogenous insulin production. Interestingly, the two proteins removed from the two ends of the rod-like collagen molecule differ from each other profoundly in their chemical natures and their further metabolic fates (see also Chapter 1). The type I collagen molecule is a long, thin, rigid rod – a shape necessary for its function as part of a collagen fiber in the tissue. The best known and most abundant form of this collagen, the “classical” type I collagen, is a

heterotrimer of two α1(I) chains and one α2(I) chain, which are wrapped around each other into the triple helix. The original gene products, the proα1(I) and proα2(I) chains of type I procollagen, are about 50% longer than the corresponding final products. The two additional, bulky domains at both ends of the molecule are usually called the amino-terminal (abbreviation PINP) and the carboxyterminal (PICP) propeptides of type I procollagen (Fig. 1) [10], despite their relatively large sizes, which are clearly outside the ordinary definition of a peptide (see Sections II.B and C). Once the molecule has reached the extracellular space, these parts are removed en bloc from the procollagen by two specific endoproteinases, the N- and C-proteinase, the latter of which is identical to BMP-1 [11]. Several functions have been proposed for the propeptide domains in the procollagen molecule (for N-propeptide, see reference [12]). The individual polypeptide chains are expressed separately, with the most amino-terminal signal peptide and propeptide parts being synthesized first, followed by the part that will form the collagen molecule itself, and followed finally by the carboxy-terminal propeptide moiety. As the chains pass into the cisternae of the endoplasmic reticulum, they are, at the same time, being post-translationally modified by several enzymes [10]. The carboxy-terminal propeptide sequences are needed for the proper selection and association of the three proαchains of type I procollagen within the cisternae of the endoplasmic reticulum. The collagen helix is formed starting from the carboxy-terminus and proceeding in the amino-terminal direction. The amino-terminal propeptide is thus the last part of the molecule to attain its native conformation, which includes an additional short collagenous triple helix within this domain. During the further transit of the procollagen, its propeptide domains are believed to prevent premature fiber formation in the cells and in the extracellular space, since there is a 1000-fold difference in solubility between procollagen and collagen. The carboxy-terminal propeptide can accomplish this on the basis of its bulky, globular form, whereas, in the amino-terminal propeptide, a similar function is probably due to its negative charge, analogous to the situation in blood fibrinogen (see Section II.C). When the carboxy-terminal propeptide has been removed, the molecules still containing the amino-terminal propeptide (so-called type I pN-collagen) can be layered onto the collagen fibrils, but just on the surface of the fibril. A delayed cleavage of the amino-terminal propeptide is believed to regulate the lateral growth of the type I collagen fibers [13]. In addition to the classical type I collagen that is most abundant both in the skeleton and in the soft tissues, there is some evidence for the existence of other molecular forms

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Figure 1

Schematic presentation of a procollagen molecule. The parts of the N-terminal propeptide are Col 1 (globular domain), Col 2 (nontriple-helical domain) and Col 3 (triple-helical domain). The nontriple-helical domains at both ends of the collagen molecule are the telopeptides. Reproduced from Prockop et al. [10], Copyright © 1979 Massachusetts Medical Society. All rights reserved.

of type I collagen. A type I α1-trimer collagen, which consists of three α1(I) chains, is the main collagen in the rare cases of osteogenesis imperfecta in which this disease is due to a genetic lack of the α2(I) chain [14]. Similar trimeric collagen has also been found in osteoarthritic bone [15]. There is evidence that some type I α1(I)-trimer collagen is also formed in people having a G to T substitution in the Sp1 binding site in the COL1A1 gene [16] (see also Chapter 30).

B. Carboxy-terminal Propeptide of Type I Procollagen (PICP) The carboxy-terminal propeptide – both in free form and as part of the type I procollagen molecule – has a globular shape and an amino acid composition of a noncollagenous protein. The parts corresponding to the propeptide are encoded by exons 49 through 52 of the COL1A1 and

Table I.

COL1A2 genes. Its overall molecular mass is about 100 000 (Table I), and those of the subunits about 33 000 each. All the three component polypeptide chains contain an N-glycosylation site, which is normally occupied by an oligosaccharide of the high-mannose type [17]. For this reason, the propeptide is bound to an affinity column of immobilized concanavalin A, a property that can be used in isolating the propeptide [18]. There are also interchain disulfide bridges between its three polypeptide chains; these are needed in the processes of chain selection and association. There also seems to be a free SH-group, since the propeptide can also be purified using an affinity column binding such groups [19]. It is relatively difficult to isolate the PICP antigen, but it has been purified from the culture media of human fibroblasts, either in the form of type I procollagen, from which the PICP has been liberated by bacterial collagenase digestion [18, 20], or directly as a propeptide cleaved in cell culture, starting from a serumfree culture medium [19].

Comparison of the Amino-terminal (PINP) and Carboxy-terminal (PICP) Propeptides of the Classical Human Type I Procollagen PINP

Location Relative molecular mass Shape Chemical nature Related serum antigen Homogeneity Size Concentration (µg/L) Clearance from blood Blocked by

PICP

Amino-terminal 35 000 Elongated Phosphorylated, partially collagenous

Carboxy-terminal 100 000 Globular Glycoprotein, oligosaccharides of the high-mannose type

One major and one minor form Same as intact PINP (major) or Col1-like (minor) Men: 20–76 Women: 19–84 Scavenger receptor of liver endothelial cells Formaldehyde-treated albumin, acetylated LDL, PIIINP

One form Same as PICP 38–202 50–170 Mannose receptor of liver endothelial cells Ovalbumin, mannan

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C. Amino-terminal Propeptide of Type I Procollagen (PINP) The aminoterminal propeptide contains a collagenous domain and thus has the shape of a short rod ending with a more globular part (Fig. 1). The subdomains of the propeptide have been named on the basis of their order of elution from a size-exclusion column after bacterial collagenase digestion. From the largest to the smallest, they are the following: Col 1 (the globular amino-terminal end with intrachain disulfide bridges); Col 2 (the short noncollagenous part joining the propeptide to the most amino-terminal part of the collagen proper); and Col 3 (the central collagenous domain that is broken down by bacterial collagenase). The parts corresponding to the propeptide are encoded by exons 1 through 6 of the COL1A1 and COL1A2 genes. The molecular mass of the propeptide is about 35 000 daltons (Table I), and masses of the subunits are about 14 250 daltons for the proα1 chain and 5500 daltons for the proα2 chain. The proα2 chain of the propeptide contains the collagenous domain (Col3) and the short carboxyterminal noncollagenous sequence (Col 2), but lacks the globular amino-terminal end (Col 1 domain). The length of the triple helix in the propeptide is 16 Gly-X-Y triplets – this is less than 5% of the length of the helical domain in the collagen molecule itself, which contains 338 such triplets. The globular Col 1 domain is phosphorylated in the type I procollagens produced by both fibroblasts and osteoblasts, the phosphate being linked to serine as phosphoserine [21]. The acidic nature of the propeptide is advantageous when the protein is being isolated from various biological starting materials, such as pleural effusions or ascitic fluids, which often contain high concentrations of the propeptide [22]. Because the amino-terminal propeptide of the α1-trimer type I procollagen contains three globular Col 1 domains, it is more acidic than the propeptide of the classical type I procollagen with its two Col 1 domains and thus elutes later in anion exchange chromatography [23].

D. Clearances of the Circulating PICP and PINP In comparison with the large amounts of the propeptides set free in the body, their circulating concentrations are surprisingly low, only 20–200 µg/L, which is in the nanomolar range (Table I). This fact suggests the existence of efficient removal mechanisms for this material. In fact, the further metabolic fates of both PICP and PINP are now known better than those of most other analytes used routinely in clinical medicine. The propeptides are

not passively lost in the urine because they are either too large (PICP) or have such an elongated shape (PINP) that they cannot pass through the glomerular filter. In addition, the negative charge of the latter, due to the covalently attached phosphate groups, obviously repels the propeptide from the negatively charged membrane. The metabolic fates of the free propeptides have been clarified by several types of experiments, such as by following the disappearance of radioactively labeled or fluorescent propeptides from the circulation in rats, by feeding the proteins to cultured cells in the presence of known competing ligands for various membrane receptors, by simultaneous measurement of the propeptide concentrations in different vascular beds, and, recently, also by studying the effect of receptor knockout on the circulating propeptide concentration in mice (Fig. 2) [24]. The propeptides are actively taken up and metabolized by a specialized part of the reticulo-endothelial system, the endothelial cells of the liver [25]. These cells bind and internalize the propeptides by receptor-mediated endocytosis using two different receptor systems, the mannose receptor for PICP [26] and the scavenger receptor for PINP [27]. The clearance follows biphasic kinetics, with half-lives measured in minutes in the rat. The endocytosed material is delivered to the lysosomes and degraded, with the resulting amino acids most probably being passed to the hepatocytes. The evolution of such a clearance mechanism could be related to the importance of having an effective recycling system for the large amounts of amino acids in the propeptides. The amount of type I collagen synthesized per year in adult humans has been estimated to be at least 1 kg [28], and for each kilogram of type I collagen, as much as 500 g of propeptide material is also produced. The oligosaccharides on PICP are of the high-mannose type that is considered relatively primitive; a special function related to such recycling could explain this feature, which has no effect on the functions or processing of the procollagen [29]. The two clearance systems, although present in the same cells, are independent of each other and also seem to be regulated separately. Some hormones, e.g., thyroxine, can affect the activities of these endocytotic systems and even sometimes cause opposite changes in the circulating concentrations of the procollagen propeptides [30]. However, the effects of such endocrinological regulation are relatively small. In contrast, at least one family is known that has an obvious genetic defect in the uptake mechanism of PICP [31]. The circulating PICP concentrations of the affected family members tend to be several standard deviations above the upper limits of the age- and sex-adjusted reference intervals, with no other abnormalities in either type I collagen metabolites or other biochemical markers of bone metabolism. The feature is inherited in an autosomal

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Figure 2

Proteomic analysis of total serum proteins in mice, two-dimensional polyacrylamidegel electrophoresis under reducing conditions. The gels were stained with SYPRO Ruby and scanned on a laser fluorescence scanner. WT = wild-type mice; KO = mannose receptor knockout mice; pI = isoelectric point; MW = molecular weight. The spot nos. 1–4 represent the proteins that were constantly found to be elevated in the knockout samples. The spots nos. 1–3 were identified by peptide mass fingerprinting as the carboxy-terminal propeptide domains of the proα1(I) and proα2(I) chains of type I procollagen and the proα1(III) chain of type III procollagen. Reproduced from Lee et al. [24], with permission.

dominant manner (Fig. 3) and is not associated with any obvious signs or symptoms. Interestingly, such grossly elevated PICP concentrations also react to factors known otherwise to increase or decrease this concentration, such as pregnancy [32] or glucocorticoid therapy [31]. Studies have shown also that the local PICP concentration in suction blisters of the skin (see Section II.E) is similarly elevated in these individuals, suggesting that there normally may also be some local uptake of PICP, at least in the soft tissues, possibly by tissue macrophages carrying mannose receptors [33].

practically noninvasively testing the validity for any new test assumed to detect type I collagen synthesis. The relative contribution of soft-tissue type I collagen synthesis to the circulating concentration of PICP has been studied in lymph flow experiments with conscious pigs [36]. There is a gradient of 10:1 from lymph to peripheral

E. Comparison of PICP and PINP in Physiological and Clinical Situations The local synthesis of soft-tissue type I procollagen can be assessed in human skin, either by collecting the interstitial fluid in surgical wounds [34] or by a suction blister method [35]. Such studies have shown that the propeptides of type I procollagen are efficiently set free from the collagen in the extracellular fluid. Their concentrations increase dramatically (up to 1000-fold within 1 week) during wound healing, when the expression of type I procollagen is induced [22, 34]. On the other hand, local treatment with glucocorticoids suppresses the synthesis of this collagen, leading to often dramatic decreases in the concentrations of the propeptides measured in the fluid in suction blisters that are made in the treated area of the skin [35]. Such experiments provide excellent means of reproducibly and

Figure 3

Family with inherited high circulating concentrations of the carboxy-terminal propeptide (PICP) of type I procollagen. The circles and squares indicate women and men, respectively. The open symbols show unaffected family members, whereas the solid symbols present individuals with high PICP concentrations. The arrow indicates the proband. AP, alkaline phosphatase; BAP, bone-specific alkaline phosphatase; OC, osteocalcin. Reproduced from Sorva et al. [31], with permission.

396 blood in the concentration of the amino-terminal propeptide of type III procollagen (PIIINP; soft-tissue origin), whereas there are no significant differences between the concentrations of PICP in lymph and blood. Stopping and restarting the lymph flow produces dramatic changes in the concentration of PIIINP in the blood, whereas there are no changes in the corresponding concentration of PICP. This study demonstrates that even large changes in the access of the products of soft-tissue type I collagen synthesis into the circulation have little effect on the blood concentration of PICP. This is not surprising because we are, in fact, not measuring the absolute amounts of the proteins in any pool in the body, but just their concentrations, and before mixing the lymph into the blood there is no gradient in this case. Because soft tissues have little effect on blood, the bone formation rate is obviously the main determinant of the concentrations of the type I procollagen propeptides in the blood (Fig. 4). This view is also supported by

Figure 4

Schematic presentation of the distribution between various pools and of the clearances of the type I procollagen propeptides. PICP, the carboxy-terminal propeptide; Intact PINP, the intact triple-helical aminoterminal propeptide of type I procollagen; PINP Col 1, the smaller, singlechain antigen related to the Col 1 domain of the proα1 chain, recognized by some assays for PINP. Thick arrows indicate the major routes. Reproduced from Risteli et al. [2], with permission.

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histomorphometric and calcium kinetic studies in patients [37, 38]. In the blood, the carboxy-terminal propeptide (PICP) is always found as the free antigen, with the size of the authentic propeptide [18], whereas the circulating antigenicity related to the amino-terminal propeptide (PINP) can be resolved into two peaks with different molecular sizes. The first is identical to the free, authentic trimeric antigen, and the second resembles part of a single polypeptide chain, probably containing at least the globular Col 1 domain of the proα1(I) chain of PINP [22]. PINP can be isolated as an intact, trimeric propeptide from pleural and ascitic fluids [22], whereas the corresponding propeptide isolated from amniotic fluid always seems to fall apart, into single, possibly truncated, chains [39, 40]. Different PINP assays may vary with respect to the extent to which their results are influenced by the smaller antigens. The corresponding propeptide of type III procollagen, PIIINP, is also present in the blood as a series of antigens with partially different metabolic origins and differing metabolic fates [1, 3]. The origin of the smaller antigenic forms of PINP is most probably the tissue degradation of newly synthesized type I procollagen or of the type I pN-collagen with the retained propeptide (Section III.A). Thus the exclusive assay of the intact PINP seems to be more sensitive than that of total PINP for detecting changes in the rate of bone collagen synthesis (e.g., estrogen treatment in postmenopausal women leads to a 42% decrease in the circulating concentration of intact PINP, compared with the 30% decrease observed with an assay measuring both antigenic forms [41]). There is also an important difference in the elimination routes, since intact PINP is cleared by the liver (see Section II.D), whereas the small PINP Col 1-like peptides can also be filtered by the kidneys (Fig. 4) [1]. Because PICP and PINP are produced in equimolar amounts during the synthesis of type I procollagen, their molar concentrations in any relevant body fluids should, in principle, be almost identical. This is true, for example, in the fluid collected from a healing wound, where both the propeptides give similar estimates for the production rate of type I procollagen [22]. The ratio of PICP:PINP in adult blood is also approximately the expected 2–3:1, when the concentrations of both propeptides are expressed as protein concentrations (µg/L). However, there are both physiological and pathological situations in which this ratio differs from equimolarity. In the blood of infants and children (up until puberty), the molar concentration of PINP is higher than that of PICP, the ratio of PICP/PINP (in µg/L) being even clearly lower than 1:1 [42]. An even more dramatic discrepancy is found in patients with active Paget’s disease of bone

Chapter 23 Products of Bone Collagen Metabolism

[43, 44], or with aggressive breast cancer [45], or ovarian cancer [46]. In these cases, there can be an up to six-fold molar excess of PINP in the blood. The physiological basis for this finding is not yet known.

III. DEGRADATION PRODUCTS OF TYPE I COLLAGEN A. Formation, Cross-linking, and Aging of Collagen Fibers The rod-like collagen molecules possess a capacity for self-assembly and spontaneous formation of fibrils and fibers (see Chapter 2). The amino-terminal propeptide often remains transiently attached to some of the molecules and can be visualized on the surface of thin type I collagen fibers in tissues undergoing active collagen synthesis (see Section II.A) [13]. One characteristic location for such fibers is the epithelial–stromal junction below a metabolically active epithelium [47]. In the newly formed osteoid, some amino-terminal propeptides of type I procollagen are retained for the time of matrix maturation, since the collagenous matrix can be stained with anti-PINP antibodies, but no substantial amounts of the propeptide are any longer present in the mineralized matrix [T. Taube, I. Elomaa, L. Risteli, J. Risteli, unpublished]. The aminoterminal propeptide can be isolated as a 24-kDa phosphoprotein from the organic matrix of fetal bone, but not from the adult bone that has a much lower rate of synthesis of type I collagen [21]. In the case of type III collagen, a major collagen of soft tissues, pN collagen molecules seem to cover the surface of virtually all fibers, although they only represent a 5% fraction of the total amount of type III collagen [48]. The biological function of a fibrillar collagen is to provide the tissue with tensile strength. This is due to covalent bonds, the cross-links, that are formed between the individual collagen molecules in a collagen fiber. The cross-links result from a complicated series of partially alternative chemical reactions that gradually lead through divalent cross-links, joining two polypeptide chains, to multivalent (tri- or even tetravalent) cross-links (see Chapter 2). Continuing cross-linking explains the fact that older collagen in soft tissues is progressively more insoluble when studied in various ways, e.g., by digestion with pepsin or bacterial collagenase. The trivalent pyridinoline cross-links, which have been much studied, are abundant in the bone, but are even more so in the soft tissues where the maturation process is mostly complete and occurs within a relatively short time after the assembly of the collagen fiber. At least in some

397 tissues, the appearance of pyridinoline cross-links seems to be related to the development of an irreversible fibrosis, e.g., in the skin [49] and in the liver [50]. There is a wide variation in the cross-links, depending on whether the precursor lysyl residues have been hydroxylated to hydroxylysines or not [51]. Bone is the major connective tissue in which the collagen cross-links remain immature to a significant extent, the content of the divalent cross-links being about 2–4 times the concentration of the pyridinolines [52]. There are two obvious explanations for this finding: first, that mineralization slows down the cross-linking process and, second, that a significant part of the mature cross-links in bone have some other structure than pyridinoline [9]. The concentration of the divalent cross-links has been found to specifically decrease in osteoporosis [52, 53]. Unfortunately, analysis of the maturity of the cross-links in bones or other tissues suffers from the lack of easy, rapid, and reliable quantitative methods that would be suitable for a large number of small tissue samples. The development of immunoassays based either on synthetic peptides or on naturally cross-linked telopeptides may help in this respect, provided that the specificity of such assays is carefully tested. The application of such assays that detect the polypeptide chains involved in the cross-linked telopeptide structures has already revealed that, in addition to the pyridinolines and the relatively recently described pyrrolic cross-links [54, 55], there must still exist other, uncharacterized mature cross-links [9, 56]. In soft tissues, immaturely cross-linked collagens are transiently encountered during normal collagen turnover and in the granulation tissues. Defective cross-linking can also be a major feature of the stroma of malignant tumor tissue [48]. Interestingly, it has been found that lysyl oxidase, the only enzyme involved in the cross-link process, is identical to the tumor suppressor protein known as rrg [57] and that its synthesis is decreased in malignant breast tumor tissue [58]. Some further modifications take place during the aging of the collagen fibers, such as the cross-linking through nonenzymatically attached sugar residues [59] and the racemization and β-isomerization of some aspartic acid residues (see below). It is not known if these modifications proceed similarly in the soft tissues and in the skeleton. A β-isomerization of an aspartic acid residue as the result of a nonenzymatic, physiological aging reaction is known to take place both in the carboxy-terminal [56] and the amino-terminal telopeptide of the α1-chain of type I collagen [60]. This occurs via the formation of an unstable succinimide ring, which will spontaneously hydrolyze either back to a normal L-aspartyl or to an L-βaspartyl residue (Fig. 5). The factors determining which

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Figure 5

The β-isomerization and racemization reactions of aspartate residues in

proteins.

Asp residues are susceptible to such isomerization are the nature of the adjacent amino acid residues and the constraints imposed by the surrounding secondary and tertiary structures [61]. On the basis of specific immunoassays, the ratio of the α:β forms of aspartic acid in the carboxy-terminal telopeptide has been reported to be 0.40 in cortical and 0.48 in trabecular bone collagen and as high as 1.33 in pagetic bone [62]. Thus, even 70–80 % of the L-Asp in this location in normal adult bone collagen could be isomerized into the β-configuration. It would also be interesting to know if a similar development takes place in soft-tissue type I collagen and if the mineralization process in bone affects the rate of β-isomerization. Several cross-linked carboxy-terminal telopeptides of type I collagen have been isolated from urine that contains either only two α or two β forms or one of both forms [56]. In addition to the isomerization, a racemization of the α-carbon of the aspartate to give the corresponding D-amino acid derivative is also possible (Fig. 5). D-Asp seems to accumulate at a rate of about 0.1% per year during aging in the human bone fraction, containing mostly collagen [63]; thus, at the age of 60 years, about 6% of the total amount of the L-Asp residues in bone collagen have been racemized into D-Asp.

as a covalently cross-linked assembly of large numbers of molecules. In particular, the helical region of a collagen molecule can only be attacked by a few proteolytic enzymes before the cross-linking ends of the molecule have been cleaved. In soft tissues, there seem to be two routes of collagen breakdown, one being active in conditions associated with rapid remodeling of the tissue and the other under steady-state conditions [64]. In bones, the corresponding pathways are most probably due to the activities of matrix metalloproteinases and cathepsin K, respectively [65]. Because of the structural complexity and the presence of alternative degradation pathways, the pattern of crosslinked peptides that arises from the degradation of a certain collagen type in a certain tissue is difficult to predict. The existence of different enzymes with differing cleaving specificities and functioning in different metabolic situations adds to the complexity. Assessment of bone collagen degradation can be based either on determining small breakdown products, such as modified amino acids (including cross-links), or on immunological quantification of peptides.

C. Products Derived from the Amino-terminal Telopeptide of Type I Collagen B. Degradation Pathways The collagen in many soft tissues and particularly that in the skeleton is actively turned over throughout the life of an individual, although the overall turnover rate is naturally at its highest in infants and children. A number of enzymes can degrade soluble forms of type I collagen in the test tube. These enzymes include matrix metalloproteinases, cysteine proteinases, and serine proteinases [64]. However, it is more difficult to elucidate the real biological significance of such an activity because, in the tissues, the collagen is not present as soluble single molecules but

The amino-terminal telopeptide regions of both the α1(I) chain (17 amino acids) and the α2(I) chain (11 amino acids) of type I collagen contain lysine (or hydroxylysine) that can be oxidized to allysine (hydroxyallysine). The further events at this location can, in principle, lead to either intermolecular or intramolecular cross-links, resulting in a complex pattern of possible structures. One class of multivalent cross-links, those with pyridinoline structures, can be found both in blood and in urine. When quantified in such fluids, they do not reveal their specific origins, however, although the lysyl pyridinoline variant is more abundant

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in bone than in soft-tissue collagen. In addition to the Nterminal telopeptide of type I collagen, the cross-links can originate in the C-terminal telopeptide of the same protein or in other collagen types. Measurement of collagen cross-links is a more specific way of assessing the breakdown of fibrillar collagen than the traditional hydroxyproline measurement, however, because the cross-links can only be derived from the degradation of collagen molecules that have participated in collagen fibers. Furthermore, the pyridinoline cross-links are not absorbed from the diet, so their excretion is only related to their endogenous formation. The N-terminal telopeptide of type I collagen also contains other multivalent cross-link structures than pyridinolines. In particular, the pyrrole cross-links are predominantly present in the N-terminus [54, 55]. An immunological assay method has been described for the N-terminal telopeptide of type I collagen that detects a related antigen in blood [66] and in urine [67]. The antigen measured has been called NTx, and it represents a neoepitope, i.e. the corresponding sequence is not recognized by the antibody if the collagen molecule is intact. The structure of the Ntx peptide and its antigenic epitope are shown in Figure 6. NTx comprises one peptide derived from the α1(I) chain and another from the α2(I) chain, incorporating a single (hydroxy)lysine residue from the helical domain of a third chain. In urine samples, the apparent concentration of the NTx peptide has been found

Figure 6 Antigenic epitope of the NTx assay for the amino-terminal telopeptide of the human type I collagen molecule. The epitope depends on a specific proteolytic cleavage of the α2(I)N-telopeptide to the sequence shown where J is pyroglutamate and K is the original lysyl residue embodied in a cross-link. Reproduced from Apone et al. [90], with permission.

to increase when the samples are exposed to UV light [68]. Such a finding could be explained by the presence of pyrrolic cross-links in the small peptides; these cross-links are known to easily aggregate, and UV light could dissolve such aggregates. An aspartate residue that is located in the α2(I) part of NTx has been shown to undergo β-isomerization [60]. Although it is possible to set up assays that differentiate between the α- and β-isomers in this location, the specificity of the commonly used NTx assay in this regard has not been published.

D. Products Derived from the Helical Domain of Type I Collagen According to a traditional view of collagen breakdown, the major triple-helical domain of type I collagen contains the cleavage site for the mammalian collagenase enzyme that is unique in its ability to degrade this protein under physiological conditions. This enzyme is a member of the group of matrix metalloproteinases. Once the collagen molecule has been cut in two, the resulting shorter helices are no longer stable at body temperature and the polypeptide chains thus become susceptible to other proteindegrading enzymes as well. However, cathepsin K, which is the major collagen-degrading enzyme of osteoclasts, seems also to cleave the helical domain of native type I collagen in many locations in a reproducible manner [65]. Peptide antigens derived from the central, helical part of type I collagen have been reported to exist in blood [69] and in urine [70–72]. These findings indicate that some of the degradation products must be relatively long peptides, in order to contain an antigenic determinant (epitope). The circulating antigens have been detected by using antibodies raised to purified human collagen [69] and the urinary ones by using monoclonal antibodies to synthetic peptides [70–72]. The collagenous helix is generally considered an extremely weak immunogen both in its native and denatured state [73]. Thus it is essential that the true chemical natures of the physiological, immunoreactive substances are carefully characterized. The collagenous nature of the circulating antigen has been verified by its sensitivity to highly purified bacterial collagenase [69]. The sequence of the immunogen that has been used in developing the urinary assay has, on the other hand, been derived from a peptide isolated from the urine of a patient with Paget’s disease [70]. Despite this, it has not yet been verified beyond doubt that what is measured in the urine by such an assay under normal circumstances is indeed derived from bone tissue, particularly as there is a 90% cross-reaction with the corresponding part of type III

400 collagen [74]. The assay has been used for some clinical studies in osteoporosis [75]. 4-hydroxyproline is necessary for the correct triplehelical conformation of collagen at body temperature. This modified imino acid makes up about 12% of the weight of the type I collagen molecule and can thus be used as a measure of the collagen content of a tissue. The acid hydrolysis, followed by a chemical color reaction used to assess this imino acid, can be applied both for tissue samples and for urine. This method has traditionally been used as a measure of bone collagen metabolism [28]. Once set free from the helical part of a collagen molecule, hydroxyproline can no longer be incorporated into new protein; however, exogenous hydroxyproline is absorbed from the diet. Most of the imino acid (up to 90%) is metabolized in the liver, leading to the formation of pyruvate and glyoxalate, but some is always passed into the urine. Excretion of hydroxyproline correlates with growth velocity in children and with the presence of bone degrading processes, e.g., bone metastases of cancers, in adults [28]. Also, a generally enhanced bone metabolic rate increases urinary hydroxyproline excretion. For this reason, hydroxyproline served as an indirect test of thyroid hormone action before the time of specific thyroid hormone assays. Some hydroxyproline is directly derived from collagen synthesis, since the helical domain of PINP contains it. A substantial part of the urinary hydroxyproline can also be derived from the C1q component of the complement, particularly in inflammatory conditions [76]. The 4-hydroxyproline content is similar in all type I collagens. The extent of other post-translational modifications in the helical domain varies from one situation and tissue to another. Such modifications include the hydroxylation of lysine, the subsequent glycosylation of hydroxylysine – first by galactose and then by glucose – and the hydroxylation of proline at position 3, leading to the formation of 3-hydroxyproline. The presence of a halfway glycosylated hydroxylysine, galactosylhydroxylysine, has been suggested to be characteristic of adult bone tissue, and the excretion of this amino acid in the urine has been used as a marker of bone collagen degradation [77]. An immunological assay has also been introduced for galactosylhydroxylysine [78].

E. Products Derived from the Carboxy-terminal Telopeptide of Type I Collagen In the carboxy-terminal telopeptides of type I collagen, lysine/hydroxylysine is only present in the α1(I) chains, making the number of possible alternative cross-linked structures smaller than the corresponding number at the

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amino-terminus where both α1(I) and α2(I) contain lysine/ hydroxylysine. Since the divalent cross-link formed first binds one telopeptide to the helical region of another collagen molecule, there is a possibility that the telopeptide of the second α1(I) chain remains free. Thus, several degradation products can be formed from the carboxyterminal part of type I collagen that differ with respect to the α1-chain; this either has no cross-link or participates in a divalent or a trivalent cross-link. The latter may be hydroxylysyl pyridinoline, lysyl pyridinoline, or even pyrrole. There is some evidence also for the presence of an as yet uncharacterized cross-link [9, 56]. When pyridinoline cross-links as such are being measured, they can be derived both from the carboxy-terminus and from the amino-terminus of the type I collagen molecule. More specifically, several immunoassays have been developed for structures involving the carboxy-terminal telopeptides and they can give different results, depending on their immunochemical specificity with respect to the length of the antigen, the maturity of the cross-link and the possible modifications of an aspartic acid residue in the telopeptide. The ICTP antigen is a trivalently cross-linked structure that was originally isolated from human femoral bone after digestion with trypsin or bacterial collagenase and shown to contain the carboxy-terminal telopeptides of two α1(I) chains and material from the helical part of a third chain [79]. The immunological assay for ICTP detects relatively large degradation products of mature type I collagen since the epitope requires the presence of two phenylalaninerich regions [80] (Fig. 7). Such antigens are detectable in the blood and in several other physiological fluids, but not in the urine. Specificity tests have indicated that cathepsin K cleaves the carboxy-terminal telopeptide of the α1-chain of type I collagen on both sides of the sequence LPQPPQE, which is located between the phenylalanine-rich region and the cross-link site; thus, the degradation product liberated by cathepsin K is not detected by the ICTP assay (Fig. 7). A series of assays called CrossLaps or CTx have been developed using synthetic peptides related to the carboxyterminal telopeptide as immunogen. The respective part of the telopeptide is situated more in C-terminal direction than the ICTP epitope (Fig. 7). The first of these could recognize related antigens in the urine; the immunogen contained a stretch of eight amino acids, including the cross-link site of the α1(I) chain as an unmodified lysine [81]. Later, variants of the assay were introduced that can differentiate between the α- and β-forms of the Asp located in the epitope [82]. The CrossLaps α- or β-assays may detect both uncross-linked and di- or trivalently crosslinked species since the epitope is a linear sequence of only

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Figure 7

Antigenic epitopes in the carboxy-terminal telopeptide of human type I collagen (ICTP and CrossLaps). The ICTP assay demands two adjacent streches that both contain a phenylalanine-rich domain FDFSF. The CrossLaps assays detect linear, eight amino acidlong sequences (bold) where the aspartic acid residue is either in an α- or a β-configuration. Cat K = cathepsin K cleavage site. Reproduced from Garnero et al. [92], with permission.

six amino acids (Fig. 7). However, the serum CrossLaps β-β-assay in which a double antibody technique is used [83] does not detect immature degradation products (Fig. 7). In addition to α to β isomerization, the aspartic acid residue in the CrossLaps epitope can undergo L to D racemization. All the four alternative structures (αL, αD, βL, and βD) have been found in the carboxy-terminal telopeptide of type I collagen in human tissues [84]. A number of clinical studies have indicated that the ICTP and CrossLaps methods reflect different processes in type I collagen metabolism, i.e. pathological states involving breakdown of bone tissue and normal turnover of the skeleton, respectively.

pathological situations, and thus to liberate the ICTP antigen into the blood. Such situations include tumor metastases, myeloma lesions, and rheumatoid arthritis (Fig. 8) [87]. In the hereditary disease pycnodysostosis, there is a genetic defect in cathepsin K, with lower than normal NTx and CrossLaps excretions and a compensatorily highly increased circulating ICTP concentration [88]. Degradation products of bone collagen have been immunochemically identified both in osteoclast cultures on bone slices [89–92] and in vitro, by degrading demineralized bone matrix with proteolytic enzymes. In the former system, peptides are liberated that react in assays for the amino-terminal [89] and the carboxy-terminal [89, 91, 92] telopeptides of type I collagen, although the exact sizes of these degradation products have not been reported.

F. The Physiological Degradation Products of Type I Collagen It first became evident from clinical studies that cathepsin K is active in the major pathway of normal bone turnover, a process that is enhanced in estrogen deficiency, for instance. The degradation phase of the bone metabolic cycle involves the specific attachment of osteoclasts to the mineralized surface, the dissolution of the inorganic mineral component (acidic microenvironment) and the degradation of matrix proteins (protease activity). Bone collagen degradation occurs mainly in the resorption lacuna and is analogous to the intracellular route of soft-tissue collagen degradation [85]. Cathepsin K, with its ability to cut the collagen molecule in both the telopeptides and in the helical domain [65, 86], liberates, among other things, NTx and CrossLaps (CTx) antigens from the bone. Another class of proteolytic enzymes, the matrix metalloproteinases, seems to be more actively involved in the localized breakdown of bone that takes place in

Figure 8 Size distribution of the ICTP and CrossLaps antigens in human serum. A 2.5-ml sample of serum from a 55-year-old woman with rheumatoid arthritis was chromatographed on a Sephacryl S-100 column at room temperature in phosphate-buffered saline containing 0.04% Tween20. The dotted line indicates the detection limit of the CrossLaps assay based on the absorbance of the blank (standard A = 0 pmol/l) of the kit. Before analyses the 20-min fractions were concentrated by lyophilization. Reproduced from Sassi et al. [87], with permission.

402 Interestingly, no free hydroxylysyl pyridinoline or lysyl pyridinoline is set free from the bone [90]. The probable liberation of peptide products originating from the helical parts of the collagen molecules or hydroxyproline and galactosylhydroxylysine have yet to be characterized in this system. When demineralized bone matrix is treated with proteolytic enzymes in the test tube, the kinetics of the appearance of the different degradation products can be followed more closely. Cathepsin K liberates an NTx antigen and generates a neoepitope that is necessary for reactivity in this immunoassay [86]. The enzyme can degrade up to 100% of a purified, decalcified bone matrix [65], cutting the collagen molecules in telopeptide areas as well as in the helical domain. The number of studies on the exact mechanism of bone collagen degradation is still quite limited. However, studies on the urinary excretion of presumed degradation products of bone collagen abound (see Chapter 35). Both hydroxyproline and the pyridinoline cross-links are present partially in free form and partially in peptide-bound form in the urine; when the turnover rate of bone increases, there is typically a concomitant increase in the total excretion of the cross-links, but also a disproportionate increase in their peptide-bound fraction. The urinary antigens that are recognized either by the NTx or the CrossLaps assay behave in a manner resembling the peptide-bound crosslinks in this respect. Furthermore, different antiresorptive pharmaceuticals seem to affect the free and peptidebound fractions differently. For example, estrogen therapy

JUHA RISTELI AND LEILA RISTELI

decreases the urinary excretion of both the free and peptide-bound cross-links in postmenopausal women, whereas bisphosphonate treatment of patients with Paget’s disease has a marked effect only on the cross-linked peptides without a change in the free cross-links [93]. It can be speculated that the bisphosphonates may be specific for osteoclasts, which do not seem to produce free pyridinolines, whereas estrogen has a more general effect on both bone and nonmineralized tissues. This reasoning suggests that the free pyridinolines in the urine would largely be derived from the degradation of soft-tissue collagens. While it is known that type III collagen contains a lot of hydroxylysyl pyridinoline, this most probably is not the whole explanation for the interesting finding, in particular because there is no significant difference between hydroxylysyl and lysyl pyridinolines in this respect. The degradation of extraskeletal type I collagen and the further metabolism of the degradation products of bone collagen are not known in detail (Fig. 9). The majority of the hydroxyproline set free from collagens is known to be metabolized in the liver. The endothelial cells of the liver possess specific gelatin receptors for the internalization of large collagenous peptides [25]. If such peptides still contain the cross-links, it is possible that the formation of free pyridinolines could partially take place in the liver. Another site for the further handling of the cross-linked peptides is the kidneys. In particular, the tubuli contain several exo- and endoproteinases, which digest small peptides to amino acids that can be reabsorbed. Most probably also the original degradation products of type I collagen are further shortened in the kidneys, for example the main urinary degradation product detected by the CrossLaps assay only contains the core sequence EKAHDGGR [56]. Also autoradiographic methods have demonstrated that the kidneys participate in the degradation of collagenous peptides [94]. The renal clearance of the free pyridinoline cross-links has been shown to be four-fold higher than that of the cross-links still present in peptides [95]. Since the fractional clearance of the former is greater than one and that of the latter less than one, some free pyridinolines excreted in urine are produced by the kidney (Fig. 9). The possible effects of pathological processes affecting the kidneys on cross-link excretion – or even on the excretion of any type I collagen telopeptide antigens – have not been studied.

IV. CLOSING REMARKS Figure 9

Schematic presentation of the distribution between various pools and of the clearances of type I collagen telopeptides.

It has been a challenge for research to assess the balance between bone collagen synthesis and degradation by the simultaneous measurement of metabolic products related

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to these two processes. There are two possible analytes related to the synthesis of type I collagen, the carboxyterminal (PICP) and the amino-terminal (PINP) propeptides of type I procollagen, and the respective assay methods are already quite well established. There is a great deal of clinical experience on their performance in different physiological and pathological situations. Although in general both propeptides give relevant information of type I collagen synthesis, in some situations the assay for the intact PINP seems to be superior to that for PICP. The general picture of the degradation of bone type I collagen has been cleared by recent research, with the clarification of the roles of the cathepsin K and matrix metalloproteinase pathways. However, a number of more specific questions remain, related to the further metabolic fates of the primary degradation products, to the breakdown products of the helical domain or to the origins of the free forms of cross-links, for instance. Much clinical experience has been gained using urine samples, but the present trend is towards serum assays. The respective contributions of the skeleton and the soft tissues to the different degradation products have so far not been seriously studied.

10.

11.

12.

13. 14.

15.

16.

17.

18.

19.

References 20. 1. Risteli, J., and Risteli, L. (1995). Analysing connective tissue metabolites in human serum. Biochemical, physiological and methodological aspects. J. Hepatol. (Suppl. 22), 77–81. 2. Risteli, J., and Risteli, L. (1997). Assays of type I procollagen domains and collagen fragments: problems to be solved and future trends. Scand. J. Clin. Lab. Invest. (Suppl. 227), 107–113. 3. Risteli, L., and Risteli, J. (2002). Extracellular matrix metabolites in body fluids. In “Connective Tissue and its Heritable Disorders: Molecular, Genetic and Medical Aspects” (P. M. Royce, and B. Steinmann, eds.), 2nd edition, pp. 1141–1160. John Wiley & Sons, New York. 4. Scariano, J. K., Walter, E. A., Glew, R. H., Hollis, B. W., Henry, A., Ocheke, I., and Isicei C. O. (1995). Serum levels of pyridinoline crosslinked carboxyterminal telopeptide of type I collagen (ICTP) and osteocalcin in rachitic children in Nigeria. Clin. Biochem. 28, 541–545. 5. Oginni, L. M., Sharp, C. A., Badru, O. S., Risteli, J., Davie M. W. J., Worsfold, M. (2003). Radiological and biochemical resolution of nutritional rickets with calcium. Arch. Dis. Child. 88, 812–817. 6. Leskelä, H.-V., Risteli, J., Niskanen, S., Koivunen, J., Ivaska, K. K., and Lehenkari, P. (2003). Osteoblast recruitment from stem cells does not decrease by age at late childhood. Biochem. Biophys. Res. Commun. 311, 1008–1013. 7. Jukkola, A., Risteli, L., Melkko, J., and Risteli, J. (1993). Procollagen synthesis and extracellular matrix deposition in MG-63 osteosarcoma cellls. J. Bone Miner. Res. 8, 651–657. 8. Mechanic, G. L., Katz, E. P., Henmi, M., Noyes, C. and Yamauchi, M. (1987). Locus of a histidine-based, stable trifunctional, helix to helix collagen cross-link: stereospecific collagen structure of type I skin fibrils. Biochemistry 26, 3500–3509. 9. Eriksen, H., Sharp, C. A., Robins, S. P., Sassi, M.-L., Risteli, L., Risteli, J. (2004). Differently cross-linked and uncross-linked carboxy-terminal

21.

22.

23.

24.

25.

26.

27.

28. 29.

telopeptides of type I collagen in human mineralised bone. Bone 34, 720–727. Prockop, D. J., Kivirikko, K. I., Tuderman, L., and Guzman, N. A. (1979). The biosynthesis of collagen and its disorders. N. Engl. J. Med. 301, 13–23, 77–85. Kessler, E., Takahara, K., Biniaminov, L., Brusel, M., and Greenspan, D. S. (1996). Bone morphogenetic protein-1: the type I procollagen C-proteinase. Science 271, 360–362. Bornstein, P. (2002). The NH2-terminal propeptides of fibrillar collagens: highly conserved domains with poorly understood functions. Matrix Biol. 21, 217–226. Fleischmajer, R., Perlish, J. S., and Timpl, R. (1985). Collagen fibrillogenesis in human skin. Ann. N. Y. Acad. Sci. 460, 246–257. Byers, P. H., and Cola, W. G. (2002). Osteogenesis imperfecta. In “Connective Tissue and its Heritable Disorders” (P. M. Royce, and B. Steinmann, eds.), 2nd edition, pp. 385–430. Wiley-Liss, New York. Bailey, A. J., Sims, T. J., and Knott, L. (2002). Phenotypic expression of osteoblast collagen in osteoarthritic bone: production of type I homotrimer. Int. J. Biochem. Cell Biol. 34, 176–182. Mann, V., Hobson, E. E., Stewart, T. L., Grant, S. F., Robins, S. P., Aspden, R. M., and Ralston, S. H. (2001). A COL1A1 Sp1 binding site polymorphism predisposes to osteoporotic fracture by affecting bone density and quality. J. Clin. Invest. 107, 899–907. Clark, C. C. (1979). The distributional initial characterization of oligosaccharide units on the COOH-terminal propeptide extensions of the pro-α1 and pro-α2 chains of type I procollagen. J. Biol. Chem. 254, 10798–10802. Melkko, J., Niemi, S., Risteli, L., and Risteli, J. (1990). Radioimmunoassay of the carboxyterminal propeptide of human type I procollagen. Clin. Chem. 36, 1328–1332. Pedersen, B. J., and Bonde, M. (1994). Purification of human procollagen type I carboxyl-terminal propeptide cleaved as in vivo from procollagen and used to calibrate a radioimmunoassay of the propeptide. Clin. Chem. 40, 811–816. Taubman, M. B., Goldberg, B., and Sherr, C. J. (1974). Radioimmunoassay for human procollagen. Science 186, 1115–1117. Fischer, L. W., Robey, P. G., Tuross, N., Otsuka, A. S. O., Tepen, D. A., Esch, F. S., Shimasaki, S., and Termine, J. D. (1987). The Mr 24 000 phosphoprotein from developing bone is the NH2-terminal propeptide of the α1 chain of type I collagen. J. Biol. Chem. 37, 13457–13463. Melkko, J., Kauppila, S., Risteli, L., Niemi, S., Haukipuro, K., Jukkola, A., and Risteli, J. (1996). Immunoassay for intact aminoterminal propeptide of human type I procollagen. Clin. Chem. 42, 947–954. Kauppila, S., Jukkola, A., Melkko, J., Risteli, L., TurpeenniemiHujanen, T., Vuorinen, K., and Risteli, J. (2001). Aminoterminal propeptide of α1-homotrimer variant of human type I collagen (hotPINP) in malignant pleural effusion. Anticancer Res. 21, 2293–2296. Lee, S. J., Evers, S., Roeder, D., Parlow, A. F., Risteli, J., Risteli, L., Lee, Y. C., Feizi, T., Langen, H., and Nussenzweig, M. C. (2002). Mannose receptor-mediated regulation of serum glycoprotein homeostasis. Science 295, 1898–1901. Smedsrød, B., Pertoft, H., Gustafsson, S., and Laurent, T. C. (1990). Scavenger functions of the liver endothelial cells. Biochem. J. 266, 313–327. Smedsrød, B., Melkko, J., Risteli, L., and Risteli, J. (1990). Circulating C-terminal propeptide of type I procollagen is cleared mainly via the mannose receptor in liver endothelial cells. Biochem. J. 271, 345–350. Melkko, J., Hellevik, T., Risteli, L., Risteli, J., and Smedsrød, B. (1994). Clearance of NH2-terminal propeptides of types I and III procollagen is a physiological function of the scavenger receptor in liver endothelial cells. J. Exp. Med. 179, 405–412. Kivirikko, K. I. (1970). Urinary excretion of hydroxyproline in health and disease. Int. Rev. Conn. Tissue Res. 5, 93–163. Lamandé, S. R., and Bateman, J. F. (1995). The type I collagen proα1(I) COOH-terminal propeptide N-linked oligosaccharide. Functional analysis by site-directed mutagenesis. J. Biol. Chem. 270, 17858–17865.

404 30. Toivonen, J., Tähtelä, R., Laitinen, K., Risteli, J., and Välimäki, M. J. (1998). Markers of bone turnover in patients with differentiated thyroid cancer on and off thyroxine suppressive therapy. Eur. J. Endocrinol. 138, 667–673. 31. Sorva, A., Tähtelä, R., Risteli, J., Risteli, L., Laitinen, K., JuntunenBackman, K., and Sorva, R. (1994). Familial high serum concentrations of the carboxyl-terminal propeptide of type I procollagen. Clin. Chem. 40, 1591–1593. 32. Puistola, U., Risteli, L., Kauppila, A., Knip, M., and Risteli, J. (1993). Markers of type I and type III collagen synthesis in serum as indicators of tissue growth during pregnancy. J. Clin. Endocrinol. Metab. 77, 178–182. 33. Sorva, A., Sorva, R., Risteli, J., Risteli, L., and Autio, P. (2001). Familial high serum PICP: high even in skin interstitial fluid, suppressible by topical corticosteroid treatment. Clin. Chim. Acta 314, 153–157. 34. Haukipuro, K., Melkko, J., Risteli, L., Kairaluoma, M. I., and Risteli, J. (1991). Synthesis of type I collagen in healing wounds in humans. Ann. Surg. 213, 75–80. 35. Oikarinen, A., Autio, P., Kiistala, U., Risteli, L., and Risteli, J. (1992). A new method to measure type I and III collagen synthesis in human skin in vivo: demonstration of decreased collagen synthesis after topical glucocorticoid treatment. J. Invest. Dermatol. 98, 220–225. 36. Jensen, L. T., Olesen, H. P., Risteli, J., and Lorenzen, I. (1990). External thoracic duct-venous shunt in conscious pigs for long term studies of connective tissue metabolism in lymph. Lab. Anim. Sci. 40, 620–624. 37. Eriksen, E. F., Charles, P., Melsen, F., Mosekilde, L., Risteli, L., and Risteli, J. (1993). Serum markers of type I collagen formation and degradation in metabolic bone disease: correlation to bone histomorphometry. J. Bone Miner. Res. 8, 127–132. 38. Charles, P., Mosekilde, L., Risteli, L., Risteli, J., and Eriksen, E. F. (1994). Assessment of bone remodelling using biochemical indicators of type I collagen synthesis and degradation: relation to calcium kinetics. Bone Miner. 24, 81–94. 39. Price, K. M., Silman, R., Armstrong, P., and Grudzinskas, J. G. (1994). Development of a radioimmunoassay for fetal antigen 2. Clin. Chim. Acta 224, 95–102. 40. Ørum, O., Hansen, M., Jensen, C. H., Sørensen, H. A., Jensen, L. B., Hørslev-Petersen, K., and Teisner, B. (1996). Procollagen type I N-terminal propeptide (PINP) as an indicator of type I collagen metabolism: ELISA development, reference interval, and hypovitaminosis D induced hyperparathyroidism. Bone 19, 157–163. 41. Suvanto-Luukkonen, E., Risteli, L., Sundström, H., Penttinen, J., Kauppila, A., and Risteli, J. (1997). Comparison of three serum assays for bone collagen formation during postmenopausal estrogen-progestin therapy. Clin. Chim. Acta 266, 105–116. 42. Tähtelä, R., Turpeinen, M., Sorva, R., and Karonen, S.-L. (1997). The amino-terminal propeptide of type I procollagen: Evaluation of a commercial radioimmunoassay kit and values in healthy subjects. Clin. Biochem. 30, 35–40. 43. Sharp, C. A., Davie, M. W. J., Worsfold, M., Risteli, L., and Risteli, J. (1996). Discrepant blood concentrations of type I procollagen propeptides in active Paget disease of bone. Clin. Chem. 42, 1121–1122. 44. Sharp, C. A., Risteli, J., Risteli, L., Davie, M. W. J. (1997). Type I procollagen propeptide ratio (PICP/PINP) responds rapidly to pamidronate in Paget´s disease of bone. J. Bone Min. Res. 12(Suppl 1), S269. 45. Jukkola, A., Tähtelä, R., Thölix, E., Vuorinen, K., Blanco, G., Risteli, L., and Risteli, J. (1997). Aggressive breast cancer leads to discrepant serum levels of the type I procollagen propeptides PINP and PICP. Cancer Res. 57, 5517–5520. 46. Simojoki, M., Santala, M., Risteli, J., and Kauppila, A. (2003). Discrepant expression of carboxy- and aminoterminal propeptides of type I procollagen predicts poor clinical outcome in epithelial ovarian cancer. Gynecol. Oncol. 88, 358–362.

JUHA RISTELI AND LEILA RISTELI 47. Rasmussen, H. B., Teisner, B., Andersen, J. A., Brandrup, F., Purkis, T., and Leigh, I. (1991). Immunohistochemical studies on the localization of fetal antigen 2 (FA 2), laminin and type IV collagen in basal cell carcinoma. J. Cutaneous Pathol. 18, 215–219. 48. Kauppila, S., Bode, M., Stenbäck. F., Risteli, L., and Risteli, J. (1999). Cross-linked telopeptides of type I and III collagens in malignant ovarian tumors in vivo. Brit. J. Cancer 81, 654–661. 49. Ricard-Blum, S., Esterre, P., and Grimaud, J. A. (1993). Collagen cross-linking by pyridinoline occurs in non-reversible skin fibrosis. Cell. Mol. Biol. 39, 723–727. 50. Ricard-Blum, S., Bresson-Hadni, S., Guerret, S., Grenard, P., Volle, P.-J., Risteli, L., and Grimaud, J. A. (1996). Mechanism of collagen network stabilization in human irreversible granulomatous liver fibrosis. Gastroenterology 111, 172–182. 51. Pornprasertsuk, S., Duarte, W. R., Mochida, Y., and Yamauchi, M. (2004). Lysyl hydroxylase-2b directs collagen cross-linking pathways in MC3T3-E1 cells. J. Bone Miner. Res. 19, 1349–1355. 52. Bailey, A. J., Wotton, S. F., Sims, T. J., and Thompson, P. W. (1993). Biochemical changes in the collagen of human osteoporotic bone matrix. Connect. Tissue Res. 29, 119–132. 53. Oxlund, H., Mosekilde, L., and Øxtoft, G. (1996). Reduced concentration of collagen reducible cross links in human trabecular bone with respect to age and and osteoporosis. Bone 19, 479–484. 54. Hanson, D. A., and Eyre, D. R. (1996). Molecular site specificity of pyridinoline and pyrrole cross-linking in type I collagen in human bone. J. Biol. Chem. 271, 26508–26516. 55. Brady, J. D., and Robins, S. P. (2001). Structural characterization of pyrrolic cross-links in collagen using a biotinylated Ehrlich’s reagent. J. Biol. Chem. 276, 18812–18818. 56. Fledelius, C., Johnsen, A. H., Cloos, P. A. C., Bonde, M., and Qvist, P. (1997). Characterization of urinary degradation products derived from type I collagen. Identification of a β-isomerized Asp-Gly sequence within the C-terminal telopeptide (α1) region. J. Biol. Chem. 272, 9755–9763. 57. Kenyon, K., Contente, S., Trackman, P. C., Tang, J., Kagan, H. M., and Friedman, R. M. (1991). Lysyl oxidase and rrg messenger RNA? Science 253, 802. 58. Peyrol, S., Raccurt, M., Gerard, F., Gleyzal, C., Grimaud, J. A., and Sommer, P. (1997). Lysyl oxidase gene expression in the stromal reactions to in situ and invasive ductal breast carcinoma. Am. J. Pathol. 150, 497–507. 59. Sell, D. R., and Monnier, V. M. (1989). Structure elucidation of a senescence cross-link from human extracellular matrix. J. Biol. Chem. 264, 21597–21602. 60. Brady, J. D., Ju, J., and Robins, S. P. (1999). Isoaspartyl bond formation within N-terminal sequences of collagen type I: implications for their use as markers of collagen degradation. Clin. Science 96, 209–215. 61. Lowenson, J., and Clarke, S. (1988). Does the chemical instability of aspartyl and asparaginyl residues in proteins contribute to erythrocyte aging. Blood Cells 14, 103–117. 62. Garnero, P., Fledelius, C., Gineyts, E., Serre, C.-M., Vignot, E., and Delmas, P. D. (1997). Decreased β-isomerization of the C-terminal telopeptide of type I collagen α1 chain in Paget’s disease of bone. J. Bone Miner. Res. 12, 1407–1415. 63. Helfman, P. M., and Bada, J. L. (1976). Aspartic acid racemisation in dentine as a measure of ageing. Nature 262, 279–281. 64. Everts, V., van der Zee, E., Creemers, L., and Beertsen, W. (1996). Phagocytosis and intracellular digestion of collagen, its role in turnover and remodelling. Histochem. J. 28, 229–245. 65. Garnero, P., Borel, O., Byrjalsen, I., Ferreras, M., Drake, F. H., McQueney, M. S., Foged, N. T., Delmas, P. D., and Delaissé, J.-M. (1998). The collagenolytic activity of cathepsin K is unique among mammalian proteinases. J. Biol. Chem. 273, 32347–32352. 66. Clemens, J. D., Herrick, M. V., Singer, F. R., and Eyre, D. R. (1997). Evidence that serum NTx (collagen-type I N-telopeptides) can act as an immunochemical marker of bone resorption. Clin. Chem. 43, 2058–2063.

405

Chapter 23 Products of Bone Collagen Metabolism 67. Hanson, D. A., Weis, M. A. E., Bollen, A.-M., Maslan, S. L., Singer, F. R., and Eyre, D. R. (1992). A specific immunoassay for monitoring human bone resorption: quantitation of type I collagen cross-linked N-telopeptides in urine. J. Bone Miner. Res. 7, 1251–1258. 68. Blumsohn, A. C., Naylor, K., and Eastell, R. (1995). Effect of light and γ-irridiation on pyridinolines and telopeptide of type I collagen in urine. Clin. Chem. 41, 1195–1197. 69. Hartmann, D., Trinchet, J. C., Ricard-Blum, S., Beaugrand, M., Callard, P., and Ville, G. (1990). Radioimmunoassay of type I collagen that mainly detects degradation products in serum: application to patients with liver diseases. Clin. Chem. 36, 421–426. 70. Ju, J., Leigh, S., Byrne, M., van der Rest, M., Liu, V., and Krane, S. (1997). A new marker for measuring bone resorption using an ELISA for a 14 amino acid peptide (residues 620–633) derived from the collagen α1(I) chain. J. Bone Miner. Res.12 (Suppl), S178. 71. Ju, H.-S. J., Maskrey, J., Krane, S., Riggs, B. L., and Allen, G. (1998). An immunoassay for measuring helical peptide/ α1(I) 620-633 derived from type I collagen α1 chain. Bone 23 (Suppl), S629. 72. Garnero, P., and Delmas, P. D. (2003). An immunoassay for type I collagen α1 helicoidal peptide 620–633, a new marker of bone resorption in osteoporosis. Bone 32, 20–26. 73. Furthmayr, H. (1982). Immunization procedures, isolation by affinity chromatography, and serological and immunochemical characterization of collagen specific antibodies. In “Immunochemistry of the Extracellular Matrix Volume I Methods” (H. Furthmayr, ed.), pp. 143–178. CRC Press, Boca Raton, FL. 74. Kit insert (2001). Metra Helical Peptide EIA kit. Quidel, 0760A (5/01) Catalog number 8022, p. 1–3 (www.quidel.com, read August 17, 2004). 75. Taxel, P., Fall, P. M., Prestwood, K. M., Dulipsingh, L., Ohannessian, C., and Raisz, L. G. (2004). Changes in urinary excretion of helical peptide during therapy for osteoporosis in older adults. Clin. Chem. 50, 747–750. 76. Robins, S. P. (1982). Turnover and cross-linking of collagen. In “Collagen in Health and Disease” (J. B. Weiss, and M. I. V. Jayson, eds.), pp. 160–178. Churchill Livingstone, Edinburgh. 77. Bettica, P., Taylor, A. K., Talbot, J., Moro, L., Talamini, R., and Baylink, D. J. (1996). Clinical performances of galactosyl hydroxylysine, pyridinoline, and deoxypyridinoline in postmenopausal osteoporosis. J. Clin. Endocrinol. Metab. 81, 542–546. 78. Leigh, S. D., Ju, H. S., Lundgard, R., Daniloff, G. Y., and Liu, V. (1998). Development of an immunoassay for urinary galactosylhydroxylysine. J. Immunol. Methods 220, 169–178. 79. Risteli, J., Elomaa, I., Niemi, S., Novamo, A., and Risteli, L. (1993). Radioimmunoassay for the pyridinoline cross-linked carboxyterminal telopeptide of type I collagen: a new serum marker of bone collagen degradation. Clin. Chem. 39, 635–640. 80. Sassi, M.-L., Eriksen, H., Risteli, L., Niemi, S., Mansell, J., Gowen, M., and Risteli, J. (2000). Immunochemical characterization of assay for carboxyterminal telopeptide of human type I collagen: Loss of antigenicity by treatment with cathepsin K. Bone 26, 367–373. 81. Bonde, M., Qvist, P., Fledelius, C., Riis, B. J., and Christiansen, C. (1994). Immunoassay for quantifying type I collagen degradation products in urine. Clin. Chem. 40, 2022–2025. 82. Bonde, M., Garnero, P., Fledelius, C., Qvist, P., Delmas, P. D., and Christiansen, C. (1997). Measurement of bone degradation products in

83.

84.

85. 86.

87.

88.

89.

90.

91.

92.

93.

94.

95.

serum using antibodies reactive with an isomerized form of an 8 amino acid sequence of the C-telopeptide of type I collagen. J. Bone Miner. Res. 12, 1028–1034. Rosenqvist, C., Fledelius, C., Christgau, S., Pedersen, B. J., Bonde, M., Qvist, P., and Christiansen, C. (1998). Serum CrossLaps One Step ELISA. First application of monoclonal antibodies for measurement in serum of bone-related degradation products of type I collagen. Clin. Chem. 44, 2281–2289. Gineyts, E., Cloos, P. A. C., Borel, O., Grimaud, L., Delmas, P. D., and Garnero, P. (2000). Racemization and isomerization of type I collagen C-telopeptides in human bone and soft tissues: assessment of tissue turnover. Biochem. J. 345, 481–485. Väänänen, H. K., Zhao, H., Mulari, M., and Halleen, J. M. (2000). The cell biology of osteoclast function. J. Cell Sci. 113, 377–381. Atley, L. M., Mort, J. S., Lalumiere, M., and Eyre, D. R. (2000). Proteolysis of human bone collagen by cathepsin K: Characterization of the cleavage sites generating the cross-linked N-telopeptide neoepitope. Bone 26, 241–247. Sassi, M.-L., Åman, S., Hakala, M., Luukkainen, R., and Risteli, J. (2003). Assay for cross-linked carboxyterminal telopeptide of type I collagen (ICTP) unlike CrossLaps assay reflects increased pathological degradation of type I collagen in rheumatoid arthritis. Clin. Chem. Lab. Med. 41, 1038–1044. Nishi, Y., Atley, L., Wyre, D. E., Edelson, J. G., Superti-Furga, A., Yasuda, T., Desnick, R. J., and Gelb, B. D. (1999). Determination of bone markers in pycnodysostosis: effects of cathepsin K deficiency on bone matrix degradation. J. Bone Min. Res. 14, 1902–1908. Foged, N. T., Delaissé, J.-M., Hou, P., Lou, H., Sato, T., Winding, B., and Bonde, M. (1996). Quantification of the collagenolytic activity of isolated osteoclasts by enzyme-linked immunosorbent assay. J. Bone Miner. Res. 11, 226–237. Apone, S., Lee, M. Y., and Eyre, D. R. (1997). Osteoclasts generate cross-linked collagen N-telopeptides (Ntx) but not free pyridinolines when cultured on human bone. Bone 21, 129–136. Parikka, V., Lehenkari, P., Sassi, M.-L., Halleen, J., Risteli, J., Härkönen, P., and Väänänen, H. K. (2001). Estrogen reduces the depth of resorption pits by disturbing the organic bone matrix degradation activity of mature osteoclasts. Endocrinology 142, 5371–5378. Garnero, P., Ferreras, M., Karsdal, K. A., Nicamhlaoibh, R., Risteli, J., Borel, O., Qvist, P., Delmas, P. D., Foged, N. T., and Delaissé, J. M. (2003). The type I collagen fragments ICTP and CTX reveal distinct enzymatic pathways of bone collagen degradation. J. Bone. Miner. Res. 18, 859–867. Garnero, P., Gineyts, E., Arbault, P., Christiansen, C., and Delmas, P. D. (1995). Different effects of bisphosphonate and estrogen therapy on free and peptide-bound bone cross-links excretion. J. Bone Miner. Res. 10, 641–649. Ruckidge, G. J., Riddoch, G. I., Williams, L. M., and Robins, S. P. (1988). Autoradiographic studies of the renal clearance of circulating Type I collagen fragments in the rat. Collagen Rel. Res. 8, 339–348. Colwell, A., and Eastell, R. (1996). The renal clearance of free and conjugated pyridinium cross-links of collagen. J. Bone Miner. Res. 11, 1976–1980.

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Chapter 24

Supramolecular Structure of Cartilage Matrix Peter Bruckner

Department of Physiological Chemistry and Pathobiochemistry, University of Münster, Münster, Germany

IX. Studies of Transgenic Mice and of Human Genetic Matrix Diseases X. Correlating Structure with the Biomechanical Role of Articular Cartilage XI. Models of Cartilage Fibril Structure XII. Future Perspectives References

I. Summary II. Introduction III. Light and Electron Micrography IV. Biochemistry of Cartilage V. Studies of Fibril Structures by X-ray Diffraction VI. Structure of Fibril Fragments Obtained by Mechanical Disruption of Tissue VII. Studies of Collagen Cross-linking in Cartilage Fibrils VIII. Reconstitution of Aggregates from Soluble Collagens and other Macromolecules

I. SUMMARY

which is rich in polyanionic carbohydrates and, thus, water. Implications of this concept are discussed for connective tissues other than cartilage and for extracellular matrices in general.

The extracellular matrix of hyaline cartilage comprises the great majority of the volume of this tissue and is responsible for its properties. The matrix consists of two basic suprastructural components: fibrils with a D = 67 nm periodic banding pattern and an electron-lucent extrafibrillar matrix with no conspicuous features. The structural properties of these matrix constituents and their functional implications are discussed. The heterotypic fibrils contain not only several types of collagen but also noncollagenous macromolecules. These molecular components specifically self-assemble, either into distinct domains within the fibrils or, more attractively, into aggregates with characteristics of metal alloys in many respects. Thus, specific properties of the fibrils are indirectly determined by the mass proportions of their molecular components. Similar considerations apply to the extrafibrillar matrix Dynamics of Bone and Cartilage Metabolism

II. INTRODUCTION Three types of cartilage have been distinguished on the basis of histological criteria and biomechanical properties: hyaline, elastic, and fibrous cartilages. The most prevalent type is hyaline cartilage, which is a visually uniform, translucent tissue found in the skeleton of all vertebrates. Articular cartilage, the most familiar hyaline cartilage, forms the smooth gliding surfaces of joints, such as the knee and hip, that permit locomotion in animals. Injuries to this tissue and common diseases such as osteoarthritis, impair joint mobility and constitute a major 407

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focus of modern medicine. Hyaline cartilage also comprises growth plate, the transient template required for endochondral bone formation in fetal development, skeletal growth, and repair. In addition, hyaline cartilage occurs as a permanent structural tissue in costal cartilages and tracheal reinforcing rings. Elastic and fibrous cartilage are less widely distributed. Flexibility is the hallmark of elastic cartilage, found for example in the outer ear, and is a consequence of its large content of elastic fibers. Meniscus and annulus fibrosus are fibrous cartilages in which bundles of banded fibers, similar to those in tendon, resist tensile forces generated by the load-bearing functions of these tissues. These classes of cartilage will not be considered further in this review. The biomechanical requirements of cartilage are met by the extracellular matrix that constitutes the predominant fraction of the tissue. By contrast, the cellular component, the chondrocytes, occupy a relatively small volume, with each cell embedded into extracellular matrix with no direct cell-to-cell contacts. The matrix properties are determined in turn by tissue-specific supramolecular structures, such as periodically banded fibrils, filaments, and proteoglycan aggregates. This review will outline the current frontiers of our knowledge in this field. Cartilage morphology has been studied by light microscopy, electron microscopy, and X-ray diffraction. These techniques have been complemented by biochemical investigation of isolated aggregates and their self-assembly in vitro, and by genetic studies of human diseases and transgenic animals.

III. LIGHT AND ELECTRON MICROGRAPHY At the level of light microscopy, articular cartilage matrix is an almost homogeneous mass of negatively charged material well stained by basic dyes such as alcian blue or toluidine blue. However, staining discontinuities do highlight specific matrix domains of adult articular cartilage. Such domains are much less conspicuous in immature cartilage, e.g. in fetal or neonatal epiphyseal cartilage. Territorial regions are seen near the cell surfaces which are distinct from the interterritorial matrix in the large intervening spaces. In his early investigations of adult articular cartilage by light microscopy, Benninghoff postulated a reinforcement of the tissue by fibers forming arcades with their bases in the subchondral bone and arches at the articular surface [1]. This arcade-like organization of the fibrils buttresses the tissue against shear forces in the superficial layers and secures the tissue to the underlying bone.

Near the osteochondral junction, a prominent lightmicroscopic feature is the sharply defined limit of calcification of the tissue which is termed the tide-mark. Growth plate, the cartilage responsible for the longitudinal growth of long bones, does not show a tide-mark, but otherwise the salient features of its matrix structure are similar to those of adult articular cartilage. Electron microscopy has been extensively used to visualize the supramolecular structures in hyaline cartilage matrix. General features include a network of banded fibrils and an extrafibrillar matrix with no conspicuous ultrastructure that was once referred to as “amorphous ground substance”. In the immediate vicinity of the cells, there is a small space apparently lacking fibrils altogether. This region is known as the pericellular matrix and mainly contains a specialized extrafibrillar matrix that is easily extractable from the tissue during conventional histological fixation (see below). As apparent already by light microscopy, the matrix structure in immature cartilage is simpler than in adult tissue. Fibrils in fetal or neonatal cartilage have a round cross-section with a uniformly small diameter of about 20 nm which exhibit a faint longitudinal banding pattern [2]. They are almost randomly orientated throughout the tissue, but tend to be parallel to tissue or cell surfaces near to those boundaries. The architecture of adult hyaline cartilages is more complex. The fibril diameters are heterogeneous and depend on the precise anatomical localization within the tissue. In articular cartilage, for example, the fibrils strongly align with joint surfaces, thereby forming conspicuous two-dimensional mats [2]. In the intermediate zones between joint surfaces and the junctures with the subchondral bone, fibrils are more randomly distributed in three dimensions. Reconstructions from thick sections show fibrils with a kinked morphology woven together in a threedimensional knit [3]. In deep zones, fibrils are nearly parallel to the long-bone axis. This distribution of fibril diameters and orientations corresponds well to the Benninghoff arcade model described above. The structure of adult cartilage fibrils also varies between the territorial and interterritorial regions defined by light microscopy. As in immature cartilage, the immediate surrounding of the cell consists of an essentially fibril-free pericellular matrix. Further removed from the cell surface there is a weave of 20-nm fibrils forming a basket around the cells. Such basket structures can be isolated after mechanical disruption of the tissue and have been termed chondrons to designate a functional unit comprising a chondrocyte in its immediate extracellular environment [4]. Chondrons may contain more than one cell, in which case individual cells are separated by a thin

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septum of territorial matrix. Moving away from the cell surface, the territorial matrix gradually becomes the interterritorial region, in which fibrils progressively aquire diameters of 50–100 nm and have striking longitudinal cross-banding patterns. The period of the banding pattern is D, the characteristic periodicity of collagen fibrils, which is typically about 65 nm in electron microscopy, but varies with the degree of shrinkage encountered in the preparation of the specimen (see below). With traditional chemical fixation and embedding in hydrophobic resins, a number of features are observed that are now thought to be artifacts. Among these is the formation of lacunae inside the territorial matrix showing cells with apparent pseudopodia anchored in the fibril network [5]. This feature appears to be a consequence of shrinking of the cells and a loss of matrix macromolecules from the cellular environment. Also, in the extrafibrillar matrix, condensations of material into electron dense globules result from the artifactual precipitation of polyanionic proteoglycans [6, 7]. The recent development of improved chemical fixation, cryopreservation and freeze substitution techniques appears to have overcome these problems and allowed a more detailed analysis of cartilage matrix suprastructures. Under these conditions, the original round shape of chondrocytes is preserved and only small tube-like cellsurface projections are visible which may be part of the secretory machinery of the cells. Lacunae, however, are no longer observed. Further, filaments are seen in the pericellular matrix with a diameter of 10–15 nm and a very faint D-periodic banding pattern [7]. Because this feature is a hallmark of collagen-containing fibrils, the filaments may represent precursors of fibrils in the territorial matrix. Such filaments are also seen throughout the matrix, including the interterritorial regions, where they are intermingled in random orientation with the large fibrils. In the interterritorial matrix, the thicker fibrils often fan out into smaller fibrils closely resembling those of the territorial matrix, giving the impression that the well-banded interterritorial fibrils arise by fusion of archetypal 20-nm fibrils [8]. This is supported by observations of fibrils in cross-section: larger fibrils have an irregular outline and are surrounded by round fibrils that closely resemble the 20-nm fibrils. Verification that large fibrils are assembled by fusion of smaller ones rather than by accretion of individual molecules presents a challenge for future studies in cartilage structural biology. Another detail visualized by the observation of unstained sections of cartilage vitrified at high pressure is the presence of a thin water-rich layer surrounding the fibrils [9]. This implies the existence of a previously unknown transition zone between the fibril surface and

the extrafibrillar matrix, perhaps 5 nm in width, which has a lower density than either of these components. It will be of great interest to find the biochemical correlate of this zone. More recent studies have revealed considerable differences in the fibrillar organization in articular cartilage. For example, tube-like structures, consisting of parallelly aligned, thin fibrils are found extensively in the territorial zones in rabbit articular cartilage [10, 11] whereas fibrils are organized into parallel sheets in the proximal joint cartilage of mouse tibiae [12]. The tubes and the sheets of fibrillar matrix in these tissues have been related to the viscoelastic properties and fluid flow upon load bearing. The biomechanical properties of bovine articular cartilage change with age, diverge within different matrix compartments, and correlate with the density of the fibrilar networks [13].

IV. BIOCHEMISTRY OF CARTILAGE A vast amount of information is available on cartilage matrix macromolecules, which is discussed in detail in Chapters 1-5 of this volume. The extrafibrillar matrix is a complex of carbohydrates and proteins with a high negative charge density which leads to the binding of large amounts of water. Its main macromolecules are aggrecan, hyaluronan, and link protein (LP) which, together, form bottle-brush-like aggregates that have been reconstituted from the purified components and observed by electron microscopy of rotary shadowed preparations [14–16]. Such complexes are known to bind matrilin-1 (also called cartilage matrix protein, CMP) as a further component [17]. Although a considerable degree of structural organization has thus been demonstrated for isolated extrafibrillar matrix components, electron microscopy has failed to visualize the ultrastructure of these components in tissues. Cartilage fibrils are mostly proteinaceous and contain collagens II and XI. Fibrils initially deposited by chondrocytes also contain collagens IX and XVI in a mutually exclusive manner [18]. Such fibrils, here termed prototypic cartilage fibrils, preferentially or exclusively occur in the territorial matrix zones, have a uniform diameter of ca. 20 nm and exhibit a weak D-periodical banding pattern. The collagens also interact with noncollagenous components, most notably with leucine-rich proteins or proteoglycans. Decorin is the most abundant representative of this protein family [19, 20]. Furthermore, binding to collagens has also been reported for fibromodulin, biglycan, matrilins-1 and -3, and cartilage oligomeric matrix protein (COMP; also called thrombospondin-5).

410 COMP directly binds to collagens in a zinc-dependent manner [21] and has been shown to be a component of the fibril periphery in situ (Budde et al., unpublished). Matrilins-1 and-3 are particularly interesting components of the fibril periphery since they have a strong binding affinity for aggrecan [17] and also bind to fibrillar components, either directly to collagens or to COMP (Budde et al., unpublished). Other cartilage components described in Chapter 4 are likely to reside within the extrafibrillar matrix, but their localization also remains to be established. The major collagenous component of hyaline cartilage is collagen II which comprises up to 50% of the dry weight of the tissue. Collagens II and XI are defined as fibril-forming collagens in that they can form fibrils as pure proteins. The other fibril-forming collagens, types I, III, and V, were conventionally thought to be absent from cartilage, but type I is reported to comprise as much as 10% of the collagenous component of normal hyaline cartilage [22]. Collagen III is a very minor component and occurs together with collagen II in the same cartilage fibrils [23]. Collagen V with the chain composition α1(V)2α2(V) is not a cartilage protein but, in adult articular cartilage, about one-half of the α1(XI)-chains in collagen XI are replaced by structurally similar α1(V)chains [24]. The levels of the minor collagens are reported to increase in osteoarthritic cartilage [25–27] but this may be a consequence of increased solubility of collagens in general in diseased tissues. Minor nonfibril-forming collagens that have been reported in cartilage include type VI [22, 28], and types XII and XIV [29]. Collagen VI predominantly occurs in the territorial regions or in isolated chondrons [30]. The protein forms filamentous structures with a periodicity of 100 nm [31] that are independent of D-periodically banded fibrils. Collagen VI, however, also can be organized into hexagonal lattices by co-polymerizing with biglycan or, to a lesser extent, with decorin [32]. By contrast, collagens XII and XIV are likely to be constituents of D-periodic fibrils in skin [33] but their exact suprastructural organization in cartilage still remains to be determined. The polypeptides of fibril-forming collagens consist of uninterrupted long sequences of repeating Gly-X-Y amino acid residue triplets, where X and Y are arbitrary, but are frequently proline and hydroxyproline, respectively. Three chains of such a repeating primary sequence fold into a triple-helical collagen molecule with a length of 300 nm and a diameter of 1.5 nm [34]. Molecules of collagen pack laterally into “quarter-staggered” arrays as shown in Figure 1, in which adjacent molecules are staggered longitudinally by a multiple of D = 67 nm, which

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Figure 1

Changes in cartilage fibril structure with diameter. (A) Uniform diameter 20-nm fibrils of quarter-staggered collagen molecules illustrating the gap–overlap structure. Collagen XI molecules (gray) are shown with amino-terminal propeptides protruding from the surface of the fibril, which may act to control diameter. Collagen IX (hatched) is shown at the surface with the COL3 and NC4 (spherical) domains projecting from the fibril axis. The dark zig-zag represents the glycosaminoglycan (GAG) component covalently attached to the type IX molecules. (B) Intermediatediameter fibril showing both collagen IX and decorin incorporated into the surface. GAG presented at the surface derives from both collagen IX and decorin. (C) Large-diameter fibril showing only decorin, and its attendant GAG on the surface. Collagen IX, perhaps processed, may remain in the interior of the fibril.

corresponds to 234 amino acid residues [35]. The molecules thus are about 4.4 D in length, which gives rise to the well-known gap–overlap longitudinal structure of collagen, where the gap, or hole zone, is about 0.6 D in length [36]. The longitudinal D period is the most obvious feature of collagenous fibrils both in electron microscopy and by X-ray diffraction, the latter technique giving the precise value of D for fully hydrated native specimens. Dermatan sulfate and keratan sulfate proteoglycans are important structural components of fibrils in cartilage and have been visualized in tissues and isolated fibrils by staining with the cationic dye cupromeronic blue under conditions of critical electrolyte concentration [37]. Filamentous chains of proteoglycan are seen with highly regular D-periodic distribution. These have been interpreted as integral components of cartilage fibrils and serve to tether fibrils in position within the matrix. Decorin has been shown to bind to individual collagen I and II molecules near the N-terminal end [38]. This location would place it at the “d” band within the gap of the fibril structure, a location to which proteoglycan has been mapped by cupromeronic blue staining [39]. The proteinaceous part of decorin is a member of a large family of proteins containing leucine-rich repeating (LRR) sequences. The structure of ribonuclease inhibitor,

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another member of the LRR family, though not obviously related to decorin, has been solved by X-ray crystallography [40]. A detailed structural model of decorin binding based on homology to the horseshoe-shaped ribonuclease inhibitor structure has been proposed [41]. Energy calculations of the model support the assignment of the binding to the “d” band of collagen fibrils, but other nearby sites are also possible. Fibromodulin, like decorin, another LRR proteoglycan, has been shown to bind specifically to collagen II [42]. Decorin and fibromodulin have one binding site each per collagen and do not compete with each other [43, 44]. Hence, they must occupy separate specific sites on the collagen molecule. Thus, proteoglycans, including decorin, fibromodulin, and collagen IX, appear to have important specific roles in establishing and maintaining the fibrils of cartilage.

notochord, the collagen II-containing fibrils are apparently crystalline, but the uniform diameter fibrils are too small, ca. 20 nm, to give high-resolution diffraction [53]. Both in this tissue and in mammalian cartilages the average center-to-center intermolecular spacing in the fully hydrated state is ca. 1.7 nm, significantly higher than in collagen I-containing tissues [54]. This is consistent with a higher number of glucosyl-galactosyl-hydroxylysine residues found in collagen II: the levels of this posttranslational modification are 50–100% higher than in type I collagen. This results in a higher intrafibrillar water content in cartilage [55, 56]. Dried cartilage shows approximately the same intermolecular spacing (ca. 1.1 nm) as tendon, demonstrating that the collagen II molecular diameter is not significantly increased by the additional sugar content. The other fibril-forming collagen of cartilage, type XI, has an even higher degree of glycosylation than collagen II; however, there are no X-ray diffraction data specific for this collagen type. Glycation, the nonenzymatic reaction by glucose with protein amino groups that occurs naturally in tissues with slow protein turnover, has shed light on the lateral organization of collagen in fibrils. Rat tail tendon was subjected to cross-linking by ribose as a model for the age-related effects of glycation [57]. It was found that the average intermolecular spacing increased with the degree of cross-linking. This was interpreted as showing that the separation of collagen molecules is a weakly constrained equilibrium such that there is little energetic cost involved in swelling or shrinking the structure by the exchange of water [53]. This supports the idea that collagen II has increased intrafibrillar water because of its increased post-translational hydroxylysyl glycosylation. The biomechanical effects of age-related glycation in cartilage fibrils have also been modeled by ribosylation, but resulting changes in intrafibrillar water content were not reported [58].

V. STUDIES OF FIBRIL STRUCTURES BY X-RAY DIFFRACTION X-ray diffraction has been used to determine the structure of collagen in rat tail tendon nearly to atomic resolution [45, 46]. For this reason and because of the biochemical similarities in fibril-forming collagens, the fibrils in this tissue have often been considered as model structures of collagen-containing fibrils in general, a notion that later has been revised [47]. Rat tail tendon is almost unique among connective tissues in that collagen molecules are packed in molecular crystals that diffract to high resolution. A analysis is consistent with a tissue comprised of microfibrils of five-triple helical collagen molecules twisted into left-handed strands distorted by crystal packing forces into a quasi-hexagonal array [48, 49]. These arrays exhibit coherently diffracting domains up to 100 nm in lateral size, considerably smaller than the size of the fibrils (up to 600 nm) in which they reside. Thus, the large fibrils seen by electron microscopy are not single crystals, but mosaics of smaller crystallites [50]. The average center-to-center intermolecular spacing in hydrated rat tail tendon is ca. 1.5 nm. Most other tendons do not give crystalline patterns, but X-ray diffraction can still be used in these specimens to evaluate the intermolecular spacing. In other fully hydrated, collagen I-containing tissues, this spacing is also near 1.5 nm [51]. In the dry state, this distance collapses to about 1.1 nm, demonstrating that the collagen molecules in fibrils are separated from each other by substantial quantities of intrafibrillar water [52]. Cartilage fibrils have also been extensively studied by X-ray diffraction. In one case, the sheath of the lamprey

VI. STRUCTURE OF FIBRIL FRAGMENTS OBTAINED BY MECHANICAL DISRUPTION OF TISSUE Another approach to study the structure of cartilage fibrils has been to prepare fragments of fibrils from cartilage or from chondrocyte cultures. Such fibril preparations, contrasted by negative or positive stain with or without immuno-gold labeling, or by rotary shadowing, have yielded considerable information. Both cell cultures and tissues yield the thin fibrils characteristic of cartilage which have been shown by immunostaining to be heterotypic, in that individual fibrils contain collagens II,

412 IX, and XI. Biochemical analysis of fragments isolated from chick embryo cartilage revealed a mass proportion of 8:1:1 for these collagen types, respectively [59]. Interestingly, collagen XI in fibril fragments from chick embryo sterna exhibited a strong immunochemical masking toward antibodies raised against the long triple helix of the protein. Upon disruption of the collagen packing by partial digestion with pepsin, however, this immunochemical masking was released [60]. Employing immunogold labeling with antibodies directed to the aminoterminal nonhelical domain, collagen XI was localized to fibrils only of small diameter in tissue sections of human rib and growth plate cartilage [61]. By contrast, undigested fibril fragments were readily labeled with antibodies to the pepsin-treated forms of both collagens II and IX [60]. Taken together, these results raise the possibility that collagen XI is buried within the fibrillar body, and that, by forming a core structure, collagen XI regulates apposition of types II and IX onto the fibrils. Rotary shadowed fibrils exhibit a D-periodic occurrence of collagen IX, with the amino-terminal domains NC4 and COL3 projecting from the fibril axis (see Chapter 1 for a discussion of these domains). This has been interpreted as showing collagen IX to be a surface component [59]. However, fibrils of older mammalian cartilage are biochemically heterogeneous, with the detailed composition depending on which tissue layer is sampled. Small, uniform fibrils characteristically have collagen IX at their surface, whereas large fibrils have decorin. Some intermediate diameter fibrils stain for both collagen IX and decorin [20] (Fig. 1). Since both of these molecules are glycoproteins carrying dermatan sulfate chains in cartilage, the fibrils are studded with anionic groups which could help to secure the extrafibrillar matrix to the fibrils.

VII. STUDIES OF COLLAGEN CROSSLINKING IN CARTILAGE FIBRILS Naturally occurring cross-links between collagen polypeptides in cartilage have been used as indicators for the fibrillar organization (the occurrence and chemistry of such cross-links is discussed in Chapter 2). Collagen IX purified from steer cartilage was found to have 1.7–2.3 mol of cross-linking residues to collagen II per mol of collagen, consistent with the heterotypic nature of cartilage fibrils [62]. If it is assumed that the collagen IX molecules lie parallel to the fibril axis, then the locations of these links indicate that the collagen IX is oriented antiparallel to the type II and covering both the gap and the overlap region (Fig. 1). However, it is also possible to

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constuct models in which the type IX wraps around, or partially penetrates, the fibril, in which cases the molecular configuration cannot be specified. Collagen IX is also cross-linked to other molecules of collagen IX. Collagen XI is cross-linked as well, largely to other type XI molecules, but to a small degree also to collagen II [63]. This observation has been interpreted as supporting the idea that collagen XI forms a core on which collagen II is deposited. Cross-links to other fibrillar and nonfibrillar components have not yet been described.

VIII. RECONSTITUTION OF AGGREGATES FROM SOLUBLE COLLAGENS AND OTHER MACROMOLECULES One of the fascinating questions concerning cartilage matrix structure is what mechanism controls the diameter of the thin, uniform fibrils. Fibril reconstitution experiments with purified components have been used to approach this question. In such experiments, collagens in solution are subjected to temperature and buffer conditions under which they spontaneously aggregate into fibrils. Chicken sternal chondrocytes from 17-day embryos grown in agarose maintain the cartilage phenotype and synthesize uniform 20-nm fibrils. Collagens II, XI, and IX have been purified from these cultures by salt fractionation and ion exchange chromatography, yielding essentially pure components in fibril-competent form [64]. Fibrils reconstituted from the mixture of the three collagen types directly extracted from the cultures gave uniform D-periodic fibrils closely resembling authentic 20-nm fibrils by their appearance in the electron microscope after negative contrasting. Collagen XI alone also formed small, uniform fibrils of about the same diameter. The kinetics of in vitro fibrillogenesis was followed by turbidometry and by electron microscopy. As reported previously [65], collagen II alone developed turbidity only after a lag time of 90–120 min. After this lag phase, the protein formed wide tactoids which, in the electron microscope, appear as relatively short, D-periodically banded aggregates with tapered ends and without apparent diameter control. Aggregation of collagen XI alone resulted in a hyperbolic increase in turbidity with no discernible lag phase at all. Immediately after initiation of in vitro fibrillogenesis, within the limits of time resolution of about 1 min, the protein assembled into long and flexible, small-diameter filaments. As turbidity developed to moderate amplitudes, the filaments turned into very weakly cross-banded fibrils with tightly controlled diameters of ca. 20 nm.

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Mixtures of collagens II and XI, at mass proportions of up to 8:1, respectively, produced turbidity characteristics without lag phase similar to those of pure collagen XI and the final fibrils also had a closely similar appearance. Therefore, the diameter control is inherent to these two components alone. In addition, the absence of a lag phase in the binary reconstitution system of collagens II and XI indicated that collagen XI was a powerful nucleator of fibril formation in cartilage. Furthermore, the 8:1 limit of collagen proportions and the 20-nm diameter match the corresponding properties of authentic chick embryo sternal cartilage fibrils. Thus, fibrils jointly formed by collagens II and XI have characteristics that vastly differ from those of aggregates formed by both individual collagens and, therefore, represent macromolecular alloys [66]. At larger proportions of collagen II, two-phase turbidity curves were observed and 20-nm fibrils and tactoids coexisted in the final reconstitution products. However, unlike the case of pure collagen II undergoing in vitro fibrillogenesis, tactoids formed in these supercritical mixtures of collagens II and XI were already present within the lag period of the second increase in turbidity. This necessitates that such tactoids were heterotypic, in that they contain both collagen types, and that the decision on the shape of the final aggregate occurs during the early phase of assembly and is determined by the biochemical composition of the early aggregates. This conclusion is illustrated by the drawing in Figure 2. In addition, many tactoids were continuous with the 20-nm

fibrils, suggesting that tactoid formation nucleated on fibrils [64]. Unlike collagen II extracted from natural sources, collagen I purified from tendons or skin can be reconstituted into large, strongly banded fibrils. This self-assembly system, again, is associated with a considerable lag phase of turbidity development and displays little if any diameter control. However, the reconstitution products share many morphological features which are characteristic for natural collagen I-containing fibrils. They are rigid ropelike objects, with each individual fibril maintaining a constant diameter over very long distances. Ends are very rarely observed, since the length of the reconstituted fibrils usually far exceeds the dimensions of the apertures of commonly used EM grids [67]. However, even exceedingly small quantities of minor fibrillar collagens, such as collagen XI, can profoundly alter both the aggregation kinetics and the morphology of the reconstitution products. The length of the lag phase of turbidity development is strongly reduced by collagen XI but, unlike in mixtures of collagens II and XI, occurs even when collagen XI represents a large or even predominant mass fraction. Therefore, collagen XI molecules can interact with those of collagen I and strongly enhance the aggregation efficiency of collagen I. The strict diameter control characteristic for the collagen II/XI system, however, is not observed in mixtures of collagens I and XI. In addition, fibrils reconstituted from collagens I and XI are composites containing alloyed cores of collagens I and XI and homotypic collagen I sheaths varying considerably

Figure 2

Illustration of the different aggregates resulting from fibril formation with different stoichiometries of collagens II and XI. (A) Tactoids are formed in mixtures with an excess of collagen II (greater than 8:1), probably as a result of separation of a collagen II-rich phase characterized by tight lateral packing of identical molecules. (B) The native proportion of collagens II and XI result in uniform 20-nm fibrils.

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in thickness. All these differences between mixtures of collagens I and XI or collagens II and XI occur in spite of the extraordinary sequence similarity in the collagenous regions of all polypeptides constituting collagens I and II [68]. The causes underlying these differences remain to be identified.

IX. STUDIES OF TRANSGENIC MICE AND OF HUMAN GENETIC MATRIX DISEASES Studies of mutations in genes encoding matrix macromolecules have also contributed to our current understanding of cartilage matrix organization. A large repertoire of mutations in human and animal genes for almost all macromolecules discussed above has been reported. Information comes from experimentally induced mutations in transgenic mice as well as from naturally occurring mutations in human and animal diseases. Consequences of mutations in collagen genes to cartilage suprastructure will primarily be discussed here. The mutations and their pathological consequences have been catalogued in recent reviews [69, 70]. In-frame deletions resulting in a loss of peptide sequences within the triple-helical domain in the collagen II gene (COL2A1 in man or Col2a1 in mice) lead to chondrodysplasia of variable severity, depending on the extent and the location of the deletion within the gene. Such mutations result in shortened α1(II) chains that are incorporated into homotrimeric collagen II molecules, together with either normal or other mutated chains. This explains the dominant phenotype of the mutations. However, the penetrance of the mutations varies not only between mice with different genetic backgrounds but even between individual inbred animals. The ability of cells to eliminate mutated mRNA or polypeptide species may be subject to variation between individual animals, which may explain the heterogeneous severity of the consequences of a given mutation. Triple-helix formation is impaired in affected molecules which, thereby, frequently have a lowered thermal stability and a compromised competence of incorporation into fibrillar aggregates. Due to the D-periodic regularity of collagen II molecules within the fibrils, incorporation of shortened molecules, even though they are secreted by the cells in reduced amounts, will destroy the entire fibrillar organization. This translates into a generalized injury to the structure of the cartilage matrix and its functions. Point mutations, particularly those exchanging glycine residues in the Gly-X-Y repeats of the triple-helical domain, have similar consequences. Such mutations have shown that the formation of

cartilaginous tissue does not depend on the presence of collagen II: in patients with hypochondrogenesis or achondrogenesis, collagen II, though synthesized, was not secreted by chondrocytes, but an abnormal mixture of collagens I and III was present [71, 72]. The resulting cartilage even matured sufficiently to produce collagen X. Bone formation, however, was severely disrupted in these lethal mutations. In general, abnormalities within carboxyterminal regions of the triple helix tend to result in more severe phenotypes, since collagen folding proceeds from the carboxyl- towards the amino-terminal end in a zipperlike fashion [73]. Interestingly, transgenic mice harboring extra copies of the normal Col2a1 gene have a perinatally lethal phenotype. Although no gross abnormalities were found in their cartilages, wide and strongly banded fibrils occurred in the interterritorial regions. In these animals, the balance in the amounts of collagens II and XI is altered due to an overexpression of collagen II resulting from transcription and translation of the transgenes. This agrees well with the formation in vitro of tactoids rather than normal cartilage-like fibrils from solutions containing collagen II above physiological levels (see above). Mice homozygous in a nonfunctional collagen II gene (type II knockout mice) also have been reported [74, 75]. These animals show extensive abnormalities of their endoskeleton but, surprisingly, survive until birth. They extensively develop cartilage tissue in which, however, collagen II is substituted, at least in part, by type I. Poorly organized fibrils occur in much smaller numbers than in normal cartilage and lack discernible longitudinal banding patterns as well as clearly defined lateral limits. The reasons for this fibrillar disorganization are not immediately clear. However, since the α3(XI) chains are the same gene product as the α1(II) chains (see Chapter 1), collagen XI production is also affected by the knockout of the Col2a1 gene. The thermal stability of collagen XI is lowered and, hence, the protein occurs in lower quantities due to degradation in the tissue. However, collagen XI very strongly nucleates formation of heterotypic fibrils containing collagens I and XI, whereas collagen I alone only slowly forms fibrils at relatively high monomer concentrations [68]. Thus, in cartilage of mice with homozygously inactivated Col2a1 alleles, the virtual absence of banded fibrils is a consequence of collagen XI deficiency rather than a lack of the major cartilage collagen type II. Consistent with this notion, the abnormally produced collagen I, due to a lack of incorporation into supramolecular aggregates, is extractable from cartilage in markedly increased quantities [75]. Heterozygous mice with only one inactivated Col2a1 allele survive into adulthood and

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also can reproduce. However, the lowered dose of active Col2a1 genes results in premature vertebral endplate ossification and mild disk degeneration [76] and weakens the fibrillar organization in articular cartilage [77]. If heterozygous inactivation of the Col2a1 gene results in reduced translation of α1(II) polypeptide chains in articular chondrocytes collagens II and XI would occur at molar fractions different from 8:1. Hence, prototypic cartilage fibrils would be formed at abnormally low levels [64], which may well explain the cartilage phenotype of heterozygous collagen II knockout mice. A variety of mutations affecting the genes of collagen XI chains also have been discovered and, again, disturbed formation of fibrils as a consequence of collagen XI abnormalities well explains the phenotypes observed. A single base deletion in the gene encoding the α1(XI) chain occurs naturally in the cho/cho mouse strain [65]. This represents a frame-shift mutation leading to a prematurely terminated α1(XI) polypeptide which either is not efficiently synthesized or is prematurely degraded. Collagen XI protein is absent in the cartilage of these mice and, again in keeping with in vitro fibrillogenesis of cartilage collagen mixtures, the cartilage of homzygous cho/cho mice contains disorganized fibrils with exceptionally large diameters. Analogous consequences are seen in patients with Stickler syndrome where a splice-site mutation probably causes the production of shortened α2(XI) chains [66]. Taken together, these observations are consistent with the notion that collagen XI is required as a powerful nucleating agent in the formation of cartilage fibrils (see above). If production of collagen XI is compromised, collagen II not only remains unincorporated into supramolecular aggregates but, as a consequence of this lack of suprastructural integration, is degraded and largely absent from the tissues [78]. The deletion of collagen IX in mice [67] by inactivation of the Col9a1 gene [68] results in a remarkably mild phenotype. The mice show no extensive abnormality in skeletal development and the fibrils in cartilage of affected animals also are close to normal. However, the birth rate of homozygous mice from heterozygous parents appears to be reduced to about one-half of the expected Mendelian proportion, which may be related to serious skeletal malformations incompatible with further progression of fetal development (Failing, Jongebloed, and Bruckner, unpublished). In addition, cartilage integrity appears to be impaired only in older animals that show premature onset of osteoarthritis-like cartilage degeneration. Again, this is consistent with in vitro fibrillogenesis showing a reduced stability of collagen IXdeficient fibrils [64] and suggests that the main role of collagen IX is at the level of tissue organization of fibrils

in cartilage, as originally proposed by Müller Glauser et al. [79] and revisited in greater detail in cross-linking studies by Eyre et al. [24, 80]. Nevertheless, it is surprising that bone development is almost normal in these animals in view of the fact that it is the thin fibrils arising in fetal and young cartilage which are rich in collagen IX [81]. It is tempting to speculate that the function of collagen IX is redundant, at least in part, in cartilage fibrils. Likely candidates are collagen XVI [18, 82] or other FACITs, including collagens XII, XIV, or XXII [83]. An exon-skipping mutation in the COL9A2 gene of patients with multiple epiphyseal dysplasia [84] leads to the transcription of a shortened α2(IX) mRNA expected to result in α2(IX) chains lacking 12 amino acids within the Col3 domain which protrudes from the fibril surface [59]. It still remains to be determined whether, in analogy to the Col9a1 knockout mice, the production of shortened α2(IX) chains leads to an overall absence of collagen IX. However, fully functional protein is not likely to be synthesized in the affected tissues. Consistent with mice lacking collagen IX, the consequences of this mutation do not include severe skeletal abnormalities. Again, the genotype–phenotype correlation agrees with the notion that collagen IX is not essential for fibrillogenesis in cartilage but that the protein is required for long-term stability of the tissue. If an abnormal variant of collagen IX exists in the patient cartilage, it will be interesting to determine the mode in which a shortened Col3 domain causes impairment of the interaction between fibrils and the extrafibrillar matrix.

X. CORRELATING STRUCTURE WITH THE BIOMECHANICAL ROLE OF ARTICULAR CARTILAGE A major function of articular cartilage is to enable unhindered motion of joints by providing a smooth, low-friction gliding surface, and by cushioning the bone against stress and impact. These properties are achieved by prestressing the cartilagenous material through the interaction of the fibrils with their proteoglycan-rich matrix. The matrix has a high fixed-charge density, deriving from the negatively charged groups of its constituent glycosaminoglycans, chiefly those of aggrecan. This charge is responsible for the high degree of hydration of the tissue, which in turn causes the matrix component to swell strongly. Swelling is counteracted by the cartilage fibrils, which act to keep the glycosaminoglycans compressed and relatively dehydrated in comparison to their free, equilibrium hydration (see also Chapter 5). The collagen fibril network is firmly

416 attached to the extrafibrillar matrix by a high density of noncovalent interactions between proteins, such as between fibrils and decorin, and between glycosaminoglycans. In turn, some of the glycosaminoglycans are covalently bound to proteins. The arrangement of fibrils into arcades is precisely the architecture needed to counteract the stress. The physical properties of cartilage have been extensively studied by mechanical testing, with the observed properties being modeled as a viscoelastic [85], or poroviscoelastic [86] material. Most such modeling investigations have the drawback that the tissue is regarded as a homogeneous substance, but the latter study did investigate samples with the surface layer, the collagen-rich top of the arcades, removed. In this case it was found that exudation of fluid under pressure was more rapid than in intact tissue, demonstrating the role of the superficial zone in controlling interstitial movement of fluid. The intratissue pressure in cartilage has been estimated by subjecting samples to various osmotic pressures and measuring changes in hydration [87]. Osmotic pressure was controlled by use of calibrated solutions of polyethylene glycol (PEG) at various concentrations. It was found that the unperturbed internal pressure of cartilage was ca. 3.9 bar. The mass of normal tissue varied surprisingly little with osmotic pressure changes: normal specimens transferred from 0.15 M NaCl to 0.015 M NaCl swelled by only 1–2 %. The collagen fibrils too respond to changes in osmotic pressure: increases in osmotic pressure reduce the hydration, and thus the intermolecular spacing, of the collagen triple-helices [48]. The tension in the fibrils has been calculated and shown to be easily within the known tensile limits of collagen [88]. Very large increases in osmotic pressure by PEG, up to 10 bar with a concomitant 20% mass loss, were required to make the collagen fibrillar network go “limp” in normal cartilage [87]. The continuous tension to which cartilage fibrils are subjected is an unusal role for collagen. In other tissues collagens are called on to resist tension only transiently. Further, the fibrils must be synthesized and repaired while maintaining tension, requiring a mechanism that is not well understood. It is perhaps not surprising that a number of medically important conditions can develop in such a system. Ameliorating the biological problem of tissue maintenance is the exceptionally long life of cartilage components under normal circumstances. Collagen II has a half-life of more than 200 years in humans [58]. Consistent with this is the rate of accumulation of glycationderived cross-links in cartilage collagen. Such cross-links increase linearly with age after 20, when the organism stops growing, and show no evidence of reaching a plateau

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level at advanced age [58]. By these measures, then, the collagen is never turned over. The rate of aspartic acid residue racemization has been used to show that the half-life of different species of aggrecan ranges from 3.4 to 25 years in man [89]. Thus, in the absence of disease or injury, cartilage is highly stable. The mechanical properties of cartilage are drastically affected in osteoarthritis. An osteoarthritic specimen transferred from 0.15 M NaCl to 0.015 M NaCl swelled by more than 10%, demonstrating the effects of destruction of the fibrillar net in this disease [87]. In this condition, the tissue has not the means to resist water uptake by the glycosaminoglycans. Thus, therapies that aim to make up the observed loss of glycosaminoglycans in osteoarthritis probably will not work as expected; the fibrillar network needs to be restored.

XI. MODELS OF CARTILAGE FIBRIL STRUCTURE What models of fibril structure are compatible with these observations? The classical scheme has collagen XI forming a core or scaffold on which the collagen II is deposited. Collagen IX then serves to coat the structure and arrest the diametral growth. We have recognized that collagen IX is not necessary to achieve the morphology of the prototypic 20-nm cartilage fibril, but the model is still attractive. First, it appears to account for the observed greater difficulty in extracting type XI from cartilage as compared to type II. Second, the cross-link data showing that collagen XI is cross-linked mostly with other homologous molecules suggests that the protein forms a filamentous core of the fibrils. The core model for cartilage fibrils, unfortunately, does not easily provide a mechanism for diameter control in the absence of type IX: the fibrils should be indefinitely large. However, collagen IX at the surface of cartilage fibrils may be suitable to stabilize interfibrillar connections by integration of its triple helical domain into more than one fibril. This concept has initially been derived from a conspicuous concentration of collagen IX immunogold labeling at fibrillar intersections [60] and has since been substantiated by studies on cross-linking of collagens within cartilage fibrils [80] (Fig. 3). The possible existence of a core filament has also been discussed in conjunction with the analogous collagen I– collagen V system in cornea. Fibrils in cornea too have tightly controlled diameters, a property that is responsible for corneal transparency. One suggestion for the control of diameter in these fibrils, and the relative inaccessibility of their collagen V component to antibody labeling, is

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Figure 4

Illustration of the core model vs. the alloy model of uniform cartilage fibrils. (A) A cross-section of the core model has collagen XI (gray) at its center, collagen II (open circles) apposed around it, and collagen IX (filled) capping the structure to limit diametral growth. It is now known that collagen IX is not required for diameter control. (B) The alloy model posits a random commingling of collagen II and XI, with either molecule being able to occupy any lattice position. Diameter control could be achieved by the necessity to accommodate the collagen XI propeptides at the surface (cf. Fig. 1). Lattice faults (*), arising from incommensurate packing of collagens II and XI, are important in both models, either providing a binding site for collagen IX in the core model, or destabilizing largediameter fibrils in the alloy model.

Figure 3 Model of the fibrillar network in cartilage and the role of collagen IX and its known cross-links between collagen II and IX and between IX and IX collagen molecules. (A) Covalent cross-links between the C-terminal, nonhelical domain NC1 and a triple-helical portion of collagen II initiate network formation, followed by other cross-links between collagens II and IX or two collagen IX molecules. (B) A model illustrating a possible cross-over of collagen IX molecules between neighboring collagen II/XI fibrils. Reproduced with permission from Eyre, D. R. et al. J. Biol. Chem. 2004; 279, 2568–2574.

could incorporate a certain amount of collagen II up to approximately 8:1, the most favorable composition for the alloy, without structural modification. Beyond that ratio, the structure doesn’t form and phase separation takes place. Collagen XI would be distributed throughout the fibril in this model. This is not necessarily contradicted by the type XI cross-link data because of the relatively great length and flexibility of the molecules. The resistance to extraction of type XI in tissues can be a simple consequence of the cross-linking, rather than of any structural model.

XII. FUTURE PERSPECTIVES the existence of a core fiber of type V [90]. As in the case of cartilage fibrils, this model does not easily provide a mechanism for diameter control in the absence of other components. A different model is that of the biological alloy, in which collagens II and XI randomly commingle to form diameter-controlled fibrils as long as the ratio is 8:1 or less [64] (Fig. 4). This model is attractive because it explains why fibril diameter does not depend on composition. With the core model, one would expect mixed fibrils to be larger than the core material alone, which is not observed. Diameter control is an innate property of the alloy, mediated by collagen XI. It is known that most of the N-propeptides of type XI are retained in the tissue [91]. The requirement to accommodate these bulky moieties, probably on the exterior of the fibrils, could inhibit diametral growth [92, 93]. This class of alloys

A great deal of information has been accumulated on the identity and structure of cartilage matrix macromolecules. However, much remains to be learned about one of the quintessential characteristics of these components: their assembly into insoluble suprastructural aggregates. The fibrillar organization of cartilage collagens is gradually emerging and we are beginning to understand the assembly of the bulk components of the extrafibrillar matrix, aggrecan, and hyaluronan, but the aggregate structures of most of the other macromolecules identified as cartilage matrix components is either unknown or controversial. As is true for most extracellular matrices, even less is known about the formation of structure at the next hierarchic level, i.e. the principles directing the assembly of cartilage matrix from fibrils and the extrafibrillar matrix. As our knowledge expands about additional molecular

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components of fibrils or the extrafibrillar matrix our chances to formulate viable working hypotheses will increase. For example, the definition of fibrillar components other than collagens, particularly those at surfaces, will be one of the challenges in the immediate future. Likewise, it will be interesting to learn about the identity of molecular components of the extrafibrillar matrix which occur in the immediate vicinity of fibrils. Methods will have to be devised to reconstitute matrix domains from suprastructural subunits to understand the forces driving matrix assembly. These efforts will be strongly supported by expanding our knowledge of the function of matrix macromolecules in situ. This goal will be achieved by defining in greater detail the connection between the genotype and the phenotype in human or animal genetic disorders or in transgenic animals. Concurrently, the refinement of techniques such as X-ray diffraction and electron and atomic force microscopy, combined with specific labeling, will provide additional opportunities to elucidate cartilage matrix structure at high resolution. The combined information will lay the groundwork of our understanding of the biomechanics of cartilage and its metabolic regulation through cell–matrix interactions under normal conditions, as well as its abnormalities under pathological conditions.

References 1. Benninghoff, A. (1922). Über den funktionellen Bau des Knorpels. Anat. Anz. 55, 250–267. 2. Hunziker, E. B. (1992). Articular cartilage structure in humans and experimental animals. In “Articular Cartilage and Osteoarthritis” (K. E. Kuettner, R. Schleyerbach, J. G. Peyron, and V. C. Hascall, eds.), pp. 183–199. Raven Press, New York. 3. Broom, N. D., and Silyn-Roberts, H. (1989). The 3-dimensional knit of collagen fibrils in articular cartilage. Connect. Tissue Res. 23, 261–277. 4. Poole, C. A. (1997). Articular cartilage chondrons: Form, function and failure. J. Anat. 191, 1–13. 5. Hunziker, E. B., Herrmann, W., and Schenk, R. K. (1983). Ruthenium hexamine trichloride (RHT)-mediated interaction between plasmalemmal components and pericellular matrix proteoglycans is responsible for the preservation of chondrocyte plasma membranes in situ during cartilage fixation. J. Histochem. Cytochem. 31, 717–727. 6. Studer, D., Michel, M., Wohlwend, M., Hunziker, E. B., and Buschmann, M. D. (1995). Vitrification of articular cartilage by highpressure freezing. J. Microsc. 179, 321–332. 7. Hunziker, E. B., Wagner, J., and Studer, D. (1996). Vitrified articular cartilage reveals novel ultra-structural features respecting extracellular matrix architecture. Histochemical Cell Biology 106, 375–382. 8. Hunziker, E. B., Michel, M., and Studer, D. (1997). Ultrastructure of adult human articular cartilage matrix after cryotechnical processing. Microsc. Res. Tech. 37, 271–284. 9. Studer, D., Chiquet, M., and Hunziker, E. B. (1996). Evidence for a distinct water-rich layer surrounding collagen fibrils in articular cartilage extracellular matrix. J. Struct. Biol. 117, 81–85.

10. ap Gwynn, I., Wade, S., Kääb, M. J., Owen, G. R., and Richards, R. G. (2000). Freeze-substitution of rabbit tibial articular cartilage reveals that radial zone collagen fibres are tubules. J. Microsc. 197, 159–172. 11. ap Gwynn, I., Wade, S., Ito, K., and Richards, R. G. (2002). Novel aspects to the structure of rabbit articular cartilage. Eur. Cell. Mater. 4, 18–29. 12. Hughes, L. C., Archer, C. W., and ap Gwynn, I. (2005). The ultrastructure of mouse articular cartilage: collagen orientation and implications for tissue functionality. A polarised light and scanning electron microscope study and review. Eur. Cell. Mater. 9, 68–84. 13. Williamson, A. K., Chen, A. C., Masuda, K., Thonar, E. J. M. A., and Sah, R. L. (2003). Tensile mechanical properties of bovine articular cartilage: Variations with growth and relationships to collagen network components. J. Orthop. Res. 21, 872–880. 14. Hascall, V. C., and Heinegård, D. (1974). Aggregation of cartilage proteoglycans. 1. The role of hyaluronic acid. J. Biol. Chem. 249, 4232–4241. 15. Rosenberg, L., Hellmann, W., and Kleinschmidt, A. K. (1975). Electron microscopic studies of proteoglycan aggegates from bovine articular cartilage. J. Biol. Chem. 250, 1877–1883. 16. Mörgelin, M., Paulsson, M., Heinegård, D., Aebi, U., and Engel, J. (1995). Evidence of a defined spatial arrangement of hyaluronate in the central filament of cartilage proteoglycan aggregates. Biochem. J. 307, 595–601. 17. Hauser, N., Paulsson, M., Heinegård, D., and Mörgelin, M. (1996). Interaction of cartilage matrix protein with aggrecan. Increased covalent cross-linking with tissue maturation. J. Biol. Chem. 271, 32247–32252. 18. Kassner, A., Hansen, U., Miosge, N., Reinhardt, D. P., Aigner, T., Bruckner-Tuderman, L., Bruckner, P., and Grässel, S. (2003). Supramolecular association of collagen XVI to different fibrillar systems in human skin and cartilage. Matrix Biol. 22, 131–143. 19. Miosge, N., Flachsbart, K., Götz, W., Schultz, W., Kresse, H., and Herken, R. (1994). Light and electron microscopical immunohistochemical localization of the small proteoglycan core proteins decorin and biglycan in human knee joint cartilage. Histochem. J. 26, 939–945. 20. Hagg, R., Bruckner, P., and Hedbom, E. (1998). Cartilage fibrils of mammals are biochemically heterogeneous: Differential distribution of decorin and collagen IX. J. Cell Biol. 142, 285–294. 21. Rosenberg, K., Olsson, H., Mörgelin, M., and Heinegård, D. (1998). Cartilage oligomeric matrix protein shows high affinity zinc-dependent interaction with triple helical collagen. J. Biol. Chem. 273, 20397–20403. 22. Wardale, R. J., and Duance, V. C. (1993). Quantification and immunolocalisation of porcine articular and growth plate cartilage collagens. J. Cell Sci. 105, 975–984. 23. Young, R. D., Lawrence, P. A., Duance, V. C., Aigner, T., and Monaghan, P. (2000). Immunolocalization of collagen types II and III in single fibrils of human articular cartilage. J. Histochem. Cytochem. 48, 423–432. 24. Eyre, D. R. (2004). Collagens and cartilage matrix homeostasis. Clin. Orthop. Rel. Res. 427S, S118–S122. 25. Adam, M., and Deyl, Z. (1983). Altered expression of collagen phenotype in osteoarthrosis. Clin. Chim. Acta 12, 25–32. 26. Aigner, T., Bertling, W., Stöss, H., Weseloh, G., and von der Mark, K. (1993). Independent expression of fibril-forming collagens I, II, and III in chondrocytes of human osteoarthritic cartilage. J. Clin. Invest. 91, 829–837. 27. Aigner, T., Glückert, K., and von der Mark, K. (1997). Activation of fibrillar collagen synthesis and phenotypic modulation of chondrocytes in early human osteoarthritic cartilage lesions. Osteoarthritis Cart. 5, 183–189. 28. Hagiwara, H., Schröter-Kermani, C., and Merker, H.-J. (1993). Localization of collagen type VI in articular cartilage of young and adult mice. Cell Tissue Res. 272, 155–160.

Chapter 24 Supramolecular Structure of Cartilage Matrix

419

29. Watt, S. L., Lunstrum, G. P., McDonough, A. M., Keene, D. R., Burgeson, R. E., and Morris, N. P. (1992). Characterization of collagen type XII and type XIV from fetal bovine cartilage. J. Biol. Chem. 267, 20093–20099. 30. Poole, C. A., Ayad, S., and Gilbert, R. T. (1992). Chondrons from articular cartilage. V. Immunohistochemical evaluation of type VI collagen organisation in isolated chondrons by light, confocal and electron microscopy. J. Cell Sci. 103, 1101–1110. 31. Furthmayr, H., Wiedemann, H., Timpl, R., Odermatt, E., and Engel, J. (1983). Electron-microscopical approach to a structural model of intima collagen. Biochem. J. 211, 303–311. 32. Wiberg, C., Heinegård, D., Wenglén, C., Timpl, R., and Mörgelin, M. (2002). Biglycan organizes collagen VI into hexagonal-like networks resembling tissue structures. J. Biol. Chem. 277, 49120–49126. 33. Keene, D. R., Lunstrum, G. P., Morris, N. P., Stoddard, D. W., and Burgeson, R. E. (1991). Two type XII-like collagens localize to the surface of banded collagen fibrils. J. Cell Biol. 113, 971–978. 34. Rich, A., and Crick, F. H. C. (1961). The molecular structure of collagen. J. Mol. Biol. 3, 483–506. 35. Hulmes, D. J. S., Miller, A., Parry, D. A. D., Piez, K. A., and Woodhead-Galloway, J. (1973). Analysis of the primary structure of collagen for the origins of molecular packing. J. Mol. Biol. 79, 137–148. 36. Hodge, A. J., and Petruska, J. A. (1963). Recent studies with the electron microscope on ordered aggregates of the tropocollagen macromolecule. In “Aspects of Protein Structure” (G. N. Ramachandran, ed.), pp. 289–300. Academic Press, New York. 37. Scott, J. E. (1990). Proteoglycan-collagen interactions and subfibrillar structure in collagen fibrils. Implications in the development and ageing of connective tissues. J. Anat. 169, 23–35. 38. Yu, L., Cummings, C., Sheehan, J. K., Kadler, K. E., Holmes, D. F., and Chapman, J. A. (1993). Visualization of individual proteoglycancollagen interactions. In “Dermatan Sulfate Proteoglycans” (J. E. Scott, ed.), pp. 183–188. Portland Press, London.. 39. Scott, J. E. and Orford, C. R. (1981). Dermatan sulphate-rich proteoglycan associates with rat tail-tendon collagen at the d band in the gap region. Biochem. J. 197, 213–216. 40. Kobe, B., and Deisenhofer, J. (1993). Crystal structure of procine ribonuclease inhibitor, a protein with leucine-rich repeats. Nature 366, 751–756. 41. Weber, I. T., Harrison, R. W., and Iozzo, R. V. (1996). Model structure of decorin and implications for collagen fibrillogenesis. J. Biol. Chem. 271, 31767–31770. 42. Hedlund, H., Mengarelli-Widholm, S., Heinegård, D., Reinholt, F. P., and Svensson, O. (1994). Fibromodulin distribution and association with collagen. Matrix Biol. 14, 227–232. 43. Hedbom, E., and Heinegård, D. (1993). Binding of fibromodulin and decorin to separate sites on fibrillar collagens. J. Biol. Chem. 268, 27307–27312. 44. Schönherr, E., Witsch-Prehm, P., Harrach, B., Robenek, H., Rauterberg, J., and Kresse, H. (1995). Interaction of biglycan with type I collagen. J. Biol. Chem. 270, 2776–2783. 45. Fraser, R. D., and MacRae, T. P. (1979). The crystalline structure of collagen fibrils in tendon. J. Mol. Biol. 127, 129–133. 46. Fraser, R. D., MacRae, T. P., Miller, A., and Suzuki, E. (1983). Molecular conformation and packing in collagen fibrils. J. Mol. Biol. 167, 497–521. 47. Brodsky, B., and Eikenberry, E. F. (1985). Supramolecular collagen assemblies. In “Biology, Chemistry, and Pathology of Collagen” (R. Fleischmajer, B. R. Olsen, and K. Kühn, eds.), Vol. 460, pp. 73–84. The New York Academy of Sciences, New York. 48. Hulmes, D. J. S., and Miller, A. (1979). Quasi-hexagonal packing in collagen fibrils. Nature 282, 878–880. 49. Wess, T. J., Hammersley, A. P., Wess, L., and Miller, A. (1998). Molecular packing of type I collagen in tendon. J. Mol. Biol. 275, 255–267.

50. Hulmes, D. J. S., Holmes, D. F., and Cummings, C. (1985). Crystalline regions in collagen fibrils. J. Mol. Biol. 184, 473–477. 51. Brodsky, B., and Eikenberry, E. F. (1982). Characterization of fibrous forms of collagen. Methods of Enzymology 82, 127–174. 52. Katz, E. P., Wachtel, E. J., and Maroudas, A. (1986). Extrafibrillar proteoglycans osmotically regulate the molecular packing of collagen in cartilage. Biochim. Biophys. Acta 882, 136–139. 53. Tanaka, S., Avigad, G., Eikenberry, E. F., and Brodsky, B. (1988). Isolation and partial characterization of collagen chains dimerized by sugar-derived cross-links. J. Biol. Chem. 263, 17650–17657. 54. Grynpas, M. D., Eyre, D. R., and Kirschner, D. A. (1980). Collagen type II differs from type I in native molecular packing. Biochim. Biophys. Acta 626, 346–355. 55. Wachtel, E., Maroudas, A., and Schneiderman, R. (1995). Age-related changes in collagen packing of human articular cartilage. Biochim. Biophys. Acta 1243, 239–243. 56. Price, R. I., Lees, S., and Kirschner, D. A. (1997). X-ray diffraction analysis of tendon collagen at ambient and cryogenic temperatures: role of hydration. Int. J. Biol. Macromol. 20, 23–33. 57. Tanaka, S., Avigad, G., Brodsky, B., and Eikenberry, E. F. (1988). Glycation induces expansion of the molecular packing of collagen. J. Mol. Biol. 203, 495–505. 58. Bank, R. A., Bayliss, M. T., Lafeber, F. P. J. G., Maroudas, A., and Tekoppele, J. M. (1998). Ageing and zonal variation in post-translational modification of collagen in normal human articular cartilage. The age-related increase in non-enzymatic glycation affects biomechanical properties of cartilage. Biochem. J. 330, 345–351. 59. Vaughan, L., Mendler, M., Huber, S., Bruckner, P., Winterhalter, K. H., Irwin, M. I., and Mayne, R. (1988). D-periodic distribution of collagen IX along cartilage fibrils. J. Cell Biol. 106, 991–997. 60. Mendler, M., Eich-Bender, S. G., Vaughan, L., Winterhalter, K. H., and Bruckner, P. (1989). Cartilage contains mixed fibrils of collagen types II, IX, and XI. J. Cell Biol. 108, 191–197. 61. Keene, D. R., Oxford, J. T., and Morris, N. P. (1995). Ultrastructural localization of collagen types II, IX, and XI in the growth plate of human rib and fetal bovine epiphyseal cartilage: Type XI collagen is restricted to thin fibrils. J. Histochem. Cytochem. 43, 967–979. 62. Wu, J. J., Woods, P. E., and Eyre, D. R. (1992). Identification of crosslinking sites in bovine cartilage type IX collagen reveals an antiparallel type II-type IX molecular relationship and type IX to type IX bonding. J. Biol. Chem. 267, 23007–23014. 63. Wu, J.-J., and Eyre, D. R. (1995). Structural analysis of cross-linking domains in cartilage type XI collagen. Insights on polymeric assembly. J. Biol. Chem. 270, 18865–18870. 64. Blaschke, U. K., Eikenberry, E. F., Hulmes, D. J. S., Galla, H. J., and Bruckner, P. (2000). Collagen XI nucleates assembly and limits lateral growth of cartilage fibrils. J. Biol. Chem. 275, 10370–10378. 65. Lee, S. L., and Piez, K. A. (1983). Type II collagen from lathyritic rat chondrosarcoma: Preparation and in vitro fibril formation. Collagen Relat. Res. 3, 89–103. 66. Bruckner, P., and van der Rest, M. (1994). Structure and function of cartilage collagens. Microsc. Res. Tech. 28, 378–384. 67. Gelman, R. A., Williams, B. R., and Piez, K. A. (1979). Collagen fibril formation. Evidence for a multistep process. J. Biol. Chem. 254, 180–186. 68. Hansen, U., and Bruckner, P. (2003). Macromolecular specificity of collagen fibrillogenesis: Fibrils of collagens I and XI contain a heterotypic alloyed core and a collagen I sheath. J. Biol. Chem. 278, 37352–37359. 69. Li, Y. F., and Olsen, B. R. (1997). Murine models of human genetic skeletal disorders. Matrix Biol. 16, 49–52. 70. Aszódi, A., Pfeifer, A., Wendel, M., Hiripi, L., and Fässler, R. (1998). Mouse models for extracellular matrix diseases. J. Mol. Med. 76, 238–252. 71. Chan, D., Cole, W. G., Chow, C. W., Mundlos, S., and Bateman, J. F. (1995). A COL2A1 mutation in achondrogenesis type II results in the

420

72.

73.

74.

75.

76.

77.

78.

79.

80.

81.

replacement of type II collagen by type I and III collagens in cartilage. J. Biol. Chem. 270, 1747–1753. Mundlos, S., Chan, D., McGill, J., and Bateman, J. F. (1995). An a1(II) Gly913 to Cys substitution prevents the matrix incorporation of type II collagen which is replaced with type I and II collagens in cartilage from a patient with hypochondrogenesis. Am. J. Med. Genet. 63, 129–136. Bruckner, P., and Eikenberry, E. F. (1984). Formation of the triple helix of type I procollagen in cellulo. Temperature-dependent kinetics support a model based on cis-trans isomerization of peptide bonds. Eur. J. Biochem. 140, 391–395. Li, S. W., Prockop, D. J., Helminen, H., Fässler, R., Lapveteläinen, T., Kiraly, K., Peltarri, A., Arokoski, J., Lui, H., Arita, M., and Khillan, J. (1995). Transgenic mice with targeted inactivation of the COL2A1 gene for collagen II develop a skeleton with membranous and periosteal bone but no endochondral bone. Genes Dev. 9, 2821–2830. Aszódi, A., Chan, D., Hunziker, E. B., Bateman, J. F., and Fässler, R. (1998). Collagen II is essential for the removal of the notochord and the formation of intervertebral discs. J. Cell Biol. 143, 1399–1412. Sahlman, J., Inkinen, R., Hirvonen, T., Lammi, M. J., Lammi, P. E., Nieminen, J., Lapveteläinen, T., Prockop, D. J., Arita, M., Li, S. W., Hyttinen, M. M., Helminen, H. J., and Puustjärvi, K. (2001). Premature vertebral endplate ossification and mild disc degeneration in mice after inactivation of one allele belonging to the Col2a1 gene for type II collagen. Spine 26, 2558–2565. Hyttinen, M. M., Toyras, J., Lapveteläinen, T., Lindblom, J., Prockop, D. J., Li, S. W., Arita, M., Jurvelin, J. S., and Helminen, H. J. (2001). Inactivation of one allele of the type II collagen gene alters the collagen network in murine articular cartilage and makes cartilage softer. Ann. Rheum. Dis. 60, 262–268. Rodriguez, R. R., Seegmiller, R. E., Stark, M. R., and Bridgewater, L. C. (2004). A type XI collagen mutation leads to increased degradation of type II collagen in articular cartilage. Osteoarthritis Cart. 12, 314–320. Müller-Glauser, W., Humbel, B., Glatt, M., Sträuli, P., Winterhalter, K. H., and Bruckner, P. (1986). On the role of type IX collagen in the extracellular matrix of cartilage: Type IX collagen is localized to intersections of collagen fibrils. J. Cell Biol. 102, 1931–1939. Eyre, D. R., Pietka, T., Weis, M. A., and Wu, J.-J. (2004). Covalent cross-linking of the NC1 domain of collagen type IX to collagen type II in cartilage. J. Biol. Chem. 279, 2568–2574. Hagg, R., Hedbom, E., Möllers, U., Aszódi, A., Fässler, R., and Bruckner, P. (1997). Absence of the α1(IX) chain leads to a functional knock-out of the entire collagen IX protein in mice. J. Biol. Chem. 272, 20650–20654.

PETER BRUCKNER 82. Kassner, A., Tiedemann, K., Notbohm, H., Ludwig, T., Mörgelin, M., Reinhardt, D. P., Chu, M.-L., Bruckner, P., and Grässel, S. (2004). Molecular structure and interaction of recombinant human type XVI collagen. J. Mol. Biol. 339, 835–853. 83. Koch, M., Schulze, J., Hansen, U., Ashwodt, T., Keene, D. R., Brunken, W. J., Burgeson, R. E., Bruckner, P., and BrucknerTuderman, L. (2004). A novel marker of tissue junctions, collagen XXII. J. Biol. Chem. 279, 22514–22521. 84. Muragaki, Y., Mariman, E. C. M., Van Beersum, S. E. C., Perälä, M., Van Mourik, J. B. A., Warman, M. L., Olsen, B. R., and Hamel, B. C. J. (1996). A mutation in the gene encoding the α2 chain of the fibrilassociated collagen IX, COL9A2, causes multiple epiphyseal dysplasia (EDM2). Nat. Genet. 12, 103–105. 85. Parsons, J. R., and Black, J. (1977). The viscoelastic shear behavior of normal rabbit articular cartilage. J. Biomech. 10, 21–29. 86. Setton, L. A., Zhu, W., and Mow, V. C. (1993). The biphasic poroviscoelastic behavior of articular cartilage: role of the surface zone in governing the compressive behavior. J. Biomech. 26, 581–592. 87. Basser, P. J., Schneiderman, R., Bank, R. A., Wachtel, E., and Maroudas, A. (1998). Mechanical properties of the collagen network in human articular cartilage as measured by osmotic stress technique. Arch. Biochem. Biophys. 351, 207–219. 88. Aspden, R. M., and Hukins, D. W. L. (1990). Stress in collagen fibrils of articular cartilage calculated from their measured orientations. Matrix 9, 486–488. 89. Maroudas, A., Bayliss, M. T., Uchitel-Kaushansky, N., Schneiderman, R., and Gilav, E. (1998). Aggrecan turnover in human articular cartilage: Use of aspartic acid racemization as a marker of molecular age. Arch. Biochem. Biophys. 350, 61–71. 90. Birk, D. E., Fitch, J. M., Babiarz, J. P., Doane, K. J., and Linsenmayer, T. F. (1990). Collagen fibrillogenesis in vitro. Interaction of type I and type V collagen regulates fibril diameter. J. Cell Sci. 95, 649–657. 91. Rousseau, J. C., Farjanel, J., Boutillon, M. M., Hartmann, D. J., van der Rest, M., and Moradi-Améli, M. (1996). Processing of type XI collagen. Processing of the matrix forms of the α1(XI) chain. J. Biol. Chem. 271, 23743–23748. 92. Chapman, J. A. (1989). The regulation of size and form in the assembly of collagen fibrils in vivo. Biopolymers 28, 1367–1382. Published erratum: Biopolymers 28, 2201–2205. 93. Hulmes, D. J. S. (1983). A possible mechanism for the regulation of collagen fibril diameter in vivo. Collagen Relat. Res. 3, 317–321.

Chapter 25

Products of Cartilage Metabolism Daniel-Henri Manicourt

Jean-Pierre Devogelaer

Laboratoire de Chimie Physiologique (Metabolic Research Group, Connective Tissue Section), Christian de Duve Institute of Cellular Pathology and Department of Rheumatology, Saint Luc University Hospital, Université Catholique de Louvain, 1200 Brussels, Belgium Department of Rheumatology, Saint Luc University Hospital, Université Catholique de Louvain, 1200 Brussels, Belgium

Eugene J.-M. A. Thonar

Departments of Biochemistry, Internal Medicine and Orthopedic Surgery, Rush Medical College, Rush-Presbyterian-St. Luke’s Medical Center, Chicago, Illinois, 60612 USA

I. Introduction II. The Chondrocyte and its Extracellular Matrix III. Products of Collagen Metabolism IV. Products of Aggrecan Metabolism V. Products of the Metabolism of other Proteoglycans

VI. Products of the Metabolism of Link Protein and Hyaluronan VII. Other Products of Chondrocyte Metabolism VIII. Concluding Statement References

I. INTRODUCTION

worth pointing out that articular cartilage in diarthrodial (synovial) joints makes up only approximately 10% of the total cartilage mass in the body of an adult dog [1]. Joint diseases and related conditions involving abnormalities in the metabolism of cartilaginous tissues cause widespread disability. While articular cartilage failure was first regarded as a passive degenerative process, modern studies have provided a large body of evidence that this is not so [2]. In osteoarthritis (OA), for example, the destruction of the articular surface is now thought to result from an imbalance between dynamic anabolic and catabolic processes that normally proceed in a harmonized and strictly regulated manner [2]. Current research on cartilage includes attempts to define the mechanisms that regulate

Cartilage comprises approximately 1% of the dry body weight of an adult dog [1]. Distributed throughout the body, this hard but resilient tissue performs a variety of functions. During development, cartilage acts as a growing scaffold for limb elongation. Articular cartilage, which covers the ends of bones in diarthrodial joints, as well as the softer and more compressible nucleus pulposus of the intervertebral disk, allows the joints in the rigid skeleton to articulate smoothly. Cartilage is the supporting structure of the respiratory tract (nose, larynx, and trachea) and, as costal cartilage, contributes to the protection of the lungs, heart, liver, and spleen from external physical stresses. It is Dynamics of Bone and Cartilage Metabolism

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the expression of cartilage-specific genes, to shed light upon the structural and functional properties of individual matrix components and to better understand how these matrix-building blocks are turned over in both health and disease. This chapter will primarily discuss the major products of articular cartilage metabolism with special emphasis upon those that are relevant as body fluid markers. The structure and supramolecular organization of major cartilage components as well as the main regulatory pathways of chondrocyte metabolism have been detailed in the preceding chapters of this volume. The following section provides a brief review of the structure–function relationships of the major matrix-building blocks.

II. THE CHONDROCYTE AND ITS EXTRACELLULAR MATRIX The prime function of articular cartilage is physical, with water, ions and anionic aggrecan molecules within the collagenous meshwork all playing key roles in endowing the tissue with its load-bearing properties. The collagenous meshwork rich in type II collagen molecules gives the tissue tensile strength and hinders the expansion of the viscoelastic under-hydrated aggrecan molecules that provide compressive stiffness. The highly sulfated aggrecan molecules interact noncovalently with a single strand of hyaluronan (HA) and link protein (LP) molecules to form supramolecular aggregates of very large size that become firmly entrapped within the collagenous meshwork (see Chapters 2 and 5). The resulting high fixed negative charge density, balanced by mobile counter ions, gives rise to high osmotic pressures (up to 2–3 atmospheres), which are counteracted by the constant tension that is developed within the meshwork of cross-linked collagenous fibrils [3]. Cartilage is thus made stiff by being swollen with water. As aggrecan molecules oppose the fluid loss and the redistribution of water within the tissue, cartilage shows resilience upon loading, with the tissue rapidly recovering its elasticity when the load is removed. During cycle loading, the fluid flowing through the fine pores of the tissue not only dissipates energy and absorbs shocks but also participates in joint lubrication, contributes to chondrocyte nutrition, and acts as a mechanical signal transducer that regulates chondrocyte synthetic/catabolic activities [4]. It is obvious, therefore, that the structural integrity of the collagenous meshwork and aggrecan molecules is critically important for the maintenance of the biomechanical properties of the tissue. The extracellular matrix (ECM) of cartilage contains many additional components that contribute to the cohesiveness of the matrix and to the regulation of

chondrocyte function. For example, types IX and XI collagen (Chapters 1 and 2) are essential players in the organization and functions of the major collagenous fibrils, whereas type VI collagen forms distinct microfibrils that appear concentrated in the capsular matrix surrounding individual chondrocytes or groups of chondrocytes; this protective structure, termed the chondron, dampens the osmotic and physicomechanical changes induced by joint loading [5, 6]. By interacting with growth factors and other macromolecules of the ECM, including collagens and fibronectin (Chapter 5), small nonaggregating proteoglycans (PGs) (including decorin, biglycan, perlecan, lumican, glypican, and fibromodulin) contribute to the regulation of chondrocyte function and to the maintenance of the supramolecular organization of the matrix, a contention strengthened by the observation that mutations in the gene encoding for several of these molecules lead to chondrodysplasias [7, 8]. An ill-defined number of noncollagenous proteins (Chapter 4) also participate in the complex interactions that together form a network in which chondrocytes and most molecules of the ECM are involved. The relevance of some of these proteins is illustrated by their involvement in genetic disorders [7]. Cartilage matrix-building blocks are synthesized, organized, and maintained by a sparse population of chondrocytes. These cells are protected from the potentially damaging forces of mechanical function by the ECM they produce. As the properties of cartilage are critically dependent upon the structure and integrity of the ECM, normal turnover is a conservative process in which the rate of degradation of each molecular species does not exceed the rate at which it is replaced by newly synthesized products. The degradation of matrix molecules involves primarily proteinases (Chapter 10) but probably also free radicals secreted by chondrocytes. In disease states, the rate of degradation of matrix molecules often exceeds the rate of synthesis and, as a consequence, the tissue becomes thin and mechanically weak. Because cartilage is avascular, chondrocyte nutrition in a diarthrodial joint has to rely upon the diffusion of nutrients from the synovial fluid that bathes the tissues and links together the different components within the joint. The oxygen tension in cartilage may be as low as 1–3%, compared to 24% in the normal atmosphere [9]; this low oxygen tension appears to be beneficial for maintenance of the chondrocyte-specific phenotype in vitro [10]. This remarkable adaptation of chondrocytes to the hypoxic environment prevailing in the cartilage tissue might be related to their relatively high constitutive expression levels of the transcriptional complex hypoxia-inducible factor-1, which regulates genes encoding growth factors, glycolytic enzymes, and glucose transporters [11, 12].

Chapter 25 Products of Cartilage Metabolism

While the majority of animal cells derive their energy by using oxygen for mitochondrial oxidative phosphorylation, mammalian articular chondrocytes use oxygen to stimulate the pathway of glycolysis, a sequence of reaction that produces lactate, does not consume oxygen and is the principal source of ATP for metabolic processes, including the synthesis of the extracellular matrix [13, 14]. To maintain intracellular pH, and hence glycolysis, the large quantities of lactate− and H+ ions produced by the glycolytic pathway are extruded into the extracellular environment via the monocarboxylate transporters and other H+ active transporters, such as the sodium–hydrogen exchanger [15, 16]. By inhibiting glycolysis, iodoacetate reduces ATP levels and lactate production markedly, and, in so doing, leads to degenerative changes in articular cartilage. Since aggrecan molecules are highly sulfated, the Donnan equilibrium ensures that chondrocytes in vivo also survive in a relatively acidic environment and at a higher osmolarity than other cell types (350–450 mOsm, compared with 280 mOsm) [17]. Osmolarity is a powerful regulator of matrix turnover. In vitro, chondrocytes respond readily to alterations in their ionic environment with maximal rates of synthesis of aggrecan and proteins at osmolalities close to those experienced in situ [17]. When cells are faced with osmotic challenges, the activation of signaling pathways (SAPK2, ERK, and others) enhances the activity of a variety of membrane transport proteins, including ionic channels and carriers as well as amino acid transport systems, which all contribute to restore the intracellular ionic milieu and, hence, cell function [18, 19]. Changes in the rate of synthesis of matrix macromolecules also occur in vivo when PG concentrations are altered during fluid loss under load and when cartilage swells because of a decrease in the tensile stiffness of the collagenous meshwork, as observed in the initial stages of OA [19]. Clinical observations, as well as in vitro and in vivo studies, point out that cartilage loading markedly influences the biosynthetic activity of chondrocytes, which then can adapt the composition and organization of the ECM in response to changes in functional demand. In postnatal cartilage, the high weight-bearing regions have a higher aggrecan content and a stiffer collagenous meshwork than low weight-bearing regions [3]. This is in contrast to neonatal cartilage which does not exhibit any topographical variation in tissue composition [20, 21]. Cartilage explants are sensitive to applied loads. Fluctuating loads stimulate matrix production, the extent of the response being influenced by the frequency, amplitude, and wave forms, whereas static pressure causes a reduction in the rate of synthesis of both aggrecan and LP molecules, without affecting HA synthesis [22–25].

423 Joint immobilization also causes a marked loss of PGs from articular cartilage [26]. This depletion, reflecting both a decline in the rate of PG synthesis and an increase in PG catabolism, is, however, completely reversible when the joints are remobilized [27]. The absence of motion appears more critical than the lack of loading because even small amounts of motion of the unloaded joint reduces the severity of the PG depletion, suppresses the signal transduction pathways of pro-inflammatory/catabolic mediators, and stimulates anabolic pathways; a finding that justifies the clinical use of continuous passive motion [28]. On the other hand, strenuous joint loading can cause disruption of the collagenous meshwork and a chondrocyte-mediated loss of matrix PGs as well as cell necrosis and apoptosis, which can all lead to osteoarthritic cartilage changes [29–31]. Further, after an injurious compression, the remaining viable chondrocytes are no longer able to respond to the stimulatory effects of moderate dynamic compression as seen in normal cartilage [32]. How joint loading affects the biosynthetic activity of chondrocytes remains poorly understood. There is, however, evidence that the intracellular composition, the rate of glycolysis, and the pattern of gene expression are all influenced by changes in chondrocyte membrane permeability, which are generated upon joint loading by changes in pericellular osmolarity and pressure, as well as by membrane stretch and shear stresses [33, 34]. Further, by deforming the intracellular organelles and nucleus, loads might change the rate of synthesis, secretion, and/or quality of matrix molecules [35]. It is also clear that, by deforming the ECM and producing local and temporal changes in gradient pressure, loads initiate intratissue fluid flows, which not only generate electrical streaming potentials and currents but also affect the transport of soluble factors and nutrients and might liberate locally basic fibroblast growth factor from an extracellular heparan sulfate-bound pool [35–38]. All these mechanical, chemical, and electrical signals are sensed by the chondrocyte and are thought to be mediated, at least in part, by integrins (Chapter 8) and extracellular signal-regulated kinase (ERK), a member of the ubiquitous family of mitogen-activated kinases (MAPs), which contributes to the transmission of these stimuli from outside the cell to the nucleus and influences a variety of cell processes, including transcriptional regulation [39]. The effects of mechanical stresses are also modulated by the quality of the matrix and more particularly its aggrecan content, which regulates swelling pressure, and the properties of the collagenous meshwork, which influences the degree of deformation. Thus, under the same load, the extent of cartilage deformation, the increase of fluid flow, the hydrostatic pressure and the change in fluid content can be very different in different areas of the same joint,

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in young relative to old cartilage, or in normal relative to arthritic tissue. The biosynthetic response of chondrocytes to mechanical stress is also affected by their origin [40]. Indeed, with distance from the articular surface, chondrocytes exhibit heterogeneity with respect to shape, surface receptors, and metabolic activities [41]. In culture, chondrocytes from the deeper layers of articular cartilage exhibit a faster rate of PG synthesis and a slower rate of PG turnover than cells from the superficial layers, a finding that is consistent with the observation that net matrix accumulation is more pronounced around deep but superficial chondrocytes. On the other hand, chondrocytes from the superficial layers produce greater amounts of nitric oxide when stimulated with cytokines and are even more sensitive to the catabolic effect of interleukin-1 [42]. The precise reasons for this type of heterogeneity among chondrocytes are not obvious. However, it is clear that the shape and magnitude of the biomechanical stresses are unevenly distributed throughout

Figure 1

the cartilage matrix. It also is clear that chondrocytes in different layers are subjected to different concentrations of nutrients and metabolites.

III. PRODUCTS OF COLLAGEN METABOLISM As stated in Chapter 1, type II collagen is synthesized as a large precursor molecule, termed procollagen, which exhibits a globular extension propeptide in both its N- and C-terminal ends (Fig. 1) [43]. Depending upon alternative splicing of exon 2, the N-propeptide either contains (form IIA) or does not contain (form IIB) a 69 aminoacid cysteine-rich domain [44]. Although the function of the domain encoded by exon 2 remains poorly understood, there is clear evidence that normal adult articular cartilage expresses high levels of type IIB procollagen, whereas type IIA procollagen is expressed during embryogenesis,

Type II collagen fibrils are made of tropocollagen molecules interconnected by cross-links between the nonhelical telopeptide regions of individual tropocollagen molecules and the helical region of adjacent molecules. Each tropocollagen molecule is composed of three identical α chains. After a first cleavage by any one of the three mammalian collagenases (MMP-1, MMP-8, and MMP-13), the cleaved triple helix unwinds at physiological temperature and exposes hidden epitopes on α chains that are not recognized on the native triple helix. Epitopes on nonhelical telopeptides and conformational triple helical dependent epitopes are also indicated. Adapted from reference [43].

Chapter 25 Products of Cartilage Metabolism

at the onset of cartilage hypertrophy and in human OA cartilage as well [45–47]. Soon after its release extracellularly, the type II procollagen molecule is processed by specific proteinases which remove its two globular propeptides: the disintegrin and metalloproteinase with thrombospondin-like motifs (ADAMTS)-3 cleaves the N-propeptide at A181 ↓ Q182 [48] whereas bone morphogenic protein-1 (BMP-1) cleaves the C-propeptide at A1241 ↓ D1242 [49]. This proteolytic cleavage allows the resulting tropocollagen molecule, consisting of a long compact triple-helical region with nontriplehelical short telopeptides in both N- and C-terminal ends, to become incorporated into a fibril (Fig. 1). Because they cannot be detected in cell layers and culture media of chondrocytes, the free N-propeptide molecules of type II collagen are thought to be quickly metabolized [50]. In contrast, the free C-propeptide of the pro [α(II)] chain (CPII), also referred to as chondrocalcin, is readily detected in the pericellular matrix of cartilage explants [47, 51]. In healthy adult articular cartilage, the concentrations of CPII are low, appear relatively constant throughout the tissue and do not change from 30 to 75 years of age whereas, in OA cartilage, CPII levels are usually elevated in the middle and deeper zones of the matrix but not in the superficial more damaged zone, therefore suggesting that the less degenerated cartilage is capable of mounting a reparative response characterized by increased synthesis of type II procollagen. Most CPII molecules are thought to diffuse out of the cartilage matrix after they are produced and their concentration in body fluids can be measured by specific immunoassays [51, 52]. Support for the contention that the level of this body fluid marker provides a measure of the rate of type II collagen synthesis in cartilage comes from the observation that the rate of synthesis of type II collagen in articular cartilage is directly proportional to the tissue content of C-propeptide: the latter was found to have a halflife of 14–20 hours [51]. This contention is further strengthened by the observation that, during longitudinal bone growth, the enhanced rate of synthesis of type II collagen in the growth plate is associated with elevated serum levels of CPII, which then drop as growth ceases [53]. Synovial fluid levels of CPII are found in higher levels in traumatic and primary osteoarthritis (OA) than in wellestablished rheumatoid arthritis (RA) or infectious arthritis [54–57], confirming the widely held view that the degradative events in OA are balanced, at least during the early stages, by an up-regulation of the biosynthetic processes. In contrast, the synovial fluid levels of CPII are low in early RA suggesting that, in the early stages of this disease, down-regulation of the rate of type II collagen synthesis might contribute to joint destruction [57]. Interestingly, the

425 enhanced levels of CPII found in the articular cartilage and synovial fluid from both RA and OA joints are reflected by an increase in the serum concentrations of CPII in RA patients, but not in OA patients who instead exhibit a significant reduction in their circulating levels of type II collagen propeptides [51, 52, 58]. It is possible that in OA patients the altered serum levels of CPII reflect a reduction of type II collagen synthesis in cartilages not affected by OA. Once tropocollagen molecules have spontaneously aggregated into fibrils (Fig. 1), complex covalent crosslinks form within and between the collagen molecules to enhance the cohesiveness of the collagenous meshwork (Chapter 2). The only step of the crosslinking process that is known to be cell-controlled is the oxidative deamination (by lysyl oxidase) of the ε-amino group in telopeptidyl lysine and hydroxylysine residues to form the aldehydes, termed allysine and hydroxyallysine, respectively [59]. The hydroxyallysine-derived cross-links predominate in cartilage and lead almost exclusively to the formation of the stable, nonreducible and trivalent 3-hydroxypyridinium cross-link termed hydroxylysyl pyridinoline (also termed pyridinoline) whose concentrations change little during adulthood. On the other hand, with increasing age, the nonenzymatic glycation of lysine and arginine residues leads to the formation and accumulation of advanced glycation end products (AGEs) and crosslinks, such as pentosidine, methylglyoxallysine dimer, and threosidine [60, 61]. AGE crosslinking of the collagen network is believed to contribute to the age-related increase in stiffness of the collagen network, which, by decreasing the resistance of this network to mechanical damage, might predispose to the development of OA. Since type II collagen is the most abundant component of the collagenous meshwork in cartilage (90–95% of the total collagen content), the proteolytic degradation of this fibrillar collagen is likely an important rate-limiting step in tissue remodeling both in health and disease. Proteinases can contribute to the extracellular degradation of type II collagen in several ways. First, by cleaving the telopeptides of collagen molecules, proteinases separate the intact triple-helical domain from the telopeptide crosslinks and thus depolymerize the collagen fibrils without actually damaging the triple helix: the latter can be recognized by monoclonal antibodies that only react with the intact native triple-helical molecule [62]. Second, proteinases can also cleave the triple-helical domain of the native and fully wound collagen molecules. At physiologic temperature the cleaved triple helix unwinds (Fig. 1) to yield denatured collagen or gelatin and exposes neoepitopes that are normally hidden and not recognized in the native triple helix but are recognizable on cyanogen bromide (CB)

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peptides of type II collagen [62]. Third, proteinases can also degrade gelatin and/or activate zymogens which hereby acquire the ability to cleave collagen molecules. Although neutrophil elastase and stromelysin-1 or metalloproteinase-3 (MMP-3) are the only mammalian proteinases known to cleave telopeptides in vitro [63, 64], studies conducted in cartilage explants [65] suggest that cleavage of the N-telopeptide by MMP-3 is unlikely to be a major mechanism in the chondrocyte-mediated degradation of collagen (see also Chapter 11). This notwithstanding, MMP-3 might play a pivotal role in collagen damage by activating collagenase-1 (MMP-1), collagenase-3 (MMP-13), and gelatinase B (MMP-9) [66, 67]. It also

should be noted that neutrophil elastase also can activate MMP-3 and MMP-8 [66]. Thus far, MMP-1, MMP-8 (neutrophil collagenase), MMP-13, and MMP-14 (membrane type I MMP or MT1MMP), which all are synthesized by chondrocytes, are the major mammalian proteinases known to be capable of hydrolyzing the intrahelical domain of native fibrillar collagens at neutral pH [68]. These collagenases cleave initially all three α chains of type II collagen at the classic G775 ↓ L776 bond, three-quarters of the way from the N-terminal end (Fig. 1). This produces denatured α chain fragments having approximately 3/4 length (TCA fragment) and 1/4 length (TCB fragment) of the native molecule.

Table I. Neoepitopes of Type II Collagen Cleavage. Summary of the Type II Collagen Neoepitopes and their Molecular Origin in the Triple Helical and C-telopeptide Domains of Type II Collagen Alpha Chains. 1. Schematic of type II collagen alpha chain cleavage by collagenases and matrix metalloproteinases First collagenase cleavage in triple helical domain -------GPPOHGPQG775 Ø LAGQRGICG-----Second collagenase cleavage in triple helical domain LAG778 Ø QRGICG-----Matrix metalloproteinase cleavage in C-telopeptide domain ------PREKGPDP Ø LQYMRA-----2. Neoepitopes located in the triple helical domain Identification

Immunizing antigen

Characteristics

COL2-1/4N1

LAGQRG

COL2-1/4N2

QRGIVG (LP)

COL2-3/4Cshort C1,2C

GPPOHGPQG GPPGPQG

COL2-3/4Clong C2C

GGEGPPOHGPQG

COL2-3/4m

APOHGEDGRPOHGP

COLL2-1 COLL2-1-NO2

HRGYPGLDG HRGY(NO2)PGLDG

Located at the N-terminus of the C-terminal α chain fragment after collagenase first cleavage of the triple helical domain of type II tropocollagen molecules This epitope is rapidly destroyed by secondary collagenase cleavage Located at the N-terminus of the C-terminal α chain fragment after collagenase second cleavage of the triple helical domain of type II tropocollagen molecules Located at the C-terminus of the N-terminal α chain fragment after collagenase first cleavage of the triple helical domain of type II tropocollagen molecules. Detected in body fluids Also present in a collagenase cleavage fragment generated from type I collagen Located at the C-terminus of the N-terminal α chain fragment after collagenase first cleavage of the triple helical domain of type II tropocollagen molecules. Detected in body fluids Digested by alpha chymotrypsin Residues 511–523 located in the N-portion of the N-terminal α chain fragment after collagenase first cleavage of the triple helical domain of type II tropocollagen molecules This neoepitope signs collagen denaturation: it is resistant to cleavage by alpha-chymotrypsin which is used to solubilize denatured collagen The sequence is also present in alpha 3 (XI) chain Residues 108–116 located in the N-terminal portion of the triple helical domain of type II tropocollagen molecules Residue Y may be nitrated by oxygen-derived free radicals Detected in body fluids The sequence is also found in type XI collagen

3. C-telopeptide neoepitope Identification

Immunizing antigen

Characteristics

CTX-II CartiLaps COL2-CTx

EKGPDP

Sequence found exclusively in human type II collagen C-telopeptide domain K indicate the lysine residue involved in intramolecular cross-linking Generated by matrix metalloproteinases. Detected in urine

Chapter 25 Products of Cartilage Metabolism

Immuno-histochemical staining and immunoassays conducted with antibodies that react specifically with denatured but not with native type II α chains (neoepitope COL2-3/4m; Table I), have established that adult normal human cartilage from large joints such as the hip or the knee, but not the ankle, may contain up to 3% of its total collagen in the denatured form: after 35 years of age, this can be detected around chondrocytes at and near the articular surface, and, with increasing age this damage may extend progressively into the cartilage [69–71]. Although denaturation of type II collagen is enhanced in cartilage obtained from joints with OA and RA, the topographical distribution of the collagen cleavage differs markedly between the two arthritides [72]. In OA cartilage, damage to collagen is first detected at and near the articular surface in territorial and interterritorial sites around chondrocytes and from there on extends into deeper cartilage layers, whereas in RA, denaturation of type II collagen is much more pronounced in the territorial matrix surrounding chondrocytes of the deeper layers [69, 70, 72]. These findings strongly suggest that cytokines derived from subchondral bone might enhance chondrocyte-derived degradation of the collagen meshwork in RA. After cleavage of the α chains by any one of the four collagenases (MMP-1, MMP-8, MMP-13, and MMP-14),

Figure 2 The three mammalian collagenases (MMP-1, MMP-8, and MMP-13) first cleave the triple helical domain of type II procollagen molecules at a site located three-quarters of the way from the N-terminus of the molecule. This cleavage unwinds the triple helix and exposes two neoepitopes, i.e. the C-terminus of the N-terminal α chain fragment, termed COL2-3/4 C, and the N-terminus of the C-terminal α chain fragment, termed COL2-1/4 N1. The N-terminal neoepitope COL2-1/4 N1 is rapidly destroyed by secondary cleavage sites of collagenases which also release the C-terminal neoepitope COL2-3/4 C that diffuses then out of the matrix to reach biological fluids where it can be detected. The amino acid sequence of the neoepitopes is also given. Adapted from reference [73].

427 unwinding of the triple helix exposes neoepitopes at the C-terminus of the TCA fragments and at the N-terminus of the TCB fragments (Fig. 2). Immunoassays conducted with antibodies reacting specifically with the C-terminal (COL2-3/4) and N-terminal (COL2-1/4 N1) neoepitopes (Table I) have demonstrated that, in both normal and OA cartilages, the content of COL2-3/4 neoepitope correlates strongly with the tissue content of denatured type II collagen [73]. Further, in OA cartilage, the enhanced denaturation of type II collagen becomes inversely proportional to the cartilage tissue content of this type of collagen [68]. These findings support the contention that collagenase activity is responsible for the cleavage of type II collagen in human articular cartilage and that any effective therapeutic intervention in OA must encompass a potent control of collagen degradation. When compared to other collagenases, MMP-13 hydrolyzes type II collagen more efficiently and also has a broader substrate specificity, cleaving type I, II, III, IV, IX, X, and XIV collagen, gelatin, fibronectin, and tenascin [74]. Studies of cartilage explants exposed to synthetic inhibitors of collagenases [75, 76] as well as the semi-quantification of MMP RNA transcripts [77], provide strong evidence that MMP-13 might be the most important collagenase involved in the degradation of type II collagen both in normal and OA cartilage, a contention further strengthened by the observation that transgenic mice over-expressing active MMP-13 exhibit joint pathology that mimics OA [78]. Further, during growth plate development, the synthesis and activity of MMP-13, but not MMP-1, by hypertrophic chondrocytes is associated with increased degradation of type II collagen [79]. On the other hand, proteolysis of newly synthesized collagen molecules might involve MMP-1 rather than MMP-13 [75], whereas other members of the MMP family such as MMP-2 (gelatinase A), MMP-3, and MMP-9 are very efficient at cleaving denatured α chains and at removing damaged collagen from the extracellular matrix [80]. It is thought that this subsequent clearance of unwound α chains may be a necessary event to facilitate repair but it is possible that the gelatinolytic enzymes cause additional damage to the cartilage matrix. Another enzyme likely to contribute to the turnover of type II collagen both in health and disease is cathepsin K. OA chondrocytes up-regulate the expression of this cysteine protease, which not only can cleave within the N-terminal portion of the helical domain of native type II collagen at near neutral pH, but also exhibits a high gelatinolytic activity in the pH range 4.0–7.0 [81]. Cathepsin K might be involved in the retinoic acid-induced type II collagen degradation which has been shown to be mainly independent of MMP activity in cartilage explants [82, 83].

428 The two epitopes present at the carboxy-terminus of the TCA 3/4 fragment (Table I and Fig. 2) and termed C2C (known previously as COL2-3/4Clong) and C1,2C (known previously as COL2-3/4Cshort) seem to be a better marker of collagen cleavage than the COL2-1/4 N1 neoepitope, since the N-terminal neoepitope is readily destroyed by a secondary collagenase-mediated cleavage. In contrast, as collagenases produce further cleavage of the TCA fragment, albeit at a slower rate, the C2C and C1,2C neoepitopes are released and can be readily detected in the conditioned medium of cartilage explants and in body fluids as well [62, 68]. Thus, in experimental canine OA and as early as at 12 weeks after joint destabilization, levels of the C2C neoepitope increase approximately 32-fold in synovial fluid and about 1.5-fold both in serum and urine [84] whereas the release of the C1,2C neoepitope is enhanced in the culture medium of human osteoarthritic cartilage explants and this increased release is selectively inhibited by a synthetic inhibitor that spares MMP-1, but not MMP-8 or MMP-13 [75]. Synovial fluid levels of the C2C neoepitope are also increased in experimental inflammatory arthritis [85] and in the early stages of human RA [57]. It is worth noting that in human RA the synovial levels of C2C correlate strongly not only with the synovial fluid levels of tumor necrosis factor alpha (TNF-α) but also with the serum levels of C-reactive protein [57]. The baseline serum ratio of C1,2C to C2C, but not the levels of individual markers, is predictive of disease progression in patients with knee OA [86]. The neoepitope COLL2-1 located near the N-terminus of the TCA fragment (Table I) and its nitrated form (COLL2-1-NO2) are also released into the synovial fluid and from there on reach the systemic circulation [87]. Serum levels of COLL-1 are increased 1.5-fold in both OA and RA whereas serum levels of COLL-1-NO2 are enhanced about two-fold in OA and three-fold in RA [88]. Since chondrocytes and other cells of the joint organ produce nitric oxide and superoxide anion, which then react to yield peroxynitrite anion, these neoepitopes might be promising tools to evaluate the contribution of oxygenderived species to the degradation of type II collagen [89–91]. Some degradation products of type II collagen may, acting in a positive-feedback mechanism, stimulate collagen gene expression and promote the differentiation of cartilage cells [68]. There is indeed evidence that cleaved, denatured type II collagen not only can induce the production of MMP-1, MMP-13, interleukin-1 (IL-1), and TNF-α by chondrocytes but also, and most importantly, can induce hypertrophy of cartilage cells, a process characterized by the expression of type X collagen and the up-regulation of cbfa1, the MMP-13 transcription factor [68].

DANIEL-HENRI MANICOURT ET AL.

Therefore, chondrocyte apoptosis and calcification of cartilage matrix, which are both commonly observed in OA cartilage, might result, at least in part, from chondrocyte hypertrophy [92]. On the other hand, an evolutionary conserved domain of type II collagen, located between amino acids 359 and 369, might be a target of pathogenic autoimmune responses in the induction or perpetuation of joint inflammation in RA [93]. Little is known about the degradation of other collagens in cartilage. Type IX collagen can be cleaved by MMP-3 in its NC2 domain [64] and by an unknown protease in its NC4 domain [94]. Loss of the NC4 domain is an early event in experimental inflammatory arthritis: it accompanies proteoglycan loss and precedes detectable damage to type II collagen within the cartilage [85]. Type IX collagen degradation might decrease the tensile stiffness of cartilage since this collagen protrudes from the surface of the type II collagen fibril where it apparently functions as a covalent bridge between collagenous fibrils [59]. On the other hand, the degradation of type IX collagen also might allow the diameter of collagen fibrils to increase during growth and maturation since the type IX molecules must be removed before fibril growth can occur, either by accretion of additional type II molecules and/or by lateral fusion of thin fibrils (see Chapter 25). When cartilage collagens are degraded, crosslinked N- and C-telopeptide fragments bearing pyridinoline crosslinks are released into proximal fluids, i.e. synovial fluid in the case of a diarthrodial joint, and from there on enter the systemic circulation before being most effectively cleared by filtration through the kidneys. Pioneering work has detected pyridinoline, but not deoxypyridinoline (the “bone-specific” crosslink), in OA and RA synovial fluids [95]. Although the urinary excretion of pyridinoline increased approximately two-fold in patients with OA and four-fold in patients with RA [96–98], it was not clear how much of this increase could be attributed to degradative changes occurring in cartilaginous structures, since the increase in urinary pyridinoline originating from cartilage is likely to be blurred by the concomitant increase in the turnover of other structures of the diseased joints, including bone and synovium [99–101], a contention strengthened by the observation that patients with knee and hip OA exhibit high urinary levels of the glycosylated pyridinoline derivative, glucosyl-galactosyl-pyridinoline, which is found in large amounts in human synovium, and in very low levels in the cartilage and other soft tissues [102]. The development of immunoassays capable of quantifying the neoepitope EKGPDP bound to the crosslinks and generated by proteolytic cleavage of the C-telopeptide domain of type II collagen fibrils has dramatically simplified

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Chapter 25 Products of Cartilage Metabolism

the interpretation of analyses of pyridinium crosslinks in body fluids [103, 104]. Several MMPs, including MMP-8, MMP-13, and MMP-14, can generate this neoepitope termed CTX-II, COL2CTX, or CartiLaps, but MMP-7 (matrilysin) is the most potent [103]. Enhanced levels of CTX-II have been found in the culture medium of human articular cartilage explants stimulated with the catabolic cytokine oncostatin M [105] as well as in the synovial fluid of patients with joint injury, OA, pseudogout, and septic arthritis [106]. In the early stages of canine experimental OA, levels are increased by an order of magnitude of six in synovial fluid and of two in serum [84]. The urinary excretion of CTX-II increases approximately 1.5-fold in patients with OA and 2.3-fold in patients with RA [52, 104]. Further, in patients with knee and hip OA, levels of the urinary excretion of CTX-II are associated with both the prevalence and the progression of radiographic OA changes, and this independently of known clinical risk factors for osteoarthritis [107]. Further, in patients with early RA, the magnitude of the changes of urinary CTX-II levels at 3 months after induction of therapy are predictive, independent of changes in disease activity, of the changes in radiologic score of joint destruction after 1 year [108]. Combining urinary CTX-II excretion, a marker of type II collagen degradation, with serum levels of N-propeptide of type IIA procollagen, a marker of type II collagen synthesis, might be more effective in predicting

Figure 3

cartilage destruction than measurements of a single marker [52].

IV. PRODUCTS OF AGGRECAN METABOLISM The numerous chondroitin sulfate (CS) and keratan sulfate (KS) chains carried by the long-extended interglobular domain (IGD) of the aggrecan core protein confer to this PG a bottle-brush structure with viscoelastic properties (Fig. 3). This elaborate architecture, however, leaves vulnerable to proteolytic attack the glycosaminoglycan (GAG)-poor N-terminal end of the molecule that separates the glycosaminoglycan (GAG) attachment region from the first globular domain (G1) that interacts with HA and LP and thus anchors the aggrecan molecule in the matrix. Newly secreted aggrecan molecules have low affinity for HA and their maturation into the high-affinity form probably involves the formation of disulfide bonds in the G1 domain (see Chapter 5). This, coupled to an accelerated transport, termed “rapid polymer transport”, may enable newly secreted molecules to diffuse away from the metabolically active pericellular environment and enter the metabolically inactive interterritorial matrix compartment before aggregating [109]. In contrast, LP molecules are fully capable of binding to both HA and aggrecan molecules

Through its globular domain G1, aggrecan interacts with hyaluronan (HA) and link protein (LP) to form supramolecular aggregates of very large size. The function of G2 is unknown and G3 has a role in intracellular translocation. Some of the antibodies that react with the core protein and the chondroitin sulfate (CS) and keratan sulfate (KS) chains are indicated. Other monoclonal antibodies (Mabs) react with the chondroitin sulfate chain that remains attached to the core protein after chondroitinase ABC lyase digestion. Adapted from reference [43].

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when they are released extracellularly. Binding of LP to newly synthesized aggrecan molecules might provide the stability the latter need for interacting early with HA [110]: this may play a key role in determining what proportion of newly synthesized aggrecan molecules become established in the pericellular/cell-associated matrix. While studies of the racemization of aspartic acid have suggested that aggrecan molecules turn over much more rapidly (average half-life = approximately 2 years) than collagen molecules within the fibrillar network [111, 112], the results of both in vivo [113] and in vitro [109, 114, 115] studies have suggested that articular cartilage contains at least two pools of aggrecan molecules with different rates of turnover. It is thought that the two pools of aggrecan molecules reside in different compartments of the matrix: a cell-associated matrix compartment in which they turn over rapidly and a further removed, more abundant, interterritorial matrix compartment in which the PGs turn over much more slowly [109, 115]. The catabolism of aggrecan is a proteolytic process and multiple cleavage sites have been observed along the protein core [116–119] (Fig. 4). GAG-rich fragments generated by proteolytic cleavage of the core protein are lost relatively rapidly from the cartilage matrix. In the case of articular cartilage, they diffuse into the synovial fluid

Figure 4

where they can be quantified by chemical assays of their sulfated glycosaminoglycans [120, 121] or by immunoassays capable of measuring specific carbohydrate epitopes [84, 122–125]. They also can be characterized by peptide sequence analysis after isolation and deglycosylation [126, 127]. The development of monospecific antibodies that recognize enzyme-generated neoepitopes has proved most useful for identifying and quantifying specific proteolytic cleavage products [128–134]. Proteinases of different classes and hydroxyl radicals [135] have the capacity to cleave the aggrecan core protein in vitro. Importantly, the use of antibodies against these types of neoepitopes has now provided strong evidence that at least two families of zinc-dependent metalloenzymes, the metalloproteinases (MMPs) and the ADAMTS or aggrecanases, play major roles in vivo in the catabolism of aggrecan molecules both in health and disease (Fig. 4). When proteolytic processing is confined to the C-terminus of the GAG-rich region, the aggrecan molecules remaining within cartilage matrix retain some GAGs and thus are able to continue to exert their physicochemical properties. The contention that progressive C-terminal trimming of aggrecan molecules does occur in vivo is supported by N-terminal sequencing of aggrecan fragments released into the conditioned medium of cartilage explants

Major sites of cleavage of aggrecan core protein by tissue proteinases. The two major sites of cleavage are located in the short extended interglobular domain between the globular domain G1 and the globular domain G2: the first one occurs between residues Asn341– Phe342 whereas the second one is located between residues Glu 373 and Ala374. Both cleavage sites separate the anchoring G1 domain from the bulk of the aggrecan molecule carrying the glycosaminoglycan side chains and thus deprive cartilage from its load-bearing properties. The N-terminal and C-terminal amino acid sequences of the neoepitopes resulting from the different cleavage sites as well as the nature of the proteinases acting at these sites are also indicated. The correct size of different domains is not respected. A disintegrin and metalloproteinase with thrombospondin-like motifs (ADAMTS); cathepsin B (cat B); matrix metalloproteinase (MMP).

Chapter 25 Products of Cartilage Metabolism

[126, 136, 137], as well as by physiochemical and electron microscopic analysis of aggrecan molecules extracted from cartilage. These studies have identified in adult articular cartilage an abundance of polydisperse molecules lacking the G3 domain and exhibiting a great variability in the length of their CS-rich domain [116, 138–143]. This C-terminal processing of aggrecan also occurs early in growth and might be viewed as a “nondestructive” tailoring of aggrecan that promotes cartilage matrix assembly and organization [144]. On the other hand, too extensive C-terminal processing of aggrecan with significant loss of CS-containing regions of the molecule might contribute significantly to loss of tissue function both in aging and disease. Although the C-termini of the majority of truncated aggrecans are apparently located in regions of the core protein with known MMP-sensitive cleavage sites, their amino acid sequence remains unknown, and thus the nature of the proteinase(s) responsible for the generation of these products is still unsolved. On the other hand, cleavage of the core protein near the G1 domain is more “destructive” as it results in rapid loss of the whole GAG-attachment region and thus has more adverse consequences, at least from the functional standpoint [145]. Two critical cleavage sites have been identified in the short extended IGD that separates the G1 and G2 domains: one occurs at the N341 ↓ F342 bond and the other site is located at the E373 ↓ A374 bond (Fig. 4). The first critical cleavage site generates a G1-containing fragment with the C-terminal neoepitope VDIPEN341 and a larger GAG-bearing fragment with the N-terminal neoepitope 342FFGVGGE [132, 133, 146–148]. It is the major cleavage site of several metalloproteinases including stromelysin-1 (MMP-3 ), the three collagenases (MMP-1, MMP- 8, and MMP-13), the two gelatinases (MMP-2 and MMP-9) as well as MMP-7 and MMP-14 (MT1-MMP). It can also be cleaved by the combined endopeptidase and carboxypeptidase activity of cathepsin B, but at low pH [149]. MMPs appear to be involved in the conservative baseline turnover of aggrecan: MMP-derived 342FFGVG fragments are found in small amounts in the culture medium of pig articular cartilage where they represent about 1% of the total amount of aggrecan released [150, 151]. Further, and as observed in human synovial fluids, these fragments recovered in culture medium usually have a small size, and, because they do not carry the NITEGE373 C-terminus generated by aggrecanases, they are likely to be the product of more extensive MMP processing [131, 133]. However, while most degraded aggrecan components are quickly released in culture, the majority of 342FFGVG fragments are retained in cartilage tissue. Even upon stimulation with IL-1, the 342FFGVG fragments increase significantly

431 in the tissue without a corresponding increase in 342FFGVG neoepitope in the medium. Because the 342FFGVG products of MMP catabolism are large, resistant to destruction by aggrecanases and selectively retained in cartilage, they may have a role in conveying signals or organizing a microenvironment [151]. The 342FFGVG fragments remaining in cartilage matrix could represent a non-negligible proportion of the “nonaggregating” large proteoglycans of cartilage and/or contribute to the metabolic pool of aggrecans which exhibit a long half-life [109, 152]. MMPs have the capacity to cleave at other sites within the short IGD of aggrecan molecules, but this requires higher enzyme concentrations. Thus, MMP-1, MMP-7, MMP-8, and MMP-13 all cleave the D441 ↓ L442 bond (Fig. 4), whereas MMP-3 also cleaves at S377 ↓ V378, MMP-8 at Q373 ↓ A374, and MMP-13 at P384 ↓ V385 [148]. Although it may be generated by both MMP-8 and MMP-14 [153], the second critical cleavage at the E373 ↓ A374 bond is believed to be the signature of the glutamyl endopeptidases termed ADAMTS [153–155]. This produces a large GAG-rich aggrecan fragment with the N-terminal neoepitope 374ARGSVI and a G1 fragment with the C-terminal neoepitope NITEGE373 (Fig. 4). The bulk of aggrecan fragments found in synovial fluid of patients with joint injury, OA, rheumatoid arthritis (RA), and other types of inflammatory arthritis all have the N-terminal sequence ARGSVI [127, 156, 157]. These fragments are also found in enhanced amounts in the culture media of both chondrocytes and cartilage explants stimulated to undergo matrix degradation with IL-1 or retinoic acid [154, 158, 159]. However, the cleavage site at the E373 ↓ A374 bond within the short IGD of aggrecan molecules is not the preferential site of recombinant ADAMTS-4 and ADAMTS-5: the enzymes first cleave within the CS-rich G2–G3 domain at the KEEE1714 ↓ 1715GLGS bond to release the COOH terminus of the molecule (Fig. 4) before contributing to the removal of an additional portion of the COOH terminus by cleaving at the ASELE1539 ↓ 1540GRGT bond [119, 155]. While the C-terminus portion of the molecule released by the first cleavage is further processed at two additional cleavage sites, at TAQE1819 ↓ 1820AGEG bond and at ISQE 1919 ↓ 1920LGQR bond, the cleavage within the short IGD at the E373 ↓ A374 bond occurs more slowly. Thus far, it is not clear whether the first cleavages in the C-terminal portion of the core protein are required and/or facilitate the cleavage in the short IGD domain of the aggrecan molecule. On the other hand, it is clear that the successive cleavages of the protein core within the GAG-rich region are facilitated by the binding of the C-terminal thrombospondin motif of ADAMTS to the GAG side chains [118] and, accordingly, exogenous chondroitin sulfate and

432 heparin both inhibit aggrecanase activity [160], whereas changes in glycosylation with age may alter susceptibility to cleavage by ADAMTS [161]. It is now obvious that ADAMTS-mediated aggrecan cleavages occur both in vivo and in vitro: the neoepitopes generated by the four cleavage sites located within the GAG-rich domain (E1539 ↓ G1540, E1714 ↓ G1715, E1819 ↓ A1820, and E1919 ↓ L1920) have been found in both human and bovine articular cartilage [137, 144], in rat chondrosarcoma [162], in the culture medium of cartilage explants stimulated with either IL-1 and/or retinoic acid [116, 119, 137] as well as in high-density aggrecan fragments recovered from human synovial fluids [144]. After cleavage at the two critical sites within the short IGD, the majority of VDIPEN341 and NITEGE373 neoepitopes resident on G1 fragments remain in tissue bound to hyaluronan. Both neoepitopes are detected not only in cartilage from joints with OA and RA, two conditions exhibiting quite contrasting pathological and clinical features, but also in cartilage from normal adult joints where they appear to accumulate with age [134, 163]. The generation of these neoepitopes is not necessarily coordinated since both the NITEGE373 and VDIPEN341 neoepitopes can map predominantly to different regions within a single normal joint, can show a temporal and spatial separation in arthritis models and in developing cartilage [67, 132, 134, 164, 165], and since turnover of aggrecan by cultured rat chondrosarcoma cells and primary bovine chondrocytes can be mediated almost exclusively by aggrecanase [130]. On the other hand, although there is evidence that MMP cleavage of the core protein at the N341 ↓ F342 bond and ADAMTS cleavage at the E373 ↓ A374 bond are independent [134, 166] and mutually exclusive [148, 167], the detection of the VDIPEN341 neoepitope does not prove that the primary cleavage of the core protein results from MMP activity as MMPs and ADAMTS4 both can cleave the N341 ↓ F342 bond of a G1NITEGE373 fragment to yield a G1- VDIPEN341 fragment [167, 168]. It is generally believed that, in vivo, ADAMTS play a major role in the degradation of aggrecan during the early stages of cartilage degradation whereas MMP-mediated catabolism of aggrecan occurs as a late event in cartilage matrix breakdown, at the time when active degradation of collagen is occurring [124, 169]. Indeed, in both models culture systems and murine arthritis, the NITEGE373 neoepitope appears very early, well before the appearance of the VDIPEN341 neoepitope, and importantly in proportion to GAG loss from the tissue [124, 170–172]. The ADAMTS-mediated early loss of aggrecan from cartilage could facilitate collagen proteolytic cleavage [173]. On the other hand, studies conducted in MMP-3-deficient mice [67] have pointed out that, although MMP-3 plays

DANIEL-HENRI MANICOURT ET AL.

no role in early aggrecan loss, this MMP is essential for generating both aggrecan and collagen neoepitopes, and that these epitopes co-localize and correlate with disease progression. Although ADAMTS engender cartilage aggrecan catabolism during the primary phases of arthritic diseases, it is still not known which iso-form(s) play(s) a major role. Regulation of the ADAMTS genes is still poorly understood, though there is evidence that the effect of growth factors, hormones, and inflammatory cytokines are dependent on age, species, and stimulus [124]. In bovine cartilage ADAMTS4 is not constitutively expressed but is up-regulated by IL-1, TNF-α, and fibronectin fragments whereas ADAMTS5 is constitutively expressed with little up-regulation in response to cytokines [154]. In another study, 90% of the aggrecanase activity in media from bovine articular cartilage stimulated with IL-1 could be inhibited by immunoprecipitation with a combination of antibodies to ADAMTS4 and ADAMTS5, indicating that these proteins are primarily responsible for the aggrecanase activity stimulated by IL-1 in this system [119]. On the other hand, retinoic acid induces a marked increase in the release of ADAMTS-generated aggrecan catabolites from human cartilage with no concomitant increase in the mRNA levels for any of ADAMTS1, 2, 3, 4, and 5 [174], whereas ADAMTS1, 4, and 5 mRNAs are all regulated by a combination of IL-1 and oncostatin M, though with differing magnitude and kinetics [175]. IL-17 also up-regulates ADAMTS4 via phosphorylation of ERK, p38, and JNK MAP kinases [176]. In human OA cartilage samples obtained at late stages of the disease process, ADAMTS1, 5, 9, and 15 were down-regulated while ADAMTS2, 12, 14, and 16 were up-regulated by comparison with phenotypically normal cartilage obtained from fracture patients [177]. Interestingly, the incorporation of n-3 fatty acids into chondrocyte membranes causes a dosedependent reduction in both the expression and activity of ADAMTS [178]. As observed for MMPs, control of ADAMTS activity is exerted at multiple points including zymogen activation, substrate accessibility, and inhibition by naturally occurring inhibitors. In the case of ADAMTS4, which is the best characterized of the aggrecanases, C-terminal processing of the 75-kDa full-length active form produces isoforms of 60 and 50 kDa [179]: this results in the release of the enzyme from the ECM and alteration of its profile activity [180]. The C-terminal spacer region of the 75-kDa isoform can act to inhibit the aggrecanase activity at the E373 ↓ A374 bond because the 75-kDa isoform cleaves at E1480 ↓ G 1481 but not at E373 ↓ A374: this is in contrast to the 60- and 50-kDa isoforms, which both lack the spacer region and cleave at the E373 ↓ A374 bond. Further, binding of the C-terminal spacer region to fibronectin could anchor

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the 75-kDa isoform to the ECM and/or modulate aggrecanase substrate specificity [181, 182]. On the other hand, although the tissue inhibitors of MMPs (TIMPs) are broadly effective inhibitors of the MMPs [183], it is clear that they display much greater selectivity towards ADAMTS. ADAMTS4 and ADAMTS5 are both potentially inhibited by TIMP-3, while being insensitive to TIMP-1, 2, and 4 [184], whereas ADAMTS1 is partially inhibited by TIMP-2 and TIMP-3 [185]. Alpha-2macroglobulin is another endogenous inhibitor of both ADAMTS4 and 5 [186]. As stated in Chapter 5, several glycosyl- and sulfotransferase enzymes are implicated in the biosynthetic pathways of aggrecan CS chains whose disaccharide units consist of glucuronic acid (GlcA) and N-acetyl-galactosamine (GalNAc). The ratio of 6- to 4-sulfation of the GalNAc residues present in the internal disaccharides increases markedly with age and within the successive cartilage zones with distance from the articular surface [187, 188]. Because the distinct 6-sulfotransferase activity, with specificity for the nonreducing terminal GalNAc, appears after skeletal maturity, the majority of terminal GalNAc residues are 4-sulfated in juvenile aggrecans and 4,6disulfated in adult aggrecans [187]. In both normal adult

Table II.

and OA articular cartilage, the size of CS chains on aggrecan is similar but smaller to that of juvenile aggrecans; however, alterations in the GAG biosynthesis pathways of OA chondrocytes lower the capacity for 4,6-disulfation of the nonreducing termini of CS chains, thus resulting in the synthesis and deposition of CS chains with predominantly nonsulfated termini. Therefore, based on CS chain length and pattern of sulfation of nonreducing termini, it is tempting to speculate that OA chondrocytes synthesize aggrecans substituted with CS chains similar to those found in juvenile aggrecans [189]. The fine structure of CS chains is sensitive to the physicochemical environment (pH, ionic strength) of the biosynthetically active chondrocytes [17] and can also be modulated by growth factors [190], cytokines [191], and mechanical factors [192]. Dynamic compression of cartilage tissue increases not only the length of CS chains but also decreases the abundance of the 4,6-disulfated nonreducing terminal N-acetylgalactosamine residues [192]. Presumably, these marked changes in the sulfation pattern of CS aggrecan chains initiated by chondrocytes enable the cartilage tissue to respond more efficiently to biological changes in the cell environment, and in turn, the physiochemical properties of aggrecans are influenced by these structural events.

Monoclonal Antibodies Commenly Used to Analyze Aggrecan and Other Components of Proteoglycan Aggregates

Monoclonal antoibody

Antibody specificity

5-D-4 and 1-B-4

Skeletal and corneal keratan sulfate. Highly sulfated sequence of serveral repeats of 6-sulfated N-acetylglucosamine-b1, 3-6sulfated galactose-b1, 4 Keratan sulfate. Sulfated poly (N-acetyl lactosamine) sequence of hepta- or larger oligosaccharides of 6-sulfated galactose and N-acetylglucosamine Generated by chondroitinase digestion of aggrecans. Delta-unsaturated disaccharides of chondroitin-4-sulfate, but not chondroitin-6-sulfate and unsulfated chondroitin Intact chondroitin-6-sulfate chains but not unsaturated disaccharides of chondroitin-6-sulfate Nonreducing termini of native chondroitin sulfate chains containing a terminal hexuronate residue adjacent to a 6-sulfated N-acetylglucosamine residue. Delta-unsaturated or saturated oligosaccharides of chondroitin-6-sulfate generated by chondroitinase or tessticular hyaluronidase digestion Nonreducing termini of native chondroitin sulfate chains containing a terminal hexuronate residue adjacent to a 4-sulfated N-acetylglucosamine residue. Delta-unsatutated or saturated oligosaccharides of chondroitin-4-sulfate generated by chondroitinase or testicular hyaluronidase digestion Delta-unsaturated or saturated oligosaccharides of chondroitin-4-sulfate generated by chondroitinase or testicular hyaluronidase digestion Delta-unsaturated or saturated oligosaccharides of unsulfated chondroitin generated by chondroitinase or testicular hyaluronidase digestion Chondroitin sulfate oligosaccharide containing 10–12 sugar residues. Probably contains several of the eight possible CS-isomer disaccharide combinations Complex epitopes residing within native chondroitin sulfate chains

AN9P1 2-B-4 (B) and 9-A-2 (B) 846 (A) 3-B-3 (C)

3-D-5 (C)

2-B-6 (B) 1-B-5 (B) 7-D-4 (A) 4-C-3 (A) 4-D-3 (A) 6-C-3 (A) 8-A-4 8-A-5

Determinant comment to link protein (LP) isolated from aggregates recovered from cartilage of different species Determinant present in the three molecular forms of LP (LP1, LP2, and LP3) Similar to that of 8-A-4. Reduction and carboxymethylation of the link protein does not alter the epitope

Note. The list is not exhaustive. Monoclonal antibodies directed against chondroitin sulfate epitopes can be divided into two categories: (1) those that recognize epitopes present on the native chondroitin sulfate glycosaminoglycan chains (A) and (2) those that require predigestion of the chondroitin sulfate glycosaminoglycan with endo- or exoglycosidases to generate their reducing teminal epitopes (B). Some antibodies (C) recognize epitopes that fall into both of these categories.

434 Monoclonal antibodies have been produced with specificities against structural carbohydrate epitopes on the CS chains (Table II and Fig. 3). With the exception of the 3-B-3 epitope, which corresponds to the nonreducing terminal sequence GluAβ1, 3GalNac-6-sulfate, the precise structure of these (neo)epitopes remains to be defined and it is not known to what extent their reactivity detected by Western blots or immuno-histochemistry could be affected by the CS chain length and/or degree of sulfation [187]. Two epitopes, termed 3-D-5 and 7-D-4, are present in normal mature articular cartilage [193]. The content of the 7-D-4 epitope in aggrecan increases markedly in diseased cartilage from different species [193–195] as well as in the synovial fluid from human knees with acute traumatic cruciate ligament and meniscal injuries [196], but it is decreased in RA synovial fluids where it shows a negative association with the degree of joint inflammation [197]. The CS epitope termed 846 does not contain unsaturated disaccharides of chondroitin-6-sulfate [84, 198]: it is found at highest concentrations in fetal cartilage; it is barely detectable in adult cartilage but reappears in enhanced amounts in cartilage from OA and RA joints [195, 198, 199]. As it is present on intact CS chains located in close proximity to the G3 domain of the core protein of newly synthesized aggrecan molecules, it may be a useful marker of aggrecan synthesis during inflammation and disease processes [84, 198]. While the percentage of CS chains bearing the 3-B-3 epitope in articular cartilage remains constant, at about 9%, throughout life [187], its immunoreactivity varies widely with the stage of development, maturity, and pathology [194, 195, 200]. This apparent discrepancy might be related, at least in part, to the observation that the recognition of CS chain termination by the 3-B-3 antibody on solid phase assay could be influenced by the sulfation and/or length of the chains ending with 3-B-3 epitope [187, 189]. Accordingly, the enhanced levels of 3-B-3 reactive aggrecan fragments detected in human synovial fluid after traumatic knee injury [196] might just reflect the release of degraded aggrecans that are resident in the healthy adult cartilage, whereas the increased levels observed in human OA synovial fluid [197, 201] might relate to a change in metabolic activity of chondrocytes, with alterations in the sulfation and length of the CS chains modulating the immunoreactivity of the chains bearing the 3-B-3 epitope. In OA patients, synovial fluid levels of the 3-B-3 epitope cannot distinguish between progressive and nonprogressive OA [202], and do not change significantly after a 12-week program of walking exercise [201]. Measurement of GAG epitopes can also provide a measure of the turnover of PGs. Immunoassays that use

DANIEL-HENRI MANICOURT ET AL.

monoclonal antibodies (Table II and Fig. 3) capable of recognizing KS epitope(s) in intact aggrecan molecules or fragments thereof [203, 204], have shown that levels of antigenic KS in the blood circulation correlate well with the concentration of aggrecan core protein-related epitopes in serum, suggesting that the serum levels of agKS offer a good measure of the level of aggrecan-derived fragments in body fluid (Chapter 32). Although the function of KS is still unclear, the rate of synthesis of this molecule appears to be inversely related to the ambient oxygen tension [205], which may explain why the the KS content of aggrecan increases with distance from the articular surface [206]. On the other hand, aggrecans exhibit an agerelated increase in the number, size, and sulfation of their KS chains whereas the KS content of aggrecans recovered from OA cartilage is markedly reduced [207, 208].

V. PRODUCTS OF THE METABOLISM OF OTHER PROTEOGLYCANS Members of the family of Small Leucine-Rich Proteoglycans (PGs), termed the SLRPs, have been identified in adult articular cartilage [209] (see also Chapter 4). Biglycan, decorin, and fibromodulin all have the capacity to bind to transforming growth factor betas (TGF-β) via their core protein with both high-affinity and low-affinity binding sites, and hence these small PGs may play an important role in regulating the activity of TGF-β and its impact on the synthesis of matrix components. On the other hand, studies conducted in cartilage explants have pointed out that TGF-β enhances the rate of synthesis of both decorin and fibromodulin, but not biglycan [210]. The surface of collagen fibers is decorated by decorin and fibromodulin, and the message level for these two members of the SLRPs family shows increases with increasing age in human articular cartilage [211]. Although fibromodulin might regulate collagen fibril diameter, this does not seem to be true for decorin as no effects on type II collagen fiber diameter in the decorin knockout mice have been reported as of yet [212]. Because of its specific localization in the pericellular space as well as on the chondrocyte cell surface, biglycan may have important functions in modulating morphogenesis and differentiation. Mice with a targeted disruption of the biglycan gene are apparently normal at birth, with no skeletal patterning defects [213]. However, the biglycan-null mice, that express unaffected levels of decorin, showed decreased postnatal skeletal growth, indicating that biglycan is a positive regulator of bone formation and bone mass. Fibromodulin and decorin can be cleaved by ADAMTS-4 [214]. Although Poole et al. [215] reported

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no overall differences in biglycan and decorin contents in human knee OA, other studies have reported an increase in the content of intact biglycan, decorin, and fibromodulin as well as fragments of these PGs in both human OA and RA cartilage [216–218], suggesting they are involved in the attempt at matrix repair by chondrocytes within diseased cartilage. On the other hand, patients with RA expressed an increased humoral immunity to biglycan whereas patients with seronegative spondyl-arthropathies exhibit enhanced decorin-specific antibodies [219]. Lumican and the protein known as PRELP (Proline arginine-Rich End Leucine-rich repeat Protein) are two other members of the SLRPs found in cartilage matrix. The glycosylation pattern of lumican by chondrocytes can be modulated by interactions of the cell with surrounding matrix components as well as by growth factors and cytokines: thus, while basic fibroblast factor, insulin-like growth factor and TGF-β all induce the secretion of a lumican form bearing KS chains, IL-1 promotes the secretion of a KS-deficient form of lumican [220]. On the other hand, PRELP has four potential N-linked glycosylation sites and apparently functions as a cartilage matrix protein with the capacity for matrix organization [221]. Since it has the capacity to simultaneously bind collagen and perlecan, PRELP might function as a linker between the matrix close to the chondrocyte and the more distant matrix via perlecan [222]. In adult articular cartilage, perlecan is a CS/heparin sulfate hybrid PG that is found in the territorial matrix with highest intensity in the pericellular region [223]. Human perlecan deficiencies have been found in chondrodysplasias [224, 225]. Mice lacking perlecan develop a severe cartilage phenotype characterized by a reduced collagen network and shorter collagen fibrils, implying an important role for perlecan in the cartilage structure [225, 226]. Perlecan might also maintain chondrocyte differentiation, perhaps working in concert with SOX9, and/or influence chondrocyte metabolism via signaling receptors or by modulating the expression of fibroblast growth factors [227, 228].

VI. PRODUCTS OF THE METABOLISM OF LINK PROTEIN AND HYALURONAN The synthesis of LP follows the biosynthetic pathway of other secretory proteins through the endoplasmic reticulum and the Golgi. SOX9 and its known transcriptional co-activators L-SOX5 and SOX6 are potent inducers of LP expression [229] whereas TNF-α suppresses LP expression through the MEK1/2 and NF-κB pathways [230]. Although the rate of LP synthesis is up-regulated by mechanical compression of cartilage [231], the

age-related decrease in the rate of LP synthesis might regulate the rate at which newly synthesized aggrecan acquires affinity for HA [232]. Human articular cartilage LP can be separated by SDS/PAGE into three components of molecular mass 48, 44, and 41 kDa, referred to as LP-1, LP-2, and LP-3, respectively [233]. LP-1 and LP-2 are different glycosylated forms of the same intact protein core, while LP-3 is proteolytically derived from either LP-1 or LP-2. In cartilage of newborns, LP-3 is a minor component, while, with age, LP-3 accumulates at the expense of LP-1 and LP-2. Three major cleavage sites have been identified in the N-terminal region of intact LPs (Fig. 5). The first one occurs at the H18 ↓ I19 bond: it is generated by stromelysins 1 and 2, gelatinases A and B as well as by collagenase [234]. Before being inactivated by rapid specific cleavage mediated by membrane-associated proteinases [235], the cleaved N-terminal peptide (DHLSDNYTLDHDRAIH or Link N) produced thereof stimulates the biosynthesis of both PG and type II collagen synthesis by human articular cartilage [236, 237] and hence this peptide may have an important role in the feedback control of cartilage matrix synthesis. On the other hand, LP3 from mature cartilage exhibits two additional amino termini: one is compatible with

Figure 5

Sites of cleavage of link protein (LP) in human articular cartilage. In vitro many proteolytic agents are able to cleave the link protein within the N-terminal region between amino acid residues 14 and 29 to produce the LP3 forms of the molecule. In contrast, only a few proteolytic agents can cleave the link protein in vitro within the disulfide-bonded loop between amino acid residues 56 and 87 to produce LP fragments upon reduction. The site for LP3 generation (1 between residues 16–17; 2 between residues 18–19 and 3 between residues 23–24) and LP fragment generation (4 between residues 65–66 and 5 between residues 72–73) in vivo occur within the same regions that are susceptible to in vitro proteolysis. The sites cleaved by metalloproteinases are also shown. Adapted from reference [233].

436 cleavage at the E18 ↓ A19 bond, a property of cathepsins B and G, whereas the other one occurs at the P23 ↓ H24 bond; the enzyme(s) responsible for this site of cleavage has not been identified. As the three forms of LP possess the three disulfide-bonded loops that are associated with LP function, LP3 generation is unlikely to affect the stability of PG aggregates. In vitro studies have also pointed out that the N-terminal disulfide-bonded loop of LP molecules can be cleaved at one site by cathepsin L [233], and at two sites by hydroxyl radicals [238]; however, none of these corresponds to the sites of cleavage observed in vivo, i.e. at the K65 ↓ W66 bond and the D72 ↓ Y73 bond. Whether these native cleavage sites result from the action of a yet to be identified enzyme remains to be established. On the other hand, the continued ability of fragmented LP molecules to interact with HA probably reflects the lack of proteolytic modification within the C-terminal disulfide-bonded loops. Very little is known about the half-life and turnover of LP in the cartilage matrix. During arthritides, the loss of LP from cartilage matrix into the synovial fluid may elicit the local production of antibodies and the expression of cellular immunity to LP: patients with RA, juvenile RA (JRA), ankylosing spondylitis, and OA express cellular immunity to LP, which is correlated with disease activity and severity in JRA [239–242]. In canine experimental OA, the synovial fluid levels of LP and the rate of loss of this glycoprotein from cultured cartilage explants increases several-fold within 3 months of disease induction [243, 244]. In contrast, the synovial fluid levels of LP are markedly decreased in disused joints [243]. These findings are in close agreement with the progressive disappearance of the link-rich, more saturated, aggregates from the cartilage matrix in experimental OA and with the maintenance of these fast-sedimenting aggregates in disuse atrophy [26]. Although articular cartilage from canine OA joints and from disused joints both exhibit enhanced levels of matrixdegrading enzymes [27], these cartilages differ in their HA content: the latter is markedly reduced in OA and unchanged in disuse. The same holds true for human OA cartilage. Since, in contrast to the disuse process, the OA process is apparently irreversible, one might argue that the loss of both HA and link-rich aggregates prognosticates articular cartilage destruction. Three mammalian HA synthase (HAS) genes have been cloned and referred to as HAS1, HAS2, and HAS3 [245]; the different HA isoforms might have different enzymatic properties as the size of HA chains secreted by HAS2 transfectants is larger than that produced following transfection with HAS1 or HAS3. While HAS2 is the major isoform expressed in chondrocytes, the presence of HAS1 is controversial and that of HAS3 could vary with age and in response to cytokine/growth factors [246–248].

DANIEL-HENRI MANICOURT ET AL.

This notwithstanding, HA is synthesized on the inner side of the plasma membrane: chain elongation occurs at the nonreducing end of the growing HA chain [249], which is extruded into the extracellular space [245, 250]. This unusual mode of synthesis allows unconstrained polymer growth and explains why HA molecules can reach exceptionally large sizes. Half of the HA molecules synthesized by cultured explants of normal articular cartilage diffuses rapidly into the culture medium [251]. The newly synthesized HA molecules remaining within the matrix are quite large, but with time they are gradually depolymerized to give a molecular size distribution similar to that observed for endogenous HA [251, 252]. This depolymerization could result from the action of either a recently discovered hyaluronidase active at neutral pH [253] and/or oxygenderived free radicals [254, 255]. In contrast to their native HA molecules, HA degradation fragments induce the expression of inflammatory genes in different cell lines [256]. By binding to the cell surface receptor CD44, the HA molecules (and their bound aggrecan molecules) contribute to the formation of the pericellular matrix and maintenance of matrix homeostasis [257]. Indeed, reduction of HAS2 expression by antisense inhibition reduces the size of the cell-associated matrix and the capacity of the chondrocytes to retain PGs [246], whereas disruption of chondrocyte CD44 receptor and HA interaction by either HA oligosaccharides and/or CD44 antisense oligonucleotides induces a cascade of events resulting in the activation of both catabolic and anabolic gene products [258]. It is worth noting that osteogenic protein-1, also known as bone morphogenic protein-7, inhibits the HA hexosaccharide-induced depletion of HA and PGs within cartilage matrix, thus providing further support for the emerging paradigm that cell–matrix interactions modulate the cellular response to morphogens or growth factors [259]. On the other hand, chondrocytes also use the CD44 receptor to internalize some of the HA molecules, which are then degraded to small oligosaccharides within a low pH lysosomal compartment [257]. Factors that up-regulate chondrocyte catabolism, such as interleukin-1 and the 29-kDa fibronectin fragment, also cause an increased expression of CD44 at the cell surface. As cartilage is bathed by synovial fluid, many investigators have studied the effects of exogenous HA upon matrix metabolism. Addition of high-molecular-weight HA to the culture medium of cartilage explants blocks PG release caused by catabolic cytokines and fibronectin fragments [260–263]. Whether high-molecular-weight HA molecules can reduce in vivo the extent of cartilage matrix degradation in OA and other arthritides remains a matter for debate [264, 265].

Chapter 25 Products of Cartilage Metabolism

VII. OTHER PRODUCTS OF CHONDROCYTE METABOLISM The 35-kDa glycoprotein termed tumor necrosis factor stimulated gene-6 (TSG-6) is not found in normal joints but is markedly expressed by the synoviocytes and chondrocytes in both OA and RA joints [266]. Because, on the one hand, the link module present in its N-terminal domain binds to both HA and the G1 domain of aggrecan, and, on the other hand, the CUB module present in its C-terminus can bind to collagen-like structures, members of the TGF-β superfamily and GAGs as well, TSG-6 might cross-link components of the ECM [267–269]. On the other hand, TGS-6 might protect cartilage from extensive cartilage degradation, even in the presence of acute inflammation: by forming a covalent complex with the inter-αinhibitor (IαI), TSG-6 enhances 100-fold the inhibitory activity of IαI against plasmin which then becomes unable to activate latent MMPs, including MT-MMPs, which are apparently required for the activation of aggrecanase activity [270]. In response to partial oxygen pressure variations, mechanical stress and inflammatory mediators, chondrocytes produce abnormal levels of reactive oxygen species (ROS): nitric oxide (NO) and superoxide anion generate derivative radicals, including peroxynitrite and hydrogen peroxide (as reviewed in reference [271]). Chondrocytes constitutively express the nitric oxide synthase-3 (NOS-3), a low-output enzyme, but when stimulated by either bacterial lipopolysaccharides (LPS), and/or pro-inflammatory cytokines such as IL-1 and TNF-α, the cartilage cells synthesize an inducible isoform of nitric oxide synthase (iNOS, NOS-2) [271, 272], an induction that can be markedly inhibited by transforming growth factor-β, antiinflammatory cytokines such as IL-4, IL-10, and IL-13 [271, 273], as well as by nonsteroidal anti-inflammatory drugs [274]. Once induced, this iNOS generates large quantities of NO which sustains the nuclear translocation of NF-κB and thus keeps NF-κB-dependent transcription persistently switched on [275]. It has been estimated that chondrocyte-derived NO contributes significantly to the total NO produced in the joint cavity in inflammatory arthritides [276–280]. On the other hand, NO inhibits the enzyme complex NADPH-oxidase that produces superoxide anion radicals [281]. As chondrocytes also synthesize myeloperoxidase whose mRNA levels are enhanced in OA, the cartilage cells are likely to produce hypochlorous acid [282]. There is increasing evidence that cellular responses to ROS generation are dependent on the cellular redox status that results from a subtle equilibrium between ROS production and the intracellular antioxidant level. When they are produced at low levels in articular chondrocytes,

437 ROS are likely to contribute to the maintenance of cartilage homeostasis. In addition to the activation of different members of signaling cascades involved in cell growth and differentiation, ROS regulate the DNA-binding activity of transcription factors, and hence modulate gene expression [271]. On the other hand, when they are produced in greater amounts, ROS become deleterious for joint tissues: NO up-regulates the expression of genes coding for MMPs [283], activates latent MMPs [284], and inhibits the synthesis of both PGs and type II collagen [285, 286]. Adding strength to this contention are the results of in vivo studies pointing out that a specific inhibitor of iNOS reduces the severity of cartilage lesions in canine experimental OA [287], whereas the addition of anti-oxidative vitamins to the diet of STR/1R mice diminishes the development of mechanically induced OA [288]. Further studies conducted in iNOS knockout mice strongly suggest that, by inhibiting insulin-like growth factor-(IGF)-1 receptor autophosphorylation, NO might contribute to the resistance of arthritic chondrocytes to IGF-1 stimulation [289]. On the other hand, NO by itself cannot initiate chondrocyte apoptosis, the programmed cell death requiring the concomitant production of superoxide anion [290]. With aging and with the development of OA, 3nitrotyrosine, a product of the reaction of peroxynitrite with tyrosine residues in nearby proteins [291], accumulates within cartilage matrix and hereby provides further indirect evidence of the production of ROS in the articular tissue [292]. The potential mechanism of matrix degradation of ROS cannot be discounted. Even at low concentrations, ROS inhibit glycolysis and induce a rapid depletion in ATP; the depletion in ATP is further compounded by a direct nonenzymatic hydrolysis of ATP leading to a rapid increase in inorganic pyrophosphate (PPi) levels [293]. This effect of ROS on ATP depletion is reversible as long as the exposure of chondrocytes to ROS does not last more than 1–2 hours. Long-term exposure to ROS leads to peroxidation of membrane lipids and cell death. ROS also promote the degradation or denaturation of various cartilaginous matrix components. Thus, ROS depolymerize HA and can degrade collagen, LP and PG either directly or indirectly by activating latent proteinases and/or inactivating inhibitors of proteinase. Articular chondrocytes have the unique ability to constitutively elaborate large amounts of extracellular inorganic pyrophosphate (ePPi) whose levels play an important role in the formation of both calcium pyrophosphate dehydrate (CPPD) crystals and basic calcium phosphate (BCP) crystals, the latter being related to hydroxyapatite, carbonate-substituted apatite and octacalcium phosphate [294, 295]. Specifically, excess of ePPi promotes CPPD crystal formation whereas ePPi deficiency leads to hydroxyapatite crystal deposition. PPi itself is the byproduct of

438 many biosynthetic processes and biochemical reactions in the cell, and the direct product of nucleoside triphosphate pyrophosphohydrolases (NTPPPHs) [296]; it is degraded by several inorganic pyrophosphatases, including alkaline phosphatase. Because PPi does not readily diffuse across healthy biomembranes, levels of ePPi are controlled by the activity of chondrocyte ecto-NTPPHs, such as plasma cell membrane glycoprotein-1 (PC-1) and cartilage intermediate-layer protein (CILP), as well as by the activity of ANKH, a multipass transmembrane protein controlling the egress of iPPi and intracellular substrates for NTPPPHs [297]. Numerous soluble factors modulate ePPi production, as does aging. By enhancing the expression of PC-1 and ANK, TGF-β is unique in its capacity to markedly up-regulate extracellular elaboration of PPi by the chondrocytes [294, 298], a response that is further increased by epidermal growth factor but suppressed by IL-1, parathyroid hormone-related peptide and insulin-like growth factor [299]. Further, aged chondrocytes are more responsive to TGF-β than chondrocytes from young subjects [300]. On the other hand, transglutaminases (type II and factor XIIIA) activate latent TGF-β to increase chondrocyte ePPi production [301, 302], and, in turn, NO and pro-inflammatory cytokines induce increased chondrocyte transglutaminase activity [303]. The linkage between calcium-containing crystal deposition and changes in ePPi has been documented in several animal models and human diseases. In mice, the reduced levels of ePPi secondary to the recessive mutation of ANK leads to spontaneous hydroxyapatite crystal deposition in articular cartilage [304, 305] whereas, in humans, the dominant mutations of ANKH are associated with a slower increase in the levels of ePPi and with deposition of CPPD crystals in cartilage matrix [306, 307]. Hypophosphatasia, a deficiency of the tissue-nonspecific form of alkaline phosphatase, that hydrolyzes PPi to inorganic phosphate, also results in accumulation of ePPi and CPPD deposition in cartilage [308]. A member of the thrombospondin family termed cartilage oligomeric matrix protein (COMP), is an enigmatic pentameric protein that despite its high expression in cartilage has not been assigned a biological function (see Chapter 4). COMP binds collagens I/II and the noncollagenous domain of collagen IX, and, accordingly, this abundant cartilage matrix protein might have several roles in cartilage tissue homeostasis, including regulating collagen fibril formation and maintenance of the integrity and properties of the collagen network [309], a contention strengthened by the observation that mutations of the gene encoding COMP cause pseudo-achondroplasia and multiple epiphyseal dysplasia [310]. The rate of synthesis of

DANIEL-HENRI MANICOURT ET AL.

COMP is enhanced in human cartilage with early OA lesions [311]. In patients with RA and OA, serum COMP levels of COMP could reflect the extent of cartilage matrix turnover [312–315]. As observed for other biomarkers of cartilage metabolism [316], the serum levels of COMP are increased after physiological cyclic loading [317]. Several components of the complement system, including C1q, have been reported to be secreted by normal and OA chondrocytes but their potential roles in matrix homeostasis and chondrocyte metabolism are still largely hypothetical [318]. Likewise, nothing is known about the possible function(s) of a glycoprotein termed YLK-40 that appears to be secreted by both chondrocytes and synoviocytes from patients with OA and RA [319]. Although IL-1 and TGF-β both suppress markedly YKL-40 production by articular chondrocytes [319], sustained enhanced levels of the glycoprotein are apparently related to progression of joint destruction in early RA patients [320]. By interacting with high affinity with decorin, the cartilage-specific (V+C)− fibronectin (Fn) isoform not only inhibits PG degradation by binding ADAMTS4 [181], but also might help stabilize the differentiated phenotype of chondrocytes [321]. Fn synthesis is selectively affected by the frequency of intermittent loads applied to cartilage [322]. In both OA and RA, the cartilage matrix exhibits a 10–20-fold increase in content of this ubiquitous protein [323]. Although TGF-β markedly increases Fn synthesis by normal chondrocytes, this growth factor alone is not sufficient to explain such an accumulation in diseased cartilage; therefore, either intact or degraded Fn molecules are likely to originate from the synovial fluid. In contrast to intact molecules, Fn fragments (fn-f) markedly affect the metabolism of chondrocytes. Indeed, at the high concentrations found in synovial fluids from diseased joints, these fragments can bind to the alpha[5]beta[1] integrin receptor subunit of chondrocytes [324] and up-regulate cartilage catabolism: they stimulate the liberation of IL-1, up-regulate iNOS [325], and cause not only the loss of PGs but also induce the release and/or degradation of COMP and chondroadherin [326]. It is most interesting that, at lower concentrations, the same fragments have an anabolic effect, thought to be mediated by IGF-1, upon matrix metabolism [327].

VIII. CONCLUDING STATEMENT The cartilage matrix contains several molecules that are found in much higher concentrations than in other tissues. The attention that aggrecan and collagen type II are currently receiving as potential markers of specific metabolic alterations in cartilage disease will likely extend to

Chapter 25 Products of Cartilage Metabolism

less abundant molecules found to play key roles in the maintenance of matrix homeostasis. As our understanding of the function and metabolism of these molecules improves, so will their potential as markers of the irreversible destabilization of the collagenous meshwork or of other specific alterations in cartilage matrix organization.

Acknowledgments The preparation of the manuscript was supported in part by the FSR of the UCL.

References 1. Atencia, L. J., McDevitt, C. A., Nile, W. B., and Sokoloff, L. (1989). Cartilage content of an immature dog. Connect Tissue Res. 18, 235–242. 2. Poole, A. R. (2005). Cartilage in health and disease. In “Arthritis and Allied Conditions. A Textbook of Rheumatology” (15th ed) (W. J. Koopman and L. W. Moreland, eds), pp. 223–269, Lippincott Williams and Wilkins Philadelphia, PA. 3. Mow, V. C., and Ratcliffe, A. (1997). Structure and function of articular cartilage and meniscus. In “Basic Orthopaedic Biomechanics” (2nd ed) (V. C. Mow and W. C. Hayes, eds), pp. 113–177, Lippincott-Raven Press Publishers, Philadelphia, PA. 4. Mow, V. C., Wang, C. C., and Hung, C. T. (1999). The extracellular matrix, interstitial fluid and ions as a mechanical signal transducer in articular cartilage. Osteoarthritis Cartilage 7, 41–58. 5. Hing, W. A., Sherwin, A. F., and Poole, C. A. (2002). The influence of the pericellular microenvironment on the chondrocyte response to osmotic challenge. Osteoarthritis Cartilage 10, 297–307. 6. Horikawa, O., Nakajima, H., Kikuchi, T., Ichimura, S., Yamada, H., Fujikawa, K., and Toyama, Y. (2004). Distribution of type VI collagen in chondrocyte microenvironment: study of chondrons isolated from human normal and degenerative articular cartilage and cultured chondrocytes. J. Orthop. Sci. 9, 29–36. 7. Roughley, P. J. (2001). Articular cartilage and changes in arthritis: noncollagenous proteins and proteoglycans in the extracellular matrix of cartilage. Arthritis Res. 3, 342–347. 8. Schwartz, N. B., and Domowicz, M. (2002). Chondrodysplasias due to proteoglycan defects. Glycobiology 12, 57R–68R. 9. Grimshaw, M. J., and Mason, R. M. (2001). Modulation of bovine articular chondrocyte gene expression in vitro by oxygen tension. Osteoarthritis Cartilage 9, 357–364. 10. Domm, C., Schunke, M., Christesen, K., and Kurz, B. (2002). Redifferentiation of dedifferentiated bovine articular chondrocytes in alginate culture under low oxygen tension. Osteoarthritis Cartilage 10, 13–22. 11. Semenza, G. L. (2001). Hypoxia-inducible factor 1: oxygen homeostasis and disease pathophysiology. Trends Mol. Med. 7, 345–350. 12. Coimbra, I. B., Jimenez, S. A., Hawkins, D. F., Piera-Velazquez, S., and Stokes, D.G. (2004). Hypoxia inducible factor-1 alpha expression in human normal and osteoarthritic chondrocytes. Osteoarthritis Cartilage 12, 336–345. 13. Mason, R. M., and Sweeney, C. (1994). The relationship between proteoglycan synthesis in Swarm chondrocytes and pathways of cellular energy and UDP-sugar metabolism. Carbohydrate Res. 255, 255–270. 14. Lee, R. B., and Urban, J. P. (2002). Functional replacement of oxygen by other oxidants in articular cartilage. Arthritis Rheum. 46, 3190–3200.

439 15. Lee, R. B., Wilkins, R. J., Razaq, S., and Urban, J. P. (2002). The effect of mechanical stress on cartilage energy metabolism. Biorheology 39, 133–143. 16. Browning, J. A., and Wilkins, R. J. (2004). Mechanisms contributing to intracellular pH homeostasis in an immortalised human chondrocyte cell line. Comp. Biochem. Physiol. A. Mol. Integr. Physiol. 137, 409–418. 17. Urban, J. P. G., Hall, A. C., and Ghel, K. A. (1993). Regulation of matrix synthesis rates by the ionic and osmotic environment of articular chondrocytes. J. Cell. Physiol. 154, 262–270. 18. Hall, A. C., Starks, I., Shoults, C. L., and Rashidbigi, S. (1996). Pathways for K+ transport across the bovine articular chondrocyte membrane and their sensitivity to cell volume. Am. J. Physiol. 270, (5 Pt 1), C1300–C1310. 19. Hopewell, B., and Urban, J. P. (2003). Adaptation of articular chondrocytes to changes in osmolality. Biorheology 40, 73–77. 20. Aydelotte, M. B., and Kuettner, K. E. (1993). Heterogeneity of articular chondrocytes and cartilage matrix. In “Joint Cartilage Degradation. Basic and Clinical Aspects” (J. F. Woessner, Jr. and D. S. Howell, eds), pp. 37–65, M. Dekker Inc., New York. 21. Little, C. B., Ghosh, P., and Bellenger, C. R. (1996). Topographic variation in biglycan and decorin synthesis by articular cartilage in the early stages of osteoarthritis: an experimental study in sheep. J. Orthop. Res. 14, 433–444. 22. Kim, Y. J., Grodzinsky, A. J., and Plaas, A. H. (1996). Compression of cartilage results in differential effects on biosynthetic pathways for aggrecan, link protein, and hyaluronan. Arch. Biochem. Biophys. 328, 331–340. 23. Wong, M., Siegrist, M., and Cao, X. (1999). Cyclic compression of articular cartilage explants is associated with progressive consolidation and altered expression pattern of extracellular matrix proteins. Matrix Biol. 18, 391–399. 24. Jin, M., Frank, E. H., Quinn, T. M., Hunziker, E. B., and Grodzinsky, A. J. (2001). Tissue shear deformation stimulates proteoglycan and protein biosynthesis in bovine cartilage explants. Arch. Biochem. Biophys. 395, 41–48. 25. Sauerland, K., Raiss, R. X., and Steimeyer, J. (2003). Proteoglycan metabolism and viability of articular cartilage explants as modulated by the frequency of intermittent loading. Osteoarthritis Cartilage 11(5), 343–350. 26. Muller, F. J., Setton, L. A., Manicourt, D. H., Mow, V. C., Howell, D. S., and Pita, J. C. (1994). Centrifugal and biochemical comparison of proteoglycan aggregates from articular cartilage in experimental joint disuse and joint instability. J. Orthop. Res. 12, 498–508. 27. Grumbles, R. M., Howell, D. S., Howard G. A., Roos, B. A., Setton, L. A., Mow, V. C., Ratcliffe A, Muller, F. J., and Altman, R. D. (1995). Cartilage metalloproteases in disuse atrophy. J. Rheumatol. Suppl. 43, 146–148. 28. Deschner, J., Hofman, C. R., Piesco, N. P., and Agarwal, S. (2003). Signal transduction by mechanical strain in chondrocytes. Curr. Opin. Clin. Nutr. Metab. Care 6, 289–293. 29. Borrelli, J. Jr, Tinsley, K., Ricci, W. M., Burns M., Karl, I. E., and Hotchkiss R. (2003). Induction of chondrocyte apoptosis following impact load. J. Orthop. Trauma. 17, 635–641. 30. Chen, C. T., Bhargava, M., Lin, P. M., and Torzilli, P. A. (2003). Time, stress, and location dependent chondrocyte death and collagen damage in cyclically loaded articular cartilage. J. Orthop. Res. 21, 888–898. 31. Kuhn, K., D’Lima, D. D., Hashimoto, S., and Lotz, M. (2004). Cell death in cartilage. Osteoarthritis Cartilage 12, 1–16. 32. Kurz, B., Jin, M., Patwari, P., Cheng, D. M., Lark, M. W., and Grodzinsky, A. J. (2001). Biosynthetic response and mechanical properties of articular cartilage after injurious compression. J. Orthop. Res. 19, 1140–1146. 33. Wilkins, R. J., Browning, J. A., and Urban, J. P. (2000). Chondrocyte regulation by mechanical load. Biorheology 37(1-2), 67–74. 34. Lee, R. B., Wilkins, R. J., Razaq, S., and Urban, J. P. (2002). The effect of mechanical stress on cartilage energy metabolism. Biorheology 39, 133–143.

440 35. Mow, V. C., Wang, C. C., and Hung, C. T. (1999). The extracellular matrix, interstitial fluid and ions as a mechanical signal transducer in articular cartilage. Osteoarthritis Cartilage 7, 41–58. 36. Guilak, F., and Mow, V. C. (2000). The mechanical environment of the chondrocyte: a biphasic finite element model of cell-matrix interactions in articular cartilage. J. Biomech. 33, 1663–1673. 37. Sun, D. D., Guo, X. E., Likhitpanichkul, M., Lai, W. M., and Mow, V. C. (2004). The influence of the fixed negative charges on mechanical and electrical behaviors of articular cartilage under unconfined compression. J. Biomech. Eng. 126, 6–16. 38. Vincent, T. L., Hermannsson, M. A., Hansen, U. N., Amis, A. A., and Saklatvala, J. (2004). Basic fibroblast growth factor mediates transduction of mechanical signals when articular cartilage is loaded. Arthritis Rheum. 50, 526–533. 39. Pearson, G., Robinson, F., Beers Gibson, T., Xu, B. E., Karandikar, M., Berman, K., and Cobb, M. H. (2001). Mitogen-activated protein (MAP) kinase pathways: regulation and physiological functions. Endocr. Rev. 22, 153–183. 40. Lee, D. A., Noguchi, T., Frean, S. P., Lees, P., and Bader, D. L. (2000). The influence of mechanical loading on isolated chondrocytes seeded in agarose constructs. Biorheology 37(1-2), 149–161. 41. Hayashi, T., Abe, E., Yamate, T., Taguchi, Y., and Jasin, H. (1997). Nitric oxide production by superficial and deep articular chondrocytes. Arthritis Rheum. 40, 261–269. 42. Hauselmann, H. J., Stefanovic-Racic, M., Michel, B. A., and Evans, C. H. (1998). Differences in nitric oxide production by superficial and deep human articular chondrocytes: implications for proteoglycan turnover in inflammatory joint diseases. J. Immunol. 160, 1444–1448. 43. Poole, A. R. (1992). Immunology of Cartilage. In “Osteoarthritis: Diagnosis and Medical / Surgical Management” (R. Moskowitz, D. S. Howell, V. M. Goldberg and, H. J. Mankin, eds), pp. 155–189, W. B. Saunders Company, Philadelphia, PA. 44. Ryan, M. C., and Sandell, L. J. (1990). Differential expression of a cysteine-rich domain in the amino-terminal propeptide of type II (cartilage) procollagen by alternative splicing of mRNA. J. Biol. Chem. 265, 10334–10339. 45. Aigner, T., Zhu, Y., Chansky, H. H., Matsen, F.A. 3rd., Maloney, W. J., and Sandell, L. J. (1999). Reexpression of type IIA procollagen by adult articular chondrocytes in osteoarthritic cartilage. Arthritis Rheum. 42, 1443–1450. 46. Nah, H. D., Swoboda, B., Birk, D. E., and Kirsch, T. (2001). Type IIA procollagen: expression in developing chicken limb cartilage and human osteoarthritic articular cartilage. Dev. Dyn. 220, 307–322. 47. Aurich, M., Poole, A. R., Reiner, A., Mollenhauer, C., Margulis, A., Kuettner, K. E., and Cole, A. A. (2002). Matrix homeostasis in aging normal human ankle cartilage. Arthritis Rheum. 46, 2903–2910. 48. Fernandes, R. J., Hirohata, S., Engle, J. M., Colige A., Cohn, D. H., Eyre, D. R., and Apte, S. S. (2001). Procollagen II amino propeptide processing by ADAMTS-3. Insights on dermatosparaxis. J. Biol. Chem. 276, 31502–31509. 49. Li, S-W., Sieron, A. L., Fertala, A., Hojima, Y., Arnold, W. V., and Prockop, D. J. (1996). The C-proteinase that processes procollagens to fibrillar collagens is identical to the protein previously identified as bone morphogenic protein-1. Proc. Natl Acad. Sci. USA 93, 5127-5130. 50. Rebuck, N., Croucher, L. J., and Hollander, A. P. (1999). Distribution of two alternatively spliced variants of the type II collagen N-propeptide compared with the C-propeptide in bovine chondrocyte pellet cultures. J. Cell. Biochem. 75, 13–21. 51. Nelson, F., Dahlberg, L., Laverty, S., Reiner, A., Pidoux, I., Ionescu, M., Fraser, G. L., Brooks, E., Tanzer, M., Rosenberg, L. C., Dieppe, P., and Robin Poole, A. (1998). Evidence for altered synthesis of type II collagen in patients with osteoarthritis. J. Clin. Invest. 102, 2115–2125. 52. Garnero, P., Ayral, X., Rousseau, J. C., Christgau, S., Sandell, L. J., Dougados, M., and Delmas, P. D. (2002). Uncoupling of type II collagen

DANIEL-HENRI MANICOURT ET AL.

53.

54.

55.

56.

57.

58.

59. 60.

61.

62.

63. 64.

65.

66. 67.

68.

69.

synthesis and degradation predicts progression of joint damage in patients with knee osteoarthritis. Arthritis Rheum. 46, 2613–2624. Carey, D. E., Alini, M., Ionescu, M., Hyams, J. S., Rowe, J. C., Rosenberg, L. C., and Poole, A. R. (1997). Serum content of the C-propeptide of the cartilage molecule type II collagen in children. Clin. Exp. Rheumatol. 15, 325–328. Shinmei, M., Ito, K., Matsuyama, S., Yoshihara, Y., and Matsuzawa, K. (1993). Joint fluid carboxy-terminal type II procollagen peptide as a marker of cartilage collagen biosynthesis. Osteoarthritis Cartilage 1, 121–128. Lohmander, L. S., Yoshihara, Y., Roos, H., Kobayashi, T., Yamada, H., and Shinmei, M. (1996). Procollagen II C-propeptide in joint fluid: changes in concentration with age, time after knee injury, and osteoarthritis. J. Rheumatol. 23, 1765–1769. Kobayashi, T., Yoshihara, Y., Samura, A., Yamada, H., Shinmei, M., Roos, H., and Lohmander, L. S. (1997). Synovial fluid concentrations of the C-propeptide of type II collagen correlate with body mass index in primary knee osteoarthritis. Ann. Rheum. Dis. 56, 500–503. Fraser, A., Fearon, U., Billinghurst, R. C., Ionescu, M., Reece, R., Barwick, T., Emery, P., Poole, A. R., and Veale, D. J. (2003). Turnover of type II collagen and aggrecan in cartilage matrix at the onset of inflammatory arthritis in humans: relationship to mediators of systemic and local inflammation. Arthritis Rheum. 48, 3085–3095. Mansson, B., Carey, D., Alini, M., Ionescu, M., Rosenberg, L. C., Poole, A. R., Heinegard, D., and Saxne, T. (1995). Cartilage and bone metabolism in rheumatoid arthritis. Differences between rapid and slow progression of disease identified by serum markers of cartilage metabolism. J. Clin. Invest. 95, 1071–1077. Eyre, D. (2002). Collagen of articular cartilage. Arthritis Res. 4, 30–35. Verzijl, N., DeGroot, J., Ben, Z. C., Brau-Benjamin, O., Maroudas, A., Bank, R. A., Mizrahi, J., Schalkwijk, C. G., Thorpe, S. R., Baynes, J. W., Bijlsma, J. W., Lafeber, F. P., and TeKoppele, J. M. (2002). Crosslinking by advanced glycation end products increases the stiffness of the collagen network in human articular cartilage: a possible mechanism through which age is a risk factor for osteoarthritis. Arthritis Rheum. 46, 114–123. DeGroot, J., Verzijl, N., Wenting-van Wijk, M. J., Jacobs, K. M., Van El, B., Van Roermund, P. M., Bank, R. A., Bijlsma, J. W., TeKoppele, J. M., and Lafeber, F. P. (2004). Accumulation of advanced glycation end products as a molecular mechanism for aging as a risk factor in osteoarthritis. Arthritis Rheum. 50, 1207–1215. Billinghurst, R. C., Mwale, F., Hollander, A., Ionescu, M., and Poole, A. R. (2004). Immunoassays for collagens in chondrocyte and cartilage explant cultures. Meth. Mol. Med. 100, 251–274. Barrett, A. J. (1978). The possible role of neutrophil proteinases in damage to articular cartilage. Agents Actions 8, 15–18. Wu, J-J., Lark, M. W., Chun, L. E., and Eyre, D .R. (1991). Sites of stromelysin cleavage in collagen, II, IX, X and, XI. J. Biol. Chem. 266, 5625–5628. Kozaci, L. D., Buttle, D. J., and Hollander, A. P. (1997). Degradation of type II collagen, but not proteoglycan correlates with matrix metalloproteinase activity in cartilage explant cultures. Arthritis Rheum. 40, 164–174. Nagase, H. (1997). Activation mechanisms of matrix metalloproteinases. Biol. Chem. 378, 151–160. van Meurs, J., van Lent, P., Holthuysen, A., Lambrou, D., Bayne, E., Singer, I., and van den Berg W. (1999). Active matrix metalloproteinases are present in cartilage during immune complex-mediated arthritis: a pivotal role for stromelysin-1 in cartilage destruction. J. Immunol. 163, 5633–5639. Poole, A. R., Nelson, F., Dahlberg, L., Tchetina, E., Kobayashi, M., Yasuda, T., Laverty, S., Squires, G., Kojima, T., and Wu, W., and Billinghurst, R. C. (2003). Proteolysis of the collagen fibril in osteoarthritis. Biochem. Soc. Symp. 70, 115–123. Hollander, A. P., Pidoux, I., Reiner, A., Rorabeck, C., Bourne, R., and Poole, A. R. (1995). Damage to type II collagen in aging and

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70.

71.

72.

73.

74.

75.

76.

77.

78.

79.

80.

81.

82.

83.

osteoarthritis starts at the articular surface, originates around chondrocytes, and extends into the cartilage with progressive degeneration. J. Clin. Invest. 96, 2859–2869. Wu, W., Billinghurst, R. C., Pidoux, I., Antoniou, J., Zukor, D., Tanzer, M., and Poole, A. R. (2002). Sites of collagenase cleavage and denaturation of type II collagen in aging and osteoarthritic articular cartilage and their relationship to the distribution of matrix metalloproteinase 1 and matrix metalloproteinase 13. Arthritis Rheum. 46, 2087–2094. Aurich, M., Poole, A. R., Reiner, A., Mollenhauer, C., Margulis, A., Kuettner, K. E., and Cole, A. A. (2002). Matrix homeostasis in aging normal human ankle cartilage. Arthritis Rheum. 46, 2903–2910. Dodge, G. R., and Poole, A. R. (1989). Immunohistochemical detection and immunochemical analysis of type II collagen degradation in human normal, rheumatoid and osteoarthritic articular cartilages and in explants of bovine articular cartilage cultured with interleukin-1. J. Clin. Invest. 83, 647–661. Billinghurst, R. C., Dahlberg, L., Ionescu, M., Reiner, A., Bourne, R., Rorabeck, C., Mitchell, P., Hambor, J., Diekmann, O., Tschesche, H., Chen, J., Van Wart, H., and Poole, A. R. (1997). Enhanced cleavage of type II collagen by collagenases in osteoarthritic articular cartilage. J. Clin. Invest. 99, 1534–1545. Knauper, V., Lopez-Otin, C., Smith, B., Knight, G., and Murphy, G. (1996). Biochemical characterization of human collagenase-3. J. Biol. Chem. 271, 1544–1550. Dahlberg, L., Billinghurst, R. C., Manner, P., Nelson, F., Webb, G., Ionescu, M., Reiner, A., Tanzer, M., Zukor, D., Chen, J., van Wart, H. E., and Poole, A. R. (2000). Selective enhancement of collagenase-mediated cleavage of resident type II collagen in cultured osteoarthritic cartilage and arrest with a synthetic inhibitor that spares collagenase 1 (matrix metalloproteinase 1). Arthritis Rheum. 43, 673–682. Billinghurst, R. C., Wu, W., Ionescu, M., Reiner, A., Dahlberg, L., Chen, J., van Wart, H., and Poole, A. R. (2000). Comparison of the degradation of type II collagen and proteoglycan in nasal and articular cartilages induced by interleukin-1 and the selective inhibition of type II collagen cleavage by collagenase. Arthritis Rheum. 43, 664–672. Bau, B., Gebhard, P. M., Haag, J., Knorr, T., Bartnik, E., and Aigner, T. (2002). Relative messenger RNA expression profiling of collagenases and aggrecanases in human articular chondrocytes in vivo and in vitro. Arthritis Rheum. 46, 2648–2657. Neuhold, L. A., Killar, L., Zhao, W., Sung, M. L., Warner, L., Kulik, J., Turner, J., Wu, W., Billinghurst, C., Meijers, T., Poole, A. R., Babij, P., and DeGennaro, L. J. (2001). Postnatal expression in hyaline cartilage of constitutively active human collagenase-3 (MMP-13) induces osteoarthritis in mice. J. Clin. Invest. 107, 35–44. Wu, C. W., Tchetina, E. V., Mwale, F., Hasty, K., Pidoux, I., Reiner, A., Chen, J., Van Wart, H. E., and Poole, A. R. (2002). Proteolysis involving matrix metalloproteinase 13 (collagenase-3) is required for chondrocyte differentiation that is associated with matrix mineralization. J. Bone Miner. Res. 17, 639–651. Murphy, G., and Reynolds, J. J. (1993). Extracellular matrix degradation. In “Connective Tissue and Its Heritable Disorders. Molecular Genetic and Medical Aspects” (P. M. Royce and B. Steinmann, eds), pp. 287–316, Wiley-Liss. New York. Kafienah, W., Bromme, D., Buttle, D. J., Croucher, L. J., and Hollander, A. P. (1998). Human cathepsin K cleaves native type I and II collagens at the N-terminal end of the triple helix. Biochem. J. 331 (Pt 3), 727–732. Price, J. S., Wang-Weigand, S., Bohne, R., Kozaci, L. D., and Hollander, A. P. (1999). Retinoic acid-induced type II collagen degradation does not correlate with matrix metalloproteinase activity in cartilage explant cultures. Arthritis Rheum. 42, 137–147. Brown, N. L., Knott, L., Halligan, E., Yarram, S. J., Mansell, J. P., and Sandy, J. R. (2003). Microarray analysis of murine palatogenesis: temporal expression of genes during normal palate development. Dev. Growth. Differ. 45, 153–165.

441 84. Matyas, J. R., Atley, L., Ionescu, M., Eyre, D. R., and Poole, A. R. (2004). Analysis of cartilage biomarkers in the early phases of canine experimental osteoarthritis. Arthritis Rheum. 50, 543–552. 85. Kojima, T., Mwale, F., Yasuda, T., Girard, C., Poole, A. R., and Laverty, S. (2001). Early degradation of type IX and type II collagen with the onset of experimental inflammatory arthritis. Arthritis Rheum. 44, 120–127. 86. Cerejo, R., Poole, A. R., Ionescu, M., Lobanok, T., Song, J., Cahue, S., Dunlop, D., and Sharma, L. (2002). The association between cartilage collagenase activity measured in serum and progression of knee osteoarthritis in patients with and without evidence of generalized disease. Arthritis Rheum. 48, (suppl 9), S144. 87. Henrotin, Y., Deberg, M., Dubuc, J. E., Quettier, E., Christgau, S., and Reginster, J. Y. (2004). Type II collagen peptides for measuring cartilage degradation. Biorheology 41, 543–547. 88. Deberg, M., Labasse, A., Christgau, S., Cloos, P., Bang Henriksen, D., Chapelle, J. P., Zegels, B., Reginster, J. Y., and Henrotin, Y. (2005). New serum biochemical markers (Coll 2-1 and Coll 2-1 NO2) for studying oxidative-related type II collagen network degradation in patients with osteoarthritis and rheumatoid arthritis. Osteoarthritis Cartilage 13, 258–265. 89. Farrell, A. J., Blake, D. R., Palmer, R. M., and Moncada, S. (1992). Increased concentrations of nitrite in synovial fluid and serum samples suggest increased nitric oxide synthesis in rheumatic diseases. Ann. Rheum. Dis. 51, 1219–1222. 90. Hiran, T. S., Moulton, P. J., and Hancock, J.T. (1998). In situ detection of superoxide anions within porcine articular cartilage. Br. J. Biomed. Sci. 55, 199–203. 91. Henrotin, Y. E., Zheng, S. X., Deby, G. P., Labasse, A. H., Crielaard, J. M., and Reginster, J. Y. (1998). Nitric oxide downregulates interleukin 1beta (IL-1beta) stimulated IL-6, IL-8, and prostaglandin E2 production by human chondrocytes. J. Rheumatol. 25, 1595–1601. 92. Hashimoto, S., Ochs, R. L., Komiya, S., and Lotz, M. (1998). Linkage of chondrocyte apoptosis and cartilage degradation in human osteoarthritis. Arthritis Rheum. 41, 1632–1638. 93. Burkhardt, H., Koller, T., Engstrom, A., Nandakumar, K. S., Turnay, J., Kraetsch, H. G., Kalden, J. R., and Holmdahl, R. (2002). Epitopespecific recognition of type II collagen by rheumatoid arthritis antibodies is shared with recognition by antibodies that are arthritogenic in collagen-induced arthritis in the mouse. Arthritis Rheum. 46, 2339–2348. 94. Mwale, F., Billinghurst, C., Wu, W., Alini, M., Webber, C., Reiner, A., Ionescu, M., Poole, J., and Poole, A. R. (2000). Selective assembly and remodelling of collagens II and IX associated with expression of the chondrocyte hypertrophic phenotype. Dev. Dyn. 218, 648–662. 95. Sinigaglia, L., Varenna, M., Binelli, L., Bartucci, F., Arrigoni, M., Ferrara, R., and Abbiati, G. (1995). Urinary and synovial pyridinium crosslink concentrations in patients with rheumatoid arthritis and osteoarthritis. Ann. Rheum. Dis. 54, 144–147. 96. Black, D., Marabani, M., Sturrock, R. D., and Robins, S. P. (1989). Urinary excretion of hydroxypyridinium crosslinks of collagen in patients with rheumatoid arthritis. Ann. Rheum. Dis. 48, 641–644. 97. Seibel, M. J., Duncan, A., and Robins, S. P., (1989). Urinary hydroxypyridinium crosslinks provide indices of cartilage and bone involvement in arthritic diseases. J. Rheumatol. 16, 964–970. 98. Astbury, C., Bird, H. A., McLaren, A. M., and Robins, S. P. (1994). Urinary excretion of pyridinium crosslinks of collagen correlated with joint damage in arthritis. Br. J. Rheumatol. 33, 11–15. 99. Eyre, D. R., Dickson, I. R., and Van Ness, K. P. (1988). Collagen cross-linking in human bone and articular cartilage. Age-related changes in the content of mature hydroxypyridinium residues. Biochem. J. 252, 495–500. 100. MacDonald, A. G., McHenry, P., Robins, S. P., and Reid, D. M. (1994). Relationship of urinary pyridinium crosslinks to disease extent and activity in osteoarthritis. Br. J. Rheumatol. 33, 16–19.

442 101. Gough, A. K. S., Peel, N. F. A., Eastell, R., Holder, R. L., Lilley, J., and Emery, P. (1994). Excretion of pyridinium crosslinks correlates with disease activity and appendicular bone loss in early rheumatoid arthritis. Ann. Rheum. Dis. 53, 14–17. 102. Garnero, P., and Delmas, P. D. (2003). Biomarkers in osteoarthritis. Curr. Opin. Rheumatol. 15, 641–646. 103. Eyre, D. R., Atley, L. M., Shao, P., Vosberg-Smith, K., Ochs, V., and Clemens, J. D. (2000). Biochemical markers of bone and cartilage collagen degradation. In “Chemistry and Biology of Mineralized Tissues” (M. Goldberg, A. Boskey, and C. Robinson, eds), pp. 347–350, American Academy of Orthopedic Surgeons, Rosemoat, IL. 104. Christgau, S., Garnero, P., Fledelius, C., Moniz, C., Ensig, M., Gineyts, E., Rosenquist, C., and Qvist, P. (2001). Collagen type II C-telopeptide fragments as an index of cartilage degradation. Bone 29, 209–215. 105. Atley, L. M., Carlson, K., Pietka, T. A., and Eyre, D. R. (2001). Oncostatin M induces collagen degradation in human articular cartilage. Osteoarthritis Research Society International, Washington DC. 106. Lohmander, L. S., Atley, L. M., Pietka, T. A., and Eyre, D. R. (2003). The release of crosslinked peptides from type II collagen into human synovial fluid is increased soon after joint injury and in osteoarthritis. Arthritis Rheum. 48, 3130–3139. 107. Reijman, M., Hazes, J. M., Bierma-Zeinstra, S. M., Koes, B. W., Christgau, S., Christiansen, C., Uitterlinden, A. G., and Pols, H. A. (2004). A new marker for osteoarthritis: cross-sectional and longitudinal approach. Arthritis Rheum. 50, 2471–2478. 108. Landewe, R., Geusens, P., Boers, M., van der Heijde, D., Lems, W., te Koppele, J., van der Linden, S., and Garnero, P. (2004). Markers for type II collagen breakdown predict the effect of disease-modifying treatment on long-term radiographic progression in patients with rheumatoid arthritis. Arthritis Rheum. 50, 1390–1399. 109. Mok, S. S., Masuda, K., Hauselmann, H. J., Aydelotte, M. B., and Thonar, E. J. (1994). Aggrecan synthesized by mature bovine chondrocytes suspended in alginate. Identification of two distinct metabolic matrix pools. J. Biol. Chem. 269, 33021–33027. 110. Heinegard, D., Lorenzo, P., and Sommarin, Y. (1995). Articular cartilage matrix proteins. In “Osteoarthritic Disorders” (K. E. Kuettner and V. M. Goldberg, eds), pp. 229–237. American Academy of Orthopedic Surgeons Rosemont, LI. 111. Maroudas, A., Bayliss, M. T., Uchitel-Kaushansky, N., Schneiderman, R., and Gilav, E. (1998). Aggrecan turnover in human articular cartilage: use of aspartic acid racemization as a marker of molecular age. Arch. Biochem. Biophys. 350, 61–71. 112. Ritz-Timme, S., and Collins, M. J. (2002). Racemization of aspartic acid in human proteins. Ageing Res. Rev. 1, 43–59. 113. van Kampen, G. P., van de Stadt, R. J., van de Laar, M. A. F. J., and van der Korst, J. K. (1992). Two distinct metabolic pools of proteoglycans in articular cartilage In “Articular Cartilage and Osteoarthritis” (K. E. Kuettner, R. Schleyerbach, J. G. Peyron, V. C. Hascall, eds), pp. 281–289, Raven Press, New York. 114. Bayliss, M. T. (1992). Metabolism of animal and human osteoarthritic cartilage. In “Articular Cartilage and Osteoarthritis” (K. E. Kuettner, R. Schleyerbach, J. G. Peyron, V. C. Hascall, eds), pp. 487–500, Raven Press, New York. 115. Hauselmann, H. J., Masuda, K., Hunziker, E. B., Neidhart, M., Mok, S. S., Michel, B. A., and Thonar, E. J. (1996). Adult human chondrocytes cultured in alginate form a matrix similar to native human articular cartilage. Am. J. Physiol. 271 (3 Pt 1), C742–C752. 116. Ilic, M. Z., Robinson, H. C., and Handley, C. J. (1998). Characterization of aggrecan retained and lost from the extracellular matrix of articular cartilage. Involvement of carboxyl-terminal processing in the catabolism of aggrecan. J. Biol. Chem. 273, 17451–17458. 117. Loulakis, P., Shrikhande, A., Davis, G., and Maniglia, C. A. (1992). N-terminal sequence of proteoglycan fragments isolated from

DANIEL-HENRI MANICOURT ET AL.

118.

119.

120.

121.

122.

123.

124.

125.

126.

127.

128.

129.

130.

131.

medium of interleukin-1-treated articular-cartilage cultures. Putative site(s) of enzymic cleavage. Biochem. J. 284 (Pt 2), 589–593. Tortorella, M., Pratta, M., Liu, R. Q., Abbaszade, I., Ross, H., Burn, T., and Arner, E. (2000). The thrombospondin motif of aggrecanase-1 (ADAMTS-4) is critical for aggrecan substrate recognition and cleavage. J. Biol. Chem. 275, 25791–25797. Tortorella, M. D., Malfait, A. M., Deccico, C., and Arner, E. (2001). The role of ADAM-TS4 (aggrecanase-1) and ADAM-TS5 (aggrecanase-2) in a model of cartilage degradation. Osteoarthritis Cartilage 9, 539–552. Ratcliffe, A. M., Doherty, M., Maini, R. N., and Hardingham, T. E. (1988). Increased levels of proteoglycan components in the synovial fluids of patients with acute joint disease. Ann. Rheum. Dis. 47, 826–832. Caroll, G. (1989). Measurement of sulfated glycosaminoglycans and proteoglycan fragments in arthritic synovial fluid. Ann. Rheum. Dis. 48, 17–24. Thonar, E. J-M. A., Manicourt, D. H., Williams, J. M., Fukuda, K., Campion, G. V., Sweet, B. M. E., Lenz, M. E., Schnitzer, T. J., and Kuettner, K.E. (1992). Serum keratan sulfate: a measure of cartilage proteoglycan metabolism. In “Articular Cartilage and Osteoarthritis” (K. E. Kuettner, R. Schleyerbach, J. G. Peyron, V. C. Hascall, eds), pp. 429–445, Raven Press, New York. Dahlberg, L., Roos, H., Saxne, T., Heinegard, D., Lark, M. W., Hoerrner, L. A., and Lohmander, L. S. (1994). Cartilage metabolism in the injured and uninjured knee of the same patient. Ann. Rheum. Dis. 53, 823–827. Caterson, B., Flannery, C. R., Hughes, C. E., and Little, C. B. (2000). Mechanisms involved in cartilage proteoglycan catabolism. Matrix Biol. 19, 333–344. Elsaid, K. A., Jay, G. D., and Chichester, C. O. (2003). Detection of collagen type II and proteoglycans in the synovial fluids of patients diagnosed with non-infectious knee joint synovitis indicates early damage to the articular cartilage matrix. Osteoarthritis Cartilage 11, 673–680. Sandy, J. D., Neame, P. J., Boynton, R. E., and Flannery, C. R. (1991). Catabolism of aggrecan in cartilage explants. identification of a major cleavage site within the interglobular domain. J. Biol. Chem. 266, 8683–8685. Lohmander, L. S., Neame, P. J., and Sandy, J. D. (1993). The structure of aggrecan fragments in human synovial fluid. Evidence that aggrecanase mediates cartilage degradation in inflammatory joint disease, joint injury and osteoarthritis. Arthritis Rheum. 36, 1214–1222. Hughes, C. E., Caterson, B., Fosang, A. J., Roughley P. J., and Mort, J. S. (1995). Monoclonal antibodies that specifically recognize neoepitope sequences generated by “aggrecanase” and matrix metalloproteinase cleavage of aggrecan: application to catabolism in situ and in vitro. Biochem. J. 305, 799–804. Lark, M. W., Williams, H., Hoernner, L. A., Weidner, J., Ayala, J. M., Harper, C. F., Christen, A., Olszewski, J., Konteatis, Z., Webber, R., and Mumford, R. A. (1995). Quantification of a matrix metalloproteinasegenerated aggrecan G1 fragment using monospecific anti-peptide serum. Biochem. J. 307, 245–252. Lark, M. W., Gordy, J. T., Weidner, J. R., Ayala, J., Kimura, J. H., Williams, H. R., Mumford, R. A., Flannery, C. R., Carlson, S. S., Iwata, M., and Sandy, J. D. (1995). Cell-mediated catabolism of aggrecan. Evidence that cleavage of the “aggrecanase” site (Glu 373Ala 374) is a primary event in proteolysis of the interglobular domain. J. Biol. Chem. 270, 2550–2556. Fosang, A. J., Last, K., Gardiner, P., Jackson, D. C., and Brown, L. (1995). Development of a cleavage-site-specific monoclonal antibody for detecting metalloproteinase-derived aggrecan fragments: detection of fragments in human synovial fluid. Biochem. J. 310, 337–343.

Chapter 25 Products of Cartilage Metabolism 132. Singer, I. I., Kawka, D. W., Bayne, E. K., Donatelli, S. A., Weidner, J. R., Williams, H. A., Ayala, J. M., Mumford, R. A., Lark, M. W., Glant, T. T., Nabozny, G. H., and David, C. S. (1995). VDIPEN, A metalloproteinasegenerated neoepitope is induced and immunolocalized in articular cartilage during inflammatory arthritis. J. Clin. Invest. 95, 2178–2186. 133. Fosang, A. J., Last, K., and Maciewicz, R. A. (1996). Aggrecan is degraded by matrix metalloproteinases in human arthritis. Evidence that matrix metalloproteinases and aggrecanase activities can be independent. J. Clin. Invest. 98, 2292–2299. 134. Lark, M.W., Bayne, E.K., Flanagan, J., Harper, C. F., Hoerrner, L. A., Hutchinson, N. I., Singer, I. I., Donatelli, S. A., Weidner, J. R., Williams, H. R., Mumford, R. A., and Lohmander, L. S. (1997). Aggrecan degradation in human cartilage. Evidence for both metalloproteinase and aggrecanase activity in normal, osteoarthritic and rheumatoid joints. J. Clin. Invest. 100, 93–106. 135. Roberts, C. R., Roughley, P. J., and Mort, J. S. (1989). Degradation of human proteoglycan aggregate induced by hydrogen peroxide. Protein fragmentation, amino acid modification and hyaluronic acid cleavage. Biochem. J. 259, 805–811. 136. Ilic, M. Z., Handley, C. H., Robinson, H. C., and Mok, M. T. (1992). Mechanism of catabolism of aggrecan by articular cartilage. Arch. Biochem. Biophys. 294, 115–122. 137. Loulakis, P., Shrikhande, A., Davis, G., and Maniglia, C. A. (1992). N-terminal sequence of proteoglycan fragments isolated from medium of interleukin-1-treated articular-cartilage cultures. Putative site(s) of enzymic cleavage. Biochem J. 284 (Pt 2), 589–593. 138. Rosenberg, L., Wolfenstein-Todel, C., Margolis, R., Pal, S., and Strider, W. (1976). Proteoglycans from bovine proximal humeral articular cartilage. Structural basis for the polydispersity of proteoglycan subunits. J. Biol. Chem. 251, 6439–6444. 139. Heinegard, D. (1977). Polydispersity of cartilage proteoglycans. Structural variation with size and buoyant density of the molecules. J. Biol. Chem. 252, 1980–1989. 140. Bayliss, M. T., and Ali, S. Y. (1978). Age-related differences in the composition and structure of human articular cartilage proteoglycans. Biochem. J. 176, 683–693. 141. Morgelin, M., Paulsson, M., Hardingham, T. E., Heinegard, D., and Engel, J. (1988). Cartilage proteoglycans. Assembly with hyaluronate and link protein as studied by electron microscopy. Biochem. J. 253, 175–185. 142. Sandy, J. D., Plaas, A. H., and Koob, T. J. (1995). Pathways of aggrecan processing in joint tissues. Implications for disease mechanism and monitoring. Acta. Orthop. Scand. Suppl. 266, 26–32. 143. Dudhia, J., Davidson, C. M., Wells, T. M., Vynios, D. H., Hardingham, T. E., and Bayliss, M. T. (1996). Age-related changes in the content of the C-terminal region of aggrecan in human articular cartilage. Biochem J. 313 (Pt 3), 933–940. 144. Sandy, J. D., and Verscharen, C. (2001). Analysis of aggrecan in human knee cartilage and synovial fluid indicates that aggrecanase (ADAMTS) activity is responsible for the catabolic turnover and loss of whole aggrecan whereas other protease activity is required for C-terminal processing in vivo. Biochem. J. 358 (Pt 3), 615–626. 145. Patwari, P., Kurz, B., Sandy, J. D., and Grodzinsky, A. J. (2000). Mannosamine inhibits aggrecanase-mediated changes in the physical properties and biochemical composition of articular cartilage. Arch. Biochem. Biophys. 374, 79–85. 146. Flannery, C. R., Lark, M. W., and Sandy, J. D. (1992). Identification of a stromelysin cleavage site within the interglobular domain of human aggrecan. Evidence for proteolysis at this site in vivo in human articular cartilage. J. Biol. Chem. 267(2), 1008–1014. 147. Fosang, A. J., Neame, P. J., Last, K., Hardingham, T. E., Murphy, G., and Hamilton, J. A. (1992). The interglobular domain of cartilage aggrecan is cleaved by Pump, gelatinases and cathepsin B. J. Biol. Chem. 267, 19470–19474.

443 148. Fosang, A. J., Last, K., Knauper, V., Murphy, G., and Neame, P. J. (1996). Degradation of cartilage aggrecan by collagenase-3 (MMP-13). FEBS Lett. 380(1-2), 17–20. 149. Mort, J. S., Magny, M. C., and Lee, E. R. (1998). Cathepsin B: an alternative protease for the generation of an aggrecan ‘metalloproteinase’ cleavage neoepitope. Biochem J. 335 (Pt 3), 491–494. 150. Little, C. B., Flannery, C. R., Hughes, C. E., Mort, J. S., Roughley, P. J., Dent, C., and Caterson, B. (1999). Aggrecanase versus matrix metalloproteinases in the catabolism of the interglobular domain of aggrecan in vitro. Biochem. J. 344 (Pt 1), 61–68. 151. Fosang, A. J., Last, K., Stanton, H., Weeks, D. B., Campbell, I. K., Hardingham, T. E., and Hembry, R. M. (2000). Generation and novel distribution of matrix metalloproteinase-derived aggrecan fragments in porcine cartilage explants. Generation and novel distribution of matrix metalloproteinase-derived aggrecan fragments in porcine cartilage explants. J. Biol. Chem. 275(42), 33027–33037. 152. Hascall, V. C., Sandy, J. D., and Handley, C. J. (1999). Regulation of proteoglycan metabolism in articular cartilage. In “Biology of the Synovial Joint” (C. W. Archer, B. Caterson, M. Benjamin, J. R. Ralphs, eds), pp. 101–120, Harwood Academic Publishers, The Netherlands. 153. Buttner, F. H., Hughes, C. E., Margerie, D., Lichte, A., Tschesche, H., Caterson, B., and Bartnik, E. (1998). Membrane type 1 matrix metalloproteinase (MT1-MMP) cleaves the recombinant aggrecan substrate rAgg1mut at the ‘aggrecanase’ and the MMP sites. Characterization of MT1-MMP catabolic activities on the interglobular domain of aggrecan. Biochem. J. 333 (Pt 1), 159–165. 154. Tortorella, M. D., Burn, T. C., Pratta, M. A., Abbaszade, I., Hollis, J. M., Liu, R., Rosenfeld, S. A., Copeland, R. A., Decicco, C. P., Wynn, R., Rockwell, A., Yang, F., Duke, J. L., Solomon, K., George, H., Bruckner, R., Nagase, H., Itoh, Y., Ellis, D. M., Ross, H., Wiswall, B. H., Murphy, K., Hillman M. C. Jr, Hollis, G. F., Newton, R. C., Magolda, R. L., Trzaskos, J. M., and Arner E. C. (1999). Purification and cloning of aggrecanase-1: a member of the ADAMTS family of proteins. Science 284, 1664–1666. 155. Tortorella, M. D., Pratta, M., Liu, R. Q., Austin, J., Ross, O. H., Abbaszade, I., Burn, T., and Arner, E. (2000). Sites of aggrecan cleavage by recombinant human aggrecanase-1 (ADAMTS-4). J. Biol. Chem. 275, 18566–18573. 156. Sandy, J. D., Flannery, C. R., Neame, P. J., and Lohmander, L. S. (1992). The structure of aggrecan fragments in human synovial fluid. Evidence for the involvement in osteoarthritis of a novel proteinase which cleaves the Glu 373-Ala 374 bond of the interglobular domain. J. Clin. Invest. 89, 1512–1516. 157. Hu, L. T., Eskildsen, M. A., Masgala, C., Steere, A. C., Arner, E. C., Pratta, M. A., Grodzinsky, A. J., Loening, A., and Perides, G. (2001). Host metalloproteinases in Lyme arthritis. Arthritis Rheum. 44, 1401–1410. 158. Lark, M. W., Gordy, J. T., Weidner, J. R., Ayala, J., Kimura, J. H., Williams, H. R., Mumford, R. A., Flannery, C. R., Carlson, S. S., Iwata, M., and Sandy, J. D. (1995). Cell-mediated catabolism of aggrecan. Evidence that cleavage of the « aggrecanase » site (Glu 373-Ala 374) is a primary event in proteolysis of the interglobular domain. J. Biol. Chem. 270, 2550–2556. 159. Arner, E. C., Pratta, M. A., Trzaskos, J. M., Decicco, C. P., and Tortorella, M. D. (1999). Generation and characterization of aggrecanase. A soluble, cartilage-derived aggrecan-degrading activity. J. Biol. Chem. 274, 6594–6601. 160. Sugimoto, K., Takahashi, M., Yamamoto, Y., Shimada, K., and Tanzawa, K. (1999). Identification of aggrecanase activity in medium of cartilage culture. J. Biochem. (Tokyo). 126, 449–455. 161. Pratta, M. A., Tortorella, M. D., and Arner, E. C. (2000). Age-related changes in aggrecan glycosylation affect cleavage by aggrecanase. J. Biol. Chem. 275, 39096–39102. 162. Sandy, J. D., Thompson, V., Doege, K., and Verscharen, C. (2000). The intermediates of aggrecanase-dependent cleavage of aggrecan

444

163.

164.

165.

166.

167.

168.

169.

170.

171.

172.

173.

174.

175.

DANIEL-HENRI MANICOURT ET AL. in rat chondrosarcoma cells treated with interleukin-1. Biochem. J. 351 (Pt 1), 161–166. Bayliss, M. T., Hutton, S., Hayward, J., and Maciewicz, R. A. (2001). Distribution of aggrecanase (ADAMts 4/5) cleavage products in normal and osteoarthritic human articular cartilage: the influence of age, topography and zone of tissue. Osteoarthritis Cartilage 9, 553–560. Chambers, M. G., Cox, L., Chong, L., Suri, N., Cover, P., Bayliss, M. T., and Mason, R. M. (2001). Matrix metalloproteinases and aggrecanases cleave aggrecan in different zones of normal cartilage but colocalize in the development of osteoarthritic lesions in STR/ort mice. Arthritis Rheum. 44, 1455–1465. Lee, E. R., Lamplugh, L., Davoli, M. A., Beauchemin, A., Chan, K., Mort, J. S., and Leblond, C. P. (2001). Enzymes active in the areas undergoing cartilage resorption during the development of the secondary ossification center in the tibiae of rats ages 0-21 days: I. Two groups of proteinases cleave the core protein of aggrecan. Dev Dyn. 222, 52–70. Sztrolovics, R., Alini, M., Roughley, P. J., and Mort, J. S. (1997). Aggrecan degradation in human intervertebral disc and articular cartilage. Biochem. J. 326 ( Pt 1), 235–241. Mercuri, F. A., Maciewicz, R. A., Tart, J., Last, K., and Fosang, A. J. (2000). Mutations in the interglobular domain of aggrecan alter matrix metalloproteinase and aggrecanase cleavage patterns. Evidence that matrix metalloproteinase cleavage interferes with aggrecanase activity. J. Biol. Chem. 275, 33038–33045. Westling, J., Fosang, A. J., Last, K., Thompson, V. P., Tomkinson, K. N., Hebert, T., McDonagh, T., Collins-Racie, L. A., LaVallie, E. R., Morris, E. A., and Sandy, J. D. (2002). ADAMTS4 cleaves at the aggrecanase site (Glu373-Ala374) and secondarily at the matrix metalloproteinase site (Asn341-Phe342) in the aggrecan interglobular domain. J. Biol. Chem. 277, 16059–16066. Epub 2002 Feb 19. Little, C. B., Hughes, C. E., Curtis, C. L., Janusz, M. J., Bohne, R., Wang-Weigand, S., Taiwo, Y. O., Mitchell, P. G., Otterness, I. G., Flannery, C. R., and Caterson, B. (2002). Matrix metalloproteinases are involved in C-terminal and interglobular domain processing of cartilage aggrecan in late stage cartilage degradation. Matrix Biol. 21, 271–288. van Meurs, J. B., van Lent, P. L., Holthuysen, A. E., Singer, I. I., Bayne, E. K., and van den Berg, W. B. (1999). Kinetics of aggrecanaseand metalloproteinase-induced neoepitopes in various stages of cartilage destruction in murine arthritis. Arthritis Rheum. 42, 1128–1139. van Meurs, J., van Lent, P., Stoop, R., Holthuysen, A., Singer, I., Bayne, E., Mudgett, J., Poole, R., Billinghurst, C., van der Kraan, P., Buma, P., and van den Berg, W. (1999). Cleavage of aggrecan at the Asn341-Phe342 site coincides with the initiation of collagen damage in murine antigen-induced arthritis: a pivotal role for stromelysin 1 in matrix metalloproteinase activity. Arthritis Rheum. 42, 2074–2084. van Meurs, J. B., van Lent, P. L., van de Loo, A. A., Holthuysen, A. E., Bayne, E. K., Singer, I. I., and van den Berg, W. B. (1999). Increased vulnerability of postarthritic cartilage to a second arthritic insult: accelerated MMP activity in a flare up of arthritis. Ann. Rheum. Dis. 58, 350–356. Pratta, M. A., Yao, W., Decicco, C., Tortorella, M. D., Liu, R. Q., Copeland, R. A., Magolda, R., Newton, R. C., Trzaskos, J. M., and Arner, E. C. (2003). Aggrecan protects cartilage collagen from proteolytic cleavage. J. Biol. Chem. 278, 45539–45545. Epub 2003 Jul 30. Flannery, C. R., Little, C. B., Hughes, C. E., Caterson, B. (1999). Expression of ADAMTS homologues in articular cartilage. Biochem. Biophys. Res. Commun. 260, 318–322. Koshy, P. J., Lundy, C. J., Rowan, A. D., Porter, S., Edwards, D. R., Hogan, A., Clark, I. M., and Cawston, T. E. (2002). The modulation of matrix metalloproteinase and ADAM gene expression in human chondrocytes by interleukin-1 and oncostatin M: a time-course study using real-time quantitative reverse transcription-polymerase chain reaction. Arthritis Rheum. 46, 961–967.

176. Sylvester, J., Liacini, A., Li, W. Q., and Zafarullah, M. (2004). Interleukin-17 signal transduction pathways implicated in inducing matrix metalloproteinase-3, -13 and aggrecanase-1 genes in articular chondrocytes. Cell Signal. 16, 469–476. 177. Kevorkian, L., Young, D. A., Darrah, C., Donell, S. T., Shepstone, L., Porter, S., Brockbank, S. M., Edwards, D. R., Parker, A. E., and Clark, I. M. (2004). Expression profiling of metalloproteinases and their inhibitors in cartilage. Arthritis Rheum. 50, 131–141. 178. Curtis, C. L., Hughes, C. E., Flannery, C. R., Little, C. B., Harwood, J. L., and Caterson, B. (2000). n-3 fatty acids specifically modulate catabolic factors involved in articular cartilage degradation. J. Biol. Chem. 275, 721–724. 179. Gao, G., Plaas, A., Thompson, V. P., Jin, S., Zuo, F., and Sandy, J. D. (2004). ADAMTS4 (aggrecanase-1) activation on the cell surface involves C-terminal cleavage by glycosylphosphatidyl inositolanchored membrane type 4-matrix metalloproteinase and binding of the activated proteinase to chondroitin sulfate and heparan sulfate on syndecan-1. J. Biol. Chem. 279, 10042–10051. Epub 2003 Dec 29. 180. Gao, G., Westling, J., Thompson, V. P., Howell, T. D., Gottschall, P. E., and Sandy, J. D. (2002). Activation of the proteolytic activity of ADAMTS4 (aggrecanase-1) by C-terminal truncation. J. Biol. Chem. 277, 11034–11041. Epub 2002 Jan 16. 181. Hashimoto, G., Shimoda, M., and Okada, Y. (2004). ADAMTS4 (aggrecanase-1) interaction with the C-terminal domain of fibronectin inhibits proteolysis of aggrecan. J. Biol. Chem. 279, 32483–32491. Epub 2004 May 25. 182. Flannery, C. R., Zeng, W., Corcoran, C., Collins-Racie, L. A., Chockalingam, P. S., Hebert, T., Mackie, S. A., McDonagh, T., Crawford, T. K., Tomkinson, K. N., LaVallie, E. R., and Morris, E. A. (2002). Autocatalytic cleavage of ADAMTS-4 (Aggrecanase-1) reveals multiple glycosaminoglycan-binding sites. J. Biol. Chem. 277, 42775–42780. Epub 2002 Aug 28. 183. Baker, A. H., Edwards, D. R., and Murphy G. (2002). Metalloproteinase inhibitors: biological actions and therapeutic opportunities. J. Cell. Sci. 115(Pt 19), 3719–3727. 184. Kashiwagi, M., Tortorella, M., Nagase, H., and Brew, K. (2001). TIMP-3 is a potent inhibitor of aggrecanase 1 (ADAM-TS4) and aggrecanase 2 (ADAM-TS5). J. Biol. Chem. 276, 12501–12504. Epub 2001 Jan 23. 185. Rodriguez-Manzaneque, J. C., Westling, J., Thai, S. N., Luque, A., Knauper, V., Murphy, G., Sandy, J. D., and Iruela-Arispe M. L. (2002). ADAMTS1 cleaves aggrecan at multiple sites and is differentially inhibited by metalloproteinase inhibitors. Biochem. Biophys. Res. Commun. 293, 501–508. 186. Tortorella, M. D., Arner, E. C., Hills, R., Easton, A., Korte-Sarfaty, J., Fok, K., Wittwer, A. J., Liu, R. Q., and Malfait, A. M. (2004). Alpha2macroglobulin is a novel substrate for ADAMTS-4 and ADAMTS-5 and represents an endogenous inhibitor of these enzymes. J. Biol. Chem. 279, 17554–17561. Epub 2004 Jan 07. 187. Plaas, A. H., Wong-Palms, S., Roughley, P. J., Midura, R. J., and Hascall, V. C. (1997). Chemical and immunological assay of the nonreducing terminal residues of chondroitin sulfate from human aggrecan. J. Biol. Chem. 272, 20603–20610. 188. Bayliss, M. T., Osborne, D., Woodhouse, S., and Davidson, C. (1999). Sulfation of chondroitin sulfate in human articular cartilage. The effect of age, topographical position, and zone of cartilage on tissue composition. J. Biol. Chem. 274, 15892–15900. 189. Plaas, A. H., West, L. A., Wong-Palms, S., and Nelson, F. R. (1998). Glycosaminoglycan sulfation in human osteoarthritis. Disease-related alterations at the non-reducing termini of chondroitin and dermatan sulfate J. Biol. Chem. 273, 12642–12649. 190. van den Berg, W. B., van der Kraan, P. M., Scharstuhl, A., and van Beuningen, H. M. (2001). Growth factors and cartilage repair. Clin. Orthop. (391 Suppl): S244–S250.

Chapter 25 Products of Cartilage Metabolism 191. Martel-Pelletier, J., Alaaeddine, N., and Pelletier, J. P. (1999). Cytokines and their role in the pathophysiology of osteoarthritis. Front Biosci. 4, D694–D703. 192. Sauerland, K., Plaas, A. H., Raiss, R. X., and Steinmeyer, J. (2003). The sulfation pattern of chondroitin sulfate from articular cartilage explants in response to mechanical loading. Biochim. Biophys. Acta. 1638, 241–248. 193. Caterson, B., Hughes, C. E., Johnstone, B., Mort, J. S. (1992). Immunological markers of cartilage proteoglycan metabolism in animal and human osteoarthritis. In “Articular Cartilage and Osteoarthritis” (K. E. Kuettner, R. Schleyerbach, J. G. Peyron, and V. C. Hascall, eds), pp. 415–427, Raven Press, New York. 194. Visco, D. M., Johnstone, B., Hill, M. A., Jolly, G. A., and Caterson, B. (1993). Immunohistochemical analysis of 3-B-(-) and 7-D-4 epitope expression in canine osteoarthritis. Arthritis Rheum. 36, 1718–1725. 195. Slater, R. R., Jr, Bayliss, M. T., Lachiewicz, P. F., Visco, D. M., and Caterson, B. (1995). Monoclonal antibodies that detect biochemical markers of arthritis in humans. Arthritis Rheum. 38, 655–659. 196. Hazell, P. K., Dent, C., Fairclough, J. A., Bayliss, M. T., and Hardingham, T. E. (1995). Changes in glycosaminoglycan epitope levels in knee joint fluid following injury. Arthritis Rheum. 38, 953–959. 197. Bautch, J. C., Clayton, M. K., Chu, Q., and Johnson, K. A. (2000). Synovial fluid chondroitin sulphate epitopes 3B3 and 7D4, and glycosaminoglycan in human knee osteoarthritis after exercise. Ann. Rheum. Dis. 59, 887–891. 198. Poole, A. R. (2003). Biochemical/immunochemical biomarkers of osteoarthritis: utility for prediction of incident or progressive osteoarthritis. Rheum. Dis. Clin. North. Am. 29, 803–818. 199. Poole, A. R., Ionescu, M., Swan, A., and Dieppe, P. A. (1994). Changes in cartilage metabolism in arthritis are reflected by altered serum and synovial fluid levels of the cartilage proteoglycan aggrecan. J. Clin. Invest. 94, 25–33. 200. Carlson, C. S., Loeser, R. F., Johnstone, B., Tulli, H. M., Dobson, D. B., and Caterson, B. (1995). Osteoarthritis in cynomolgus macaques. II. Detection of modulated proteoglycan epitopes in cartilage and synovial fluid. J. Orthop. Res. 13, 399–409. 201. Belcher, C., Yaqub, R., Fawthrop, F., Bayliss, M., and Doherty, M. (1997). Synovial fluid chondroitin and keratan sulphate epitopes, glycosaminoglycans, and hyaluronan in arthritic and normal knees. Ann. Rheum. Dis. 56, 299–307. 202. Fawthrop, F., Yaqub, R., Belcher, C., Bayliss, M., Ledingham, J., and Doherty, M. (1997). Chondroitin and keratan sulphate epitopes, glycosaminoglycans, and hyaluronan in progressive versus nonprogressive osteoarthritis. Ann. Rheum. Dis. 56, 119–122. 203. Thonar, E. J-M. A., Manicourt, D. H., Williams, J. M., Fukuda, K., Campion, G. V., Sweet, B. M. E., Lenz, M. E., Schnitzer, T. J., and Kuettner, K. E. (1992). Serum keratan sulfate: a measure of cartilage proteoglycan metabolism. In “Articular Cartilage and Osteoarthritis” (K. E. Kuettner, R. Schleyerbach, J. G. Peyron, and V. C. Hascall, eds), pp. 429–445, Raven Press, New York. 204. Webber, C., Glant, T. T., Roughley, P. J., and Poole, A. R. (1987). The identification and characterization of two populations of aggregating proteoglycans of high buoyant density isolated from post-natal human articular cartilages of different ages. Biochem. J. 248, 735–740. 205. Scott, J. E. (1992). Oxygen and the connective tissues. T I B S 17, 340–343. 206. Manicourt, D-H., Pita, J. C., Thonar, E. J-M., and Howell, D. S. (1991). Proteoglycans non-dissociatively extracted from different zones of canine normal articular cartilage: variations in the sedimentation profile of aggrecan with degree of physiological stress. Connect. Tissue Res. 26, 231–246. 207. Sweet, M. B., Thonar, E. J., Immelman, A. R., and Solomon, L. (1977). Biochemical changes in progressive osteoarthrosis. Ann. Rheum. Dis. 36, 387–398.

445 208. Brown, G. M., Huckerby, T. N., Bayliss, M. T., and Nieduszynski, I. A. (1998). Human aggrecan keratan sulfate undergoes structural changes during adolescent development. J. Biol. Chem. 273, 26408–26414. 209. Iozzo R. V. (1999). The biology of the small leucine-rich proteoglycans. Functional network of interactive proteins J. Biol. Chem. 274, 18843–18846. 210. Burton-Wurster, N., Liu, W., Matthews, G. L., Lust, G., Roughley, P. J., and Glant, T. T. (2003). Cs-Szabo G. TGF beta 1 and biglycan, decorin, and fibromodulin metabolism in canine cartilage. Osteoarthritis Cartilage 11, 167–176. 211. Knudson, C. B., and Knudson, W. (2001). Cartilage proteoglycans. Semin. Cell Dev. Biol. 12, 69–78. 212. Danielson, K. G., Baribault, H., Holmes, D. F., Graham, H., Kadler, K. E., and Iozzo, R. V. (1997). Targeted disruption of decorin leads to abnormal collagen fibril morphology and skin fragility. J. Cell. Biol. 136, 729–743. 213. Xu, T., Bianco, P., Fisher, L. W., Longenecker, G., Smith, E., Goldstein, S., Bonadio, J., Boskey, A., Heegaard, A. M., Sommer, B., Satomura, K., Dominguez, P., Zhao, C., Kulkarni, A. B., Robey, P. G., and Young, M. F. (1998). Targeted disruption of the biglycan gene leads to an osteoporosis-like phenotype in mice. Nat Genet. 20, 78–82. 214. Kashiwagi, M., Enghild, J. J., Gendron, C., Hughes, C., Caterson, B., Itoh, Y., and Nagase, H. (2004). Altered proteolytic activities of ADAMTS-4 expressed by C-terminal processing. J. Biol. Chem. 279, 10109–10119. Epub 2003 Dec 08. 215. Poole, A. R., Rosenberg, L. C., Reiner, A., Ionescu, M., Bogoch, E., and Roughley, P. J. (1996). Contents and distributions of the proteoglycans decorin and biglycan in normal and osteoarthritic human articular cartilage. J. Orthop. Res. 14, 681–689. 216. Cs-Szabo, G., Roughley, P. J., Plaas, A. H., and Glant, T. T. (1995). Large and small proteoglycans of osteoarthritic and rheumatoid articular cartilage. Arthritis Rheum. 38, 660–668. 217. Cs-Szabo, G., Melching, L. I., Roughley, P. J., and Glant, T. T. (1997). Changes in messenger RNA and protein levels of proteoglycans and link protein in human osteoarthritic cartilage samples. Arthritis Rheum. 40, 1037–1045. 218. Bock, H. C., Michaeli, P., Bode, C., Schultz, W., Kresse, H., Herken, R., and Miosge, N. (2001). The small proteoglycans decorin and biglycan in human articular cartilage of late-stage osteoarthritis. Osteoarthritis Cartilage 9, 654–663. 219. Polgar, A., Falus, A., Koo, E., Ujfalussy, I., Sesztak, M., Szuts, I., Konrad, K., Hodinka, L., Bene, E., Meszaros, G., Ortutay, Z., Farkas, E., Paksy, A., and Buzas, E. I. (2003). Elevated levels of synovial fluid antibodies reactive with the small proteoglycans biglycan and decorin in patients with rheumatoid arthritis or other joint diseases. Rheumatology (Oxford) 42, 522–527. 220. Melching, L. I., and Roughley, P. J. (1999). Modulation of keratan sulfate synthesis on lumican by the action of cytokines on human articular chondrocytes. Matrix Biol. 18, 381–390. 221. Bengtsson, E., Neame, P. J., Heinegard, D., and Sommarin, Y. (1995). The primary structure of a basic leucine-rich repeat protein, PRELP, found in connective tissues. J. Biol. Chem. 270, 25639–25644. 222. Bengtsson, E., Morgelin, M., Sasaki, T., Timpl, R., Heinegard, D., and Aspberg, A. (2002). The leucine-rich repeat protein PRELP binds perlecan and collagens and may function as a basement membrane anchor. J. Biol. Chem. 277, 15061–15068. Epub 2002 Feb 14. 223. SundarRaj, N., Fite, D., Ledbetter, S., Chakravarti, S., and Hassell, J. R. (1995). Perlecan is a component of cartilage matrix and promotes chondrocyte attachment. J. Cell Sci. 108 (Pt 7), 2663–2672. 224. Nicole, S., Davoine, C. S., Topaloglu, H., Cattolico, L., Barral, D., Beighton, P., Hamida, C. B., Hammouda, H., Cruaud, C., White, P. S., Samson, D., Urtizberea, J. A., Lehmann-Horn, F., Weissenbach, J., Hentati, F., and Fontaine, B. (2000). Perlecan, the major proteoglycan

446

225.

226.

227.

228.

229.

230.

231.

232.

233.

234.

235.

236.

237.

238.

239.

240.

DANIEL-HENRI MANICOURT ET AL. of basement membranes, is altered in patients with Schwartz-Jampel syndrome (chondrodystrophic myotonia). Nat Genet. 26, 480–483. Arikawa-Hirasawa, E., Watanabe, H., Takami, H., Hassell, J. R., and Yamada, Y. (1999). Perlecan is essential for cartilage and cephalic development. Nat. Genet. 23, 354–358. Costell, M., Gustafsson, E., Aszodi, A., Morgelin, M., Bloch, W., Hunziker, E., Addicks, K., Timpl, R., and Fassler, R. (1999). Perlecan maintains the integrity of cartilage and some basement membranes. J. Cell Biol. 147, 1109–1122. Handler, M., Yurchenco, P. D., and Iozzo, R. V. (1997). Developmental expression of perlecan during murine embryogenesis. Dev. Dyn. 210, 130–145. French, M. M., Smith, S. E., Akanbi, K., Sanford, T., Hecht, J., FarachCarson, M. C., and Carson, D. D. (1999). Expression of the heparan sulfate proteoglycan, perlecan, during mouse embryogenesis and perlecan chondrogenic activity in vitro. J Cell Biol. 145, 1103–1115. Kou, I., and Ikegawa, S. (2004). SOX9-dependent and -independent transcriptional regulation of human cartilage link protein. J. Biol. Chem. 279, 50942–50948. Epub 2004 Sep 28. Seguin, C. A., Bernier, S. M. (2003). TNFalpha suppresses link protein and type II collagen expression in chondrocytes: Role of MEK1/2 and NF-kappaB signaling pathways. J. Cell. Physiol. 197, 356–369. Fitzgerald, J. B., Jin, M., Dean, D., Wood, D. J., Zheng, M. H., and Grodzinsky, A. J. (2004). Mechanical compression of cartilage explants induces multiple time-dependent gene expression patterns and involves intracellular calcium and cyclic AMP. J. Biol. Chem. 279, 19502–19511. Epub 2004 Feb 11. Bolton, M. C., Dudhia, J., and Bayliss, M. T. (1999). Age-related changes in the synthesis of link protein and aggrecan in human articular cartilage: implications for aggregate stability. Biochem. J. 337 (Pt 1), 77–82. Roughley, P. J., Nguyen, Q., and Mort, J. S. (1992). The role of proteinases and oxygen radicals in the degradation of human articular cartilage. In “Articular Cartilage and Osteoarthritis” (K. E. Kuettner, R. Schleyerbach, J. G. Peyron and, V. C. Hascall, eds), pp. 305–317, Raven Press, New York. Mwale, F., Demers, C. N., Petit, A., Roughley, P., Poole, A. R., Steffen, T., Aebi, M., and Antoniou, J. (2003). A synthetic peptide of link protein stimulates the biosynthesis of collagens, II, IX and proteoglycan by cells of the intervertebral disc. J. Cell. Biochem. 88, 1202–1213. Dean, M. F., and Sansom, P. (2000). Link peptide cartilage growth factor is degraded by membrane proteinases. Biochem J. 349 (Pt 2), 473–479. McKenna, L. A., Liu, H., Sansom, P. A., and Dean, M. F. (1998). An N-terminal peptide from link protein stimulates proteoglycan biosynthesis in human articular cartilage in vitro. Arthritis Rheum. 41, 157–162. Liu, H., McKenna, L. A., and Dean, M. F. (2000). An N-terminal peptide from link protein can stimulate biosynthesis of collagen by human articular cartilage. Arch. Biochem. Biophys. 378, 116–122. Ilic, M. Z., Handley., C. H., Robinson., H. C., and Mok, M. T. (1992). Mechanism of catabolism of aggrecan by articular cartilage. Arch. Biochem. Biophys. 294, 115–122. Goodstone, N. J., Doran, M. C., Hobbs, R. N., Butler, R. C., Dixey, J. J., and Ashton, B. A. (1996). Cellular immunity to cartilage aggrecan core protein in patients with rheumatoid arthritis and non-arthritic controls. Ann. Rheum. Dis. 55, 40–46. Guerassimov, A., Duffy, C., Zhang, Y., Banerjee, S., Leroux, J. Y., Reimann, A., Webber, C., Delaunay, N., Vipparti, V., Ronbeck, L., Cartman, A., Arsenault, L., Rosenberg, L. C., and Poole, A. R. (1997). Immunity to cartilage link protein in patients with juvenile rheumatoid arthritis. J. Rheumatol. 24, 959–964.

241. Guerassimov, A., Zhang, Y., Banerjee, S., Cartman, A., Webber, C., Esdaile, J., Fitzcharles, M. A., and Poole, A. R. (1998). Autoimmunity to cartilage link protein in patients with rheumatoid arthritis and ankylosing spondylitis. J. Rheumatol. 25, 1480–1484. 242. Guerassimov, A., Zhang, Y., Cartman, A., Rosenberg, L. C., Esdaile, J., Fitzcharles, M. A., and Poole, A. R. (1999). Immune responses to cartilage link protein and the G1 domain of proteoglycan aggrecan in patients with osteoarthritis. Arthritis Rheum. 42, 527–533. 243. Ratcliffe, A., Beauvais, P. J., Saed-Nejad, F. (1994). Differential levels of synovial fluid aggrecan aggregate components in experimental osteoarthritis and joint disuse. J. Othop. Res. 12, 464–473. 244. Ratcliffe, A., Billingham, M. E., Saed-Nejad, F., Muir, H., and Hardingham, T. E. (1992). Increased release of matrix components from articular cartilage in experimental canine osteoarthritis. J. Orthop. Res. 10, 350–358. 245. Itano, N., and Kimata, K. (2002). Mammalian hyaluronan synthases. IUBMB Life 54, 195–199. 246. Nishida, Y., Knudson, C. B., Nietfeld, J. J., Margulis, A., and Knudson, W. (1999). Antisense inhibition of hyaluronan synthase-2 in human articular chondrocytes inhibits proteoglycan retention and matrix assembly. J. Biol. Chem. 274, 21893–21899. 247. Hiscock, D. R., Caterson, B., and Flannery, C. R. (2000). Expression of hyaluronan synthases in articular cartilage. Osteoarthritis Cartilage. 8, 120–126. 248. Recklies, A. D., White, C., Melching, L., and Roughley, P. J. (2001). Differential regulation and expression of hyaluronan synthases in human articular chondrocytes, synovial cells and osteosarcoma cells. Biochem. J. 354 (Pt 1), 17–24. 249. Bodevin-Authelet, S., Kusche-Gullberg, M., Pummill, P. E., DeAngelis, P. L., and Lindahl, U. (2005). Biosynthesis of hyaluronan: direction of chain elongation. J. Biol. Chem. 280, 8813–8818. Epub 2004 Dec 28. 250. Itano, N., Sawai, T., Yoshida, M., Lenas, P., Yamada, Y., Imagawa, M., Shinomura, T., Hamaguchi, M., Yoshida, Y., Ohnuki, Y., Miyauchi, S., Spicer, A. P., McDonald, J. A., and Kimata K. (1999). Three isoforms of mammalian hyaluronan synthases have distinct enzymatic properties. J. Biol. Chem. 274, 25085–25092. 251. Ng, C. K., Handley, C. J., Mason, R. M., and Robinson, H. C. (1989). Synthesis of hyaluronate in cultured bovine articular cartilage. Biochem. J. 263, 761–767. 252. Holmes, M. W. A., Bayliss, M. T., and Muir, H. (1988). Hyaluronic acid in human articular cartilage. Age-related changes in content and size. Biochem. J. 250, 435–441. 253. El Hajjaji, H., Cole, A. A., and Manicourt, D. H. (2005). Chondrocytes, synoviocytes and dermal fibroblasts all express PH-20, a hyaluronidase active at neutral pH. Arthritis Res. Ther. 7, R756–R768. 254. Kvam, B. J., Fragonas, E., Degrassi, A., Kvam, C., Matulova, M., Pollesello, P., Zanetti, F., and Vittur, F. (1995). Oxygen-derived free radical (ODFR) action on hyaluronan (HA), on two HA ester derivatives, and on the metabolism of articular chondrocytes. Exp. Cell Res. 218, 79–86. 255. Parsons, B., Al-Assaf, S., Navaratnam, S., and Phillips, G. (2002). Comparison of the reactivity of different reactive oxidative species towards hyaluronan. In “Hyaluronan.” Volume 1 (J. Kennedy, G. Phillips, P. Williams, V. Hascall, eds), pp. 141–150, MFK Group Ltd. Cambridge, England. 256. Noble, P. W. (2002). Hyaluronan and its catabolic products in tissue injury and repair. Matrix Biol. 21, 25–29. 257. Knudson, C. B., and Knudson, W. (2004). Hyaluronan and CD44: modulators of chondrocyte metabolism. Clin. Orthop. (427 Suppl), S152–S162. 258. Knudson, W., Casey, B., Nishida, Y., Eger, W., Kuettner, K. E., and Knudson, C. B. (2000). Hyaluronan oligosaccharides perturb cartilage matrix homeostasis and induce chondrocytic chondrolysis. Arthritis Rheum. 43, 1165–1174.

Chapter 25 Products of Cartilage Metabolism 259. Nishida, Y., Knudson, C. B., and Knudson, W. (2004). Osteogenic Protein-1 inhibits matrix depletion in a hyaluronan hexasaccharideinduced model of osteoarthritis. Osteoarthritis Cartilage 12, 374–382. 260. Fukuda, K., Dan, H., Takayama, M., Kumano, F., Saitoh, M., and Tanaka, S. (1996). Hyaluronic acid increases proteoglycan synthesis in bovine articular cartilage in the presence of interleukin-1. J. Pharmacol. Exp. Ther. 277, 1672–1675. 261. Fukuda, K., Takayama, M., Ueno, M., Oh, M., Asada, S., Kumano, F., and Tanaka, S. (1997). Hyaluronic acid inhibits interleukin-1-induced superoxide anion in bovine chondrocytes. Inflamm. Res. 46, 114–117. 262. Homandberg, G. A., Hui, F., Wen, C., Kuettner, K. E., and Williams, J. M. (1997). Hyaluronic acid suppresses fibronectin fragment mediated cartilage chondrolysis: I. in vitro. Osteoarthritis Cartilage 5, 309–319. 263. Williams, J. M., Zhang, J., Kang, H., Ummadi, V., and Homandberg, G. A. (2003). The effects of hyaluronic acid on fibronectin fragment mediated cartilage chondrolysis in skeletally mature rabbits. Osteoarthritis Cartilage 11, 44–49. 264. Uebelhart, D., and Williams, J. M. (1999). Effects of hyaluronic acid on cartilage degradation. Curr. Opin. Rheumatol. 11, 427–435. 265. Altman, R. D., Akermark, C., Beaulieu, A. D., and Schnitzer, T., Durolane International Study Group. (2004). Efficacy and safety of a single intra-articular injection of non-animal stabilized hyaluronic acid (NASHA) in patients with osteoarthritis of the knee. Osteoarthritis Cartilage 12, 642–649. 266. Bayliss, M. T., Howat, S. L., Dudhia, J., Murphy, J. M., Barry, F. P., Edwards, J. C., and Day, A. J. (2001). Up-regulation and differential expression of the hyaluronan-binding protein TSG-6 in cartilage and synovium in rheumatoid arthritis and osteoarthritis. Osteoarthritis Cartilage 9, 42–48. 267. Bork, P., and Beckmann, G. (1999). The CUB domain. A widespread module in developmentally regulated proteins. J. Mol. Biol. 231, 539–545. 268. Wisniewski, H. G., and Vilcek, J. (1997). TSG-6: an IL-1/TNFinducible protein with anti-inflammatory activity. Cytokine Growth Factor Rev. 8, 143–156. 269. Solis, D., Romero, A., Jimenez, M., Diaz-Maurino, T., and Calvete, J. J. (1998). Binding of mannose-6-phosphate and heparin by boar seminal plasma PSP-II., a member of the spermadhesin protein family. FEBS Lett. 431, 273–278. 270. Glant, T. T., Kamath, R. V., Bardos, T., Gal, I., Szanto, S., Murad, Y. M., Sandy, J. D., Mort, J. S., Roughley, P. J., and Mikecz, K. (2002). Cartilage-specific constitutive expression of TSG-6 protein (product of tumor necrosis factor alpha-stimulated gene 6) provides a chondroprotective, but not antiinflammatory, effect in antigen-induced arthritis. Arthritis Rheum. 46, 2207–2218. 271. Henrotin, Y. E., Bruckner, P., and Pujol, J. P. (2003). The role of reactive oxygen species in homeostasis and degradation of cartilage. Osteoarthritis Cartilage 11, 747–755. 272. Amin, A. R., and Abramson, S. B. (1998). The role of nitric oxide in articular cartilage breakdown in osteoarthritis. Curr. Opin. Rheumatol. 10, 263–268. 273. Lyons, C. R. (1995). The role of nitric oxide in inflammation. Adv. Immunol. 60, 323-371. 274. Aeberhard, E. E., Henderson, S. A., Arabolos, N. S., Griscavage, J. M., Castro, F. E., Barrett, C. T., and Ignarro, L. J. (1995). Nonsteroidal anti-inflammatory drugs inhibit expression of the inducible nitric oxide synthase gene. Biochem. Biophys. Res. Commun. 208, 1053–1059. 275. Clancy, R. M., Gomez, P. F., and Abramson, S. B. (2004). Nitric oxide sustains nuclear factor kappaB activation in cytokine-stimulated chondrocytes. Osteoarthritis Cartilage 12, 552–558. 276. Stefanovic-Racic, M., Stadler, J., and Evans, C. H. (1993). Nitric oxide and arthritis. Arthritis Rheum. 36, 1036–1044.

447 277. Baddour, V. T., and Bradley, J. D. (1999). Clinical assessment and significance of inflammation in knee osteoarthritis. Curr. Rheumatol. Rep. 1, 59–63. 278. Spreng, D., Sigrist, N., Schweighauser, A., Busato, A., and Schawalder, P. (2001). Endogenous nitric oxide production in canine osteoarthritis: Detection in urine, serum, and synovial fluid specimens. Vet. Surg. 30, 191–199. 279. Karan, A., Karan, M. A., Vural, P., Erten, N., Tascioglu, C., Aksoy, C., Canbaz, M., and Oncel, A. (2003). Synovial fluid nitric oxide levels in patients with knee osteoarthritis. Clin. Rheumatol. 22, 397–399. Epub 2003 Oct 18. 280. Kobayashi, K., Matsuzaka, S., Yoshida, Y., Miyauchi, S., Wada, Y., and Moriya, H. (2004). The effects of intraarticularly injected sodium hyaluronate on levels of intact aggrecan and nitric oxide in the joint fluid of patients with knee osteoarthritis. Osteoarthritis Cartilage 12, 536–542. 281. Moulton, P. J., Goldring, M. B., and Hancock, J. T. (1998). NADPH oxidase of chondrocytes contains an isoform of the gp91phox subunit. Biochem. J. 329 (Pt 3), 449–451. 282. Attur, M. G., Dave, M., Akamatsu, M., Katoh, M., and Amin, A. R. (2002). Osteoarthritis or osteoarthrosis: the definition of inflammation becomes a semantic issue in the genomic era of molecular medicine. Osteoarthritis Cartilage 10, 1–4. 283. Sasaki, K., Hattori, T., Fujisawa, T., Takahashi, K., Inoue, H., and Takigawa, M. (1995). Nitric oxide mediates interleukin-1-induced gene expression of matrix metalloproteinases and basic fibroblast growth factor in cultured rabbit articular chondrocytes. J. Biochem. (Tokyo) 123, 431–439. 284. Murrell, G. A., Jang, D., and Williams, R. J. (1995). Nitric oxide activates metalloprotease enzymes in articular cartilage. Biochem. Biophys. Res. Commun. 206, 15–21. 285. Hickery, M. S., and Bayliss, M. T. (1998). Interleukin-1 induced nitric oxide inhibits sulphation of glycosaminoglycan chains in human articular chondrocytes. Biochim. Biophys. Acta. 1425, 282–290. 286. Cao, M., Westerhausen-Larson, A., Niyibizi, C., Kavalkovich, K., Georgescu, H. I., Rizzo, C. F., Hebda, P. A., Stefanovic-Racic, M., and Evans, C. H. (1997). Nitric oxide inhibits the synthesis of type-II collagen without altering Col2A1 mRNA abundance: prolyl hydroxylase as a possible target. Biochem J. 324 (Pt 1), 305–310. 287. Pelletier, J. P., Jovanovic, D. V., Lascau-Coman, V., Fernandes, J. C., Manning, P. T., Connor, J. R., and Currie, M. G. (1997). MartelPelletier J. Selective inhibition of inducible nitric oxide synthase reduces progression of experimental osteoarthritis in vivo: possible link with the reduction in chondrocyte apoptosis and caspase 3 level. Arthritis Rheum. 43, 1290–1299. 288. Kurz, B., Jost, B., and Schunke, M. (2002). Dietary vitamins and selenium diminish the development of mechanically induced osteoarthritis and increase the expression of antioxidative enzymes in the knee joint of STR/1N mice. Osteoarthritis Cartilage 10, 119–126. 289. Studer, R. K., Levicoff, E., Georgescu, H., Miller, L., Jaffurs, D., and Evans, C. H. (2000). Nitric oxide inhibits chondrocyte response to IGF-I: inhibition of IGF-IRbeta tyrosine phosphorylation. Am. J. Physiol. Cell. Physiol. 279, C961–C969. 290. Del Carlo, M. Jr, and Loeser, R. F. (2002). Nitric oxide-mediated chondrocyte cell death requires the generation of additional reactive oxygen species. Arthritis Rheum. 46, 394–403. 291. Reiter, C. D., Teng, R. J., and Beckman, J. S. (2000). Superoxide reacts with nitric oxide to nitrate tyrosine at physiological pH via peroxynitrite. J. Biol. Chem. 275, 32460–32466. 292. Loeser, R. F., Carlson, C. S., Del Carlo, M., and Cole, A. (2002). Detection of nitrotyrosine in aging and osteoarthritic cartilage: Correlation of oxidative damage with the presence of interleukin1beta and with chondrocyte resistance to insulin-like growth factor 1. Arthritis Rheum. 6, 2349–2357.

448 293. Kwam, B. J., Fragonas, E., Degrassi, A., Kwam, C., Matulova, M., Pollesello, P., Zanetti, F., and Vittur, F. (1995). Oxygen-derived free radical (ODFR) action on hyaluronan (HA), on two HA ester derivatives, and on the metabolism of articular chondrocytes. Exp. Cell. Res. 218, 79–86. 294. Johnson, K., Vaingankar, S., Chen, Y., Moffa, A., Goldring, M. B., Sano, K., Jin-Hua, P., Sali, A., Goding, J., and Terkeltaub, R. (1999). Differential mechanisms of inorganic pyrophosphate production by plasma cell membrane glycoprotein-1 and B10 in chondrocytes. Arthritis Rheum. 42, 1986–1997. 295. Ea, H. K., and Liote, F. (2004). Calcium pyrophosphate dihydrate and basic calcium phosphate crystal-induced arthropathies: update on pathogenesis, clinical features, and therapy. Curr. Rheumatol. Rep. 6, 221–227. 296. Howell, D. S., Martel-Pelletier, J., Pelletier, J-P., Morales, S., and Muniz, O. (1984). NTP pyrophosphohydrolase in human chondrocalcinotic and osteoarthritic cartilage II Further studies on histologic and subcellular distribution. Arthritis Rheum. 27, 193–199. 297. Hirose, J., Ryan, L. M., and Masuda, I. (2002). Up-regulated expression of cartilage intermediate-layer protein and ANK in articular hyaline cartilage from patients with calcium pyrophosphate dihydrate crystal deposition disease. Arthritis Rheum. 46, 3218–3229. 298. Sohn, P., Crowley, M., Slattery, E., and Serra, R. (2002). Developmental and TGF-beta-mediated regulation of Ank mRNA expression in cartilage and bone. Osteoarthritis Cartilage 10, 482–490. 299. Terkeltaub, R. A. (2001). Inorganic pyrophosphate generation and disposition in pathophysiology. Am. J. Physiol. Cell. Physiol. 281, C1-C11. 300. Hirose, J., Masuda, I., and Ryan, L. M. (2000). Expression of cartilage intermediate layer protein/nucleotide pyrophosphohydrolase parallels the production of extracellular inorganic pyrophosphate in response to growth factors and with aging. Arthritis Rheum. 43, 2703–2711. 301. Rosenthal, A. K., Masuda, I., Gohr, C. M., Derfus, B. A., and Le, M. (2001). The transglutaminase, factor XIIIA, is present in articular chondrocytes. Osteoarthritis Cartilage 9, 578–581. 302. Rosenthal, A. K., Mosesson, M. W., Gohr, C. M., Masuda, I., Heinkel, D., and Seibenlist, K. R. (2004). Regulation of transglutaminase activity in articular chondrocytes through thrombin receptor-mediated factor XIII synthesis. Thromb. Haemost. 91, 558–568. 303. Johnson, K., Hashimoto, S., Lotz, M., Pritzker, K., and Terkeltaub, R. (2001). Interleukin-1 induces pro-mineralizing activity of cartilage tissue transglutaminase and factor XIIIa. Am. J. Pathol. 159, 149–163. 304. Ho, A. M., Johnson, M. D., and Kingsley, D. M. (2000). Role of the mouse ank gene in control of tissue calcification and arthritis. Science 289 (5477), 265–270. 305. Okawa, A., Nakamura, I., Goto, S., Moriya, H., Nakamura, Y., and Ikegawa, S. (1998). Mutation in Npps in a mouse model of ossification of the posterior longitudinal ligament of the spine. Nat. Genet. 19, 271–273. 306. Pendleton, A., Johnson, M. D., Hughes, A., Gurley, K. A., Ho, A. M., Doherty, M., Dixey, J., Gillet, P., Loeuille, D., McGrath, R., Reginato, A., Shiang, R., Wright, G., Netter, P., Williams, C., and Kingsley, D. M. (2002). Mutations in ANKH cause chondrocalcinosis. Am. J. Hum. Genet. 71, 933–940. Epub 2002 Sep 20. 307. Williams, C. J., Pendleton, A., Bonavita, G., Reginato, A. J., Hughes, A. E., Peariso, S., Doherty, M., McCarty, D. J., and Ryan, L. M. (2003). Mutations in the amino terminus of ANKH in two US families with calcium pyrophosphate dihydrate crystal deposition disease. Arthritis Rheum. 48, 2627–2631.

DANIEL-HENRI MANICOURT ET AL. 308. Wendling, D., Jeannin-Louys, L., Kremer, P., Fellmann, F., Toussirot, E., and Mornet, E. (2001). Adult hypophosphatasia. Current Aspects Joint Bone Spine 68, 120–124. 309. Rosenberg, K., Olsson, H., Morgelin, M., and Heinegard, D. (1998). Cartilage oligomeric matrix protein shows high affinity zinc-dependent interaction with triple helical collagen. J. Biol. Chem. 273, 20397–20403. 310. Mabuchi, A., Manabe, N., Haga, N., Kitoh, H., Ikeda, T., Kawaji, H., Tamai, K., Hamada, J., Nakamura, S., Brunetti-Pierri, N., Kimizuka, M., Takatori, Y., Nakamura, K., Nishimura, G., Ohashi, H., and Ikegawa, S. (2003). Novel types of COMP mutations and genotype-phenotype association in pseudoachondroplasia and multiple epiphyseal dysplasia. Hum. Genet. 112, 84–90. Epub 2002 Oct 29. 311. Lorenzo, P., Bayliss, M. T., and Heinegard, D. (2004). Altered patterns and synthesis of extracellular matrix macromolecules in early osteoarthritis. Matrix Biol. 23, 381–391. 312. Vilim, V., Vytasek, R., Olejarova, M., Machacek, S., Gatterova, J., Prochazka, B., Kraus, V. B., and Pavelka, K. (2001). Serum cartilage oligomeric matrix protein reflects the presence of clinically diagnosed synovitis in patients with knee osteoarthritis. Osteoarthritis Cartilage 9, 612–618. 313. Skoumal, M., Kolarz, G., and Klingler, A. (2003). Serum levels of cartilage oligomeric matrix protein. A predicting factor and a valuable parameter for disease management in rheumatoid arthritis. Scand. J. Rheumatol. 32, 156–161. 314. Crnkic, M., Mansson, B., Larsson, L., Geborek, P., Heinegard, D., and Saxne, T. (2003). Serum cartilage oligomeric matrix protein (COMP) decreases in rheumatoid arthritis patients treated with infliximab or etanercept. Arthritis Res. Ther. 5, R181–R185. Epub 2003 Apr 29. 315. Jordan, J. M. (2004). Cartilage oligomeric matrix protein as a marker of osteoarthritis. Rheumatol. Suppl. 70, 45–49. 316. Manicourt, D. H., Poilvache, P., Nzeusseu, A., van Egeren, A., Devogelaer, J. P., Lenz, M. E., and Thonar, E. J. (1999). Serum levels of hyaluronan, antigenic keratan sulfate, matrix metalloproteinase 3, and tissue inhibitor of metalloproteinases 1 change predictably in rheumatoid arthritis patients who have begun activity after a night of bed rest. Arthritis Rheum. 42, 1861–1869. 317. Mundermann, A., Dyrby, C. O., Andriacchi, T. P., and King, K. B. (2005). Serum concentration of cartilage oligomeric matrix protein (COMP) is sensitive to physiological cyclic loading in healthy adults. Osteoarthritis Cartilage 13, 34–38. 318. Bradley, K., North, J., Saunders, D., Schwaeble, W., Jeziorska, M , Woolley, D. E. (1996). Synthesis of classical pathway complement components by chondrocytes. Immunology 88, 648–656. 319. Johansen, J. S., Olee, T., Price, P. A., Hashimoto, S., Ochs, R. L., and Lotz, M. (2001). Regulation of YKL-40 production by human articular chondrocytes. Arthritis Rheum. 44, 826–837. 320. Johansen, J. S., Kirwan, J. R., Price, P. A., and Sharif, M. (2001). Serum YKL-40 concentrations in patients with early rheumatoid arthritis: relation to joint destruction. Scand. J. Rheumatol. 30, 297–304. 321. Gendelman, R., Burton-Wurster, N. I., MacLeod, J. N., and Lust, G. (2003). The cartilage-specific fibronectin isoform has a high affinity binding site for the small proteoglycan decorin. J. Biol. Chem. 278, 11175–11181. Epub 2002 Dec. 322. Wolf, A., Raiss, R. X., and Steinmeyer, J. (2003). Fibronectin metabolism of cartilage explants in response to the frequency of intermittent loading. J. Orthop. Res. 21, 1081–1089. 323. Lust, G., and Burton-Wurster, N. (1992). Fibronectin in osteoarthritis: comparison of animal and human diseases. In “Articular Cartilage and Osteoarthritis” (K. E. Kuettner, R. Schleyerbach, J. G. Peyron, and V. C. Hascall, eds), pp. 447–500, Raven Press, New York.

Chapter 25 Products of Cartilage Metabolism 324. Homandberg, G. A., Costa, V., and Wen, C. (2002). Fibronectin fragments active in chondrocytic chondrolysis can be chemically crosslinked to the alpha5 integrin receptor subunit. Osteoarthritis Cartilage 10, 938–949. 325. Pichika, R., and Homandberg, G. A. (2004). Fibronectin fragments elevate nitric oxide (NO) and inducible NO synthetase (iNOS) levels in bovine cartilage and iNOS inhibitors block fibronectin fragment mediated damage and promote repair. Inflamm. Res. 53, 405–412. Epub 2004 Aug 10.

449 326. Johnson, A., Smith, R., Saxne, T., Hickery, M., and Heinegard, D. (2004). Fibronectin fragments cause release and degradation of collagen-binding molecules from equine explant cultures. Osteoarthritis Cartilage 12, 149–159. 327. Homandberg, G. A., Hui, F. (1994). High concentrations of fibronectin fragments cause short-term catabolic effects in cartilage tissue while lower concentrations cause continuous anabolic effects. Archives Biochem. Biophys. 311, 213–218.

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Chapter 26

Fluid Dynamics of the Joint Space and Trafficking of Matrix Products Peter A. Simkin, M. D.

Professor of Medicine, Adjunct Professor of Orthopaedics, University of Washington

I. Introduction II. Interpretation of Marker Data and Strategies for Dealing with Them

III. Conclusion References

I. INTRODUCTION

subchondral bone. Although this assumption clearly deserves more intensive study, particularly in the setting of inflammatory joint disease, it will be accepted for the purpose of this chapter. It means that any product of cartilaginous injury or degradation must leave that tissue by passing into the overlying synovial fluid. Sampling of that fluid and measurements of its marker content may therefore provide insights into the extent of the lesion. Further, serial observations may help evaluate the healing response to injury and the effectiveness of therapeutic interventions. Since synovial fluid is not always accessible and markers are subsequently cleared from the joint into plasma, it has also been thought that simple measurements in blood may have even greater value. Whether in synovial fluid or in serum, measurements of concentration comprise virtually all of the published marker data. How useful are such static findings in this book about Dynamics of Bone and Cartilage Metabolism? All synovial fluid solutes are molecules in transit. Some move mainly from plasma into the joint (i.e. glucose and oxygen), others move mainly from the joint back into

A large and growing literature attests to the broad, continuing interest in cartilage-derived molecules as “markers” of damage and repair across a wide variety of joint diseases [1–5]. Specific markers and the rationale for their use are examined in detail elsewhere in this volume (see Chapters 24 and 25). This chapter will aim to put such data in the context of normal and pathologic joint physiology. In so doing, we will introduce important caveats in the interpretation of marker data and discuss potential strategies for dealing with them.

II. INTERPRETATION OF MARKER DATA AND STRATEGIES FOR DEALING WITH THEM The underlying assumptions are straightforward. Both in normal and in abnormal joints, articular cartilage is thought to be backed by an impermeable barrier of Dynamics of Bone and Cartilage Metabolism

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452 plasma (i.e. lactate, carbon dioxide, and cartilaginous “markers”), and most simply move back and forth in the ongoing equilibrium which characterizes any specific compartment of the body’s extracellular space. When we aspirate an aliquot of synovial fluid, we arrest this process. Measurements in such aspirates characterize a moment in the joint just as a still frame freezes but a moment in a moving picture. To extend the analogy, a single photograph of a busy avenue would reveal measurable numbers of pedestrians and automobiles. No matter how carefully one counted, however, such a snapshot could tell little or nothing about how fast each individual moved, how great was the overall traffic flow, or where everyone was going. If one’s interest was narrower, as in police cars or men in overcoats, the photo would reveal even less. To begin to understand what is really going on, one must incorporate time in the measurements and evaluate the differing rates that characterize each participant in the scene. This limitation poses a fundamental problem for those interested in specific synovial fluid solutes and underscores the importance of interpreting the findings of aspirates within a dynamic context. In considering the dynamics of large synovial solutes, the most relevant data have come not from cartilaginous markers but from studies of serum albumin. Such data are useful primarily for the perspective they provide on synovial lymphatic outflow. Small molecules (roughly those having molecular weights less than 10 000) are thought to diffuse directly into plasma across the fenestrae of synovial blood vessels and are therefore cleared more rapidly from the joint. For larger molecules, however, the sole route of egress is thought to be convective transport through lymphatics. These valved vessels lie within the synovial interstitium. There, they take up interstitial fluid, including all of its dissolved solutes, and cyclic joint usage then provides the pump that sends it on its way back to the bloodstream. That path has classically been thought to end at the thoracic duct, but subsequent evidence suggests that vascular access may also occur through blood vessels perfusing regional lymph nodes [6]. Albumin kinetics have been studied in joints primarily through studies using radiolabeled molecules [7–9]. Briefly, serum albumin is tagged in vitro with radioiodide and a precisely quantified dose is injected into the joint space. Serial gamma counts are then obtained over the joint for a period of hours and their log values are plotted against time. Such studies usually yield a highly linear declining function whose slope can be readily expressed either as a simple rate constant (in min−1) or as a half-life (in minutes)(Fig. 1). At the conclusion of the experiment, the joint space is aspirated and the residual labeled albumin is quantified in a measured volume of synovial fluid.

Peter A. Simkin

Figure 1 Kinetics of albumin removal from a human knee. Radiolabeled albumin (RISA) was injected at time 0 and followed serially by external counting (∆) and in synovial fluid aspirates (䊐). After a brief equilibration period, both curves decline at the same constant rate which may be expressed as a half life or in min−1. The latter rate constant may be multiplied by the volume of distribution to provide a clearance value in ml/min. The more rapid curve labeled 123I illustrates removal of free iodide which was followed concurrently. (Wallis WJ, Simkin PA, Nelp WB, Foster DM. Intraarticular volume and clearance in human synovial effusions. Arthritis Rheum. 28: 441–449, 1985.)

It is then a simple matter to take the initial counts injected, correct that number for the fraction known to have left the joint and the physical decay of the isotope, and calculate the distribution volume of the remainder by mass balance. The product of that volume (in ml) and the removal rate constant (in min−1) provides a clearance rate for albumin (in ml/min) that is considered the best available measure of articular lymphatic flow [8–10]. Proteins up to the size of IgM macroglobulin have been found to leave human knees at the same rate as albumin [11–13]. This means that one may multiply the concentration of such a protein (in, for instance, mg/ml) by the albumin clearance (in ml/min) to calculate a flux rate for the protein of interest (mg/min). Since most joints would presumably be studied under steady-state condition, the flux out should equal the flux in. This influx rate is of great potential interest in converting marker studies from a static to a dynamic basis. The influx into the joint of a solely cartilage-derived molecule should provide a true indicator of “release” and thus of the rate of cartilage catabolism. Similarly, of course, the product of the distribution volume of albumin and the concentration of a comparable protein should provide an appropriate estimate of the intra-articular mass of that protein. Previous investigators interested in marker mass have based their calculations on the synovial fluid volume, either that recovered by aspiration, that measured by indicator dilution, or a combination of the two [14, 15]. It seems most likely

Chapter 26 Fluid Dynamics of the Joint Space and Trafficking of Matrix Products

however, that any marker found in the synovial fluid will also be distributed through the synovial interstitial tissue. Studies with radiolabeled albumin invariably show an articular distribution volume that is larger than that of synovial fluid. Our pilot studies also found that this larger volume did not change between 24 and 72 hours (see Fig. 1). These findings are consistent with the concept of a synovial interstitial volume that is limited by a functionally impermeable capsule and that remains in passive equilibrium with a smaller volume of included synovial fluid. It is the larger, interstitial volume that should be considered in estimating the mass and/or the dynamics of an intra-articular solute (Fig. 2). Figure 2 also illustrates some of the inherent problems when concentrations are measured not in synovial fluid but in the far more accessible plasma. If a given arthritic joint is indeed in the steady state, the rate of cartilage products released into the joint will be equalled by the rate at which those same products are passed on to the plasma pool. There, however, these products will be admixed with and diluted by the same products from all other joints and tissues of the body. This major problem limits the hope that plasma values will be useful in interpreting the degree of involvement in any single joint. The plasma pooling of markers could be seen as an advantage in assessments of therapeutic response in a polyarticular process such as rheumatoid arthritis. Two additional concerns must be addressed, however, in any

Figure 2 In a hypothetical knee (joint A), cartilage matrix turnover reflects the balance between ongoing synthesis and loss. “Lost” molecules enter the synovial fluid and equilibrate throughout the synovial interstitium. Degradation, which can potentially occur at many sites, produces putative “marker” molecules that are cleared from the joint into plasma by the lymphatics. In plasma, the markers mix with like molecules from other joints as well as from nonarticular sources. From plasma, markers may redistribute into other interstitial spaces, may be further processed (primarily in the liver), or may be excreted into the urine. The marker literature consists mainly of concentration measurements in synovial fluid and plasma. (Simkin PA, Bassett JE. Cartilage matrix molecules in serum and synovial fluid. Current Opin. Rheumatol. 7: 346–351, 1995.)

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such attempt. First is the fact that most hyaline cartilage is nonarticular and many “cartilaginous” molecules are also found in tissues other than cartilage such as the synovial matrix. Input from these sources will obviously dilute the interpretation of plasma findings as indicators of wellbeing in articular cartilage. The second and more important concern is that concentrations in plasma, like those in synovial fluid, are just as dependent on removal as they are on input. Most of this removal is thought to be hepatic but some of it may be renal, some may be redistribution into other tissue compartments, and we presently know very little about how these rates vary between individuals or within the same person over time. This concern is potentially addressable, but plasma concentrations will remain of limited value until we can place them in a meaningful kinetic context. These concerns can be illustrated by some of the findings to date on hyaluronan. This polymeric disaccharide enters the plasma from many different tissues with indirect evidence indicating that the peritoneum may be quantitatively the most important. Local kinetics vary greatly among tissues with the posterior chamber of the eye having perhaps the slowest turnover (t1/2 ≅ 70 h), the anterior chamber the fastest (t1/2 ≅ 1 h) and other tissues, including the synovium, falling somewhere in between. Entry into plasma from joints is accelerated by local inflammation and is also related clearly to the timing and extent of physical activity. Hyaluronan degradation may occur locally in tissues or subsequently in the liver. There, the rate of removal from the blood is size-dependent with larger polymers being taken up more rapidly than shorter ones. The complexity of these factors illustrates some of the formidable difficulties inherent in any attempt to use plasma measurements of hyaluronan to provide meaningful information about what is happening in any specific joint. Unfortunately, the same concern applies to most plasma measurements of markers derived from cartilage. In practice, the articular methodology shows clearly that the clearance of radiolabeled albumin is significantly faster from the knees of patients with rheumatoid arthritis than it is from patients with osteoarthritis. This was true both in the initial series from Seattle and in a large study from London that focused on corticosteroid effects on hyaluronan levels [7, 8, 16]. Mean values were the same in both studies: 0.07 ml/min in RA (and in ankylosing spondylitis) and 0.04 ml/min in OA. In the British study, intra-articular corticosteroids were administered to all subjects and the study was repeated after 2 weeks and again after 2 months. Albumin clearance fell to mean rates of 0.03 and 0.04 ml/min at these intervals in RA but remained steady at 0.04 ml/min in the OA patients.

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Peter A. Simkin Table I.

Clearance of Labeled Albumin from Arthritic Knees of Human Patients*

Ref

Conditions

n

CRISA

CRISA 2 wks/steroid

7 7 16 16 16

OA RA OA RA AS

11 9 8 8 6

0.039 ± 0.010 0.071 ± 0.008 0.04 ± 0.01 0.07 ± 0.01 0.07 ± 0.02

n.d. n.d. 0.04 ± 0.01 0.03 ± 0.01 0.04 ± 0.01

CRISA 2 months/steroid n.d. n.d. 0.04 ± 0.01 0.04 ± 0.01 0.03 ± 0.01

*CRISA is clearance of radioiodinated serum albumin in ml/min. OA, RA, and AS are osteoarthritis, rheumatoid arthritis, and ankylosing spondylitis. Values are in ml/min (mean SEM).

Patients with ankylosing spondylitis were also studied and their albumin clearance also fell markedly after corticosteroid injections. Thus, the available evidence in humans indicates that albumin clearance (which can be considered as a measure of lymphatic drainage) is accelerated in inflammatory disease and is markedly diminished when the inflammation is effectively treated (Table I). Comparable findings have been shown in experimental animals. Using both intra-articular pyrophosphate crystals and anterior cruciate ligament resection, Myers et al. have found that inflammation markedly accelerates the clearance of albumin from canine knees [17, 18]. This increase (estimated by comparison with studies of the unmanipulated, contralateral knee) was by approximately 20-fold in the intense inflammation induced by large amounts of crystals. It was also highly significant, however, when the inflammatory response was low grade as it was with small amounts of crystals and in a common surgically induced model of osteoarthritis (Table II). In unpublished work, we have used IL-1α to induce synovitis

in caprine carpal and tarsal joints and in each case the clearance of albumin from inflamed joints was substantially greater than when the same joints were studied under control conditions, both before and after the acute experimental inflammation [19]. Thus, in animals as well as in studies of people with arthritis, the articular inflammatory response includes a significant acceleration of protein clearance. These rates of protein clearance are essential determinants of the concentration of every synovial solute. In fact, the rate of removal from the joint is every bit as critical to the intrasynovial concentration as is the rate of entry. This simple fact has obvious applications for the interpretation of marker studies. A number of these investigations have, for instance, compared synovial fluid concentrations found in rheumatoid arthritis with those found in osteoarthritis. Since the clearance from RA knees is approximately twice that found in OA, a finding of comparable SF concentrations would mean that the release rate into the RA knee was not the same but twice

Table II. Albumin Clearance and Volume Determinations in Canine Knees with and without Experimental Joint Disease* Ref

Condition

n

Va (ml)

Vd (ml)

17

500 µg CPPD 0.5 µg CPPD 0.05 µg CPPD contra cruciate contra

3

33.7

2.2 ± 0.6

36.6 ± 11.0

3

6.7

1.4 ± 1.1

6.4 ± 2.1

6

2.7

0.8 ± 0.6

3.8 ± 1.1

13 6 6

1.5 3.8 1.4

0.2 ± 0.2 1.1 ± 0.9 0.1 ± 0.1

2.7 ± 0.1 9.2 ± 3.6 3.7 ± 0.9

17 17 17 18 18

*CRISA is the clearance of radioiodinated serum albumin. CPPD is the mass of calcium phyrophosphate dihydrate crystals injected. cruciate – indicates surgical transection of the anterior cruciate ligament. contra is the contralateral knee in each series of animals. Va is the aspirated volume of synovial fluid. Vd is the distribution volume of labeled albumin. Values are mean ± S.D.

CRISA (ml/min)

Chapter 26 Fluid Dynamics of the Joint Space and Trafficking of Matrix Products

as great. Similarly, an RA marker concentration after a corticosteroid injection that was unchanged from the preinjection value would mean that the release of that marker had doubled. This is true because the articular clearance after injections is halved. These simple illustrations show that it is, and will always be, extremely dangerous to use marker concentrations to draw conclusions about differences between diseases, responses to therapy or any other comparison between joints, unless the clearance kinetics have also been assessed. Note that the critical determinants are the clearance rates (in ml/min). The flux rate (mg/min) is not critical because the flux in will always equal the flux out in the steady state. The intrasynovial volume is also not critical in determining concentration. A constant input of a marker molecule will rapidly lead to a steady concentration when the SF volume is small and will take considerably longer when the volume is large. The end-point steady-state concentration will be the same, however, regardless of the effusion volume (Fig. 3). One of the most disappointing aspects of marker studies to date has been the wide range of SF concentrations observed for virtually every cartilaginous marker in aspirates from virtually every diagnostic category. Values spanning two or more orders of magnitude are more the rule than the exception and this broad range has greatly limited the practical application of the method. Much of this variation, of course, reflects lack of homogeneity in

Figure 3 Effect of volume (ml) and clearance rate (ml per unit of time) on the solute concentration (U/ml) in a well-mixed knee receiving solute at a constant rate of 1 U per unit of time. The knee of patient A has a volume of 10 ml, whereas the knee of patient B has a volume of 100 ml. When both knees are cleared at a constant rate of 2 ml per unit of time, they both attain the same solute concentration of 0.5 U/ml, although the larger volume requires much more time to reach equilibrium. The knee of patient C also has a volume of 100 ml and receives material at the same rate. However, it is cleared at twice the rate in patient B (4 ml per unit of time) and therefore goes to an equilibrium solute concentration half as great. This figure illustrates that steady-state concentrations of any solute reflect the balance of input and output and are independent of volume. (Simkin PA, Bassett JE. Cartilage matrix molecules in serum and synovial fluid. Current Opin. Rheumatol. 7: 346–351, 1995.)

455

any patient group. Among rheumatoid subjects, for instance, both sexes are likely to be represented and there will always be a significant range of patient age, of disease duration, of effusion duration, of therapeutic measures in use, and in the intensity of the inflammatory response. Each of these factors may play a role in the intrasynovial input or clearance of specific marker molecules. Some of the observed variation must reflect real differences in marker release that carry important information about the condition and the prognosis of the studied joint. One may hope that quantification of lymphatic outflow, by allowing conversion of static SF concentrations to dynamic marker release rates, will help to sort out the logical mechanisms that underlie the current diversity of findings. The foregoing discussion has focused on proteins “comparable” to albumin. Many of the intrasynovial molecules of greater current interest do not fit this description. As already mentioned, solutes with molecular weights of 10 000 or less will be cleared not only by lymphatics but will also pass through microvascular pores and be cleared directly into the venous circulation. Since the rate of synovial plasma flow greatly exceeds that of lymphatic drainage, venous clearance could easily accelerate the clearance of IL-1 or TNF-α, for instance, by an order of magnitude compared to that of albumin. As yet, there are no available data that allow us to estimate the passive transport rates for such solutes. Obviously, the specific cellular uptake of such cytokine molecules adds huge additional complications to any consideration of their articular dynamics. At the other end of the size spectrum, markers released from cartilage may not resemble the compact, globular structure of albumin but may instead be much larger and more linear strands of aggrecan (or its catabolities), of type II collagen, or of other cartilage constituents. It is now clear that such macromolecules do not leave the joint as readily as does albumin. This issue was addressed by Page-Thomas et al. who found the mean clearance t1/2 for large proteoglycan molecules (Mr = 2.5 × 106) leaving the rabbit knee to be 12.5 hours, in contrast to 3.9 hours for albumin in the same joints [20]. A more striking difference was reported by Coleman et al., who also used rabbit knees but studied hyaluronan kinetics with a different experimental approach. Their mean t1/2 values were 26.3 h for hyaluronan and 1.23 h for albumin [21]. These findings lead to an inescapable conclusion: marker molecules, which are large in mass and/or configuration, are partially reflected back into the joint space as fluid flows into terminal lymphatics through the interstitial matrix of the synovium. More work remains to be done in defining just what are the critical molecular dimensions, in evaluating

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Peter A. Simkin

the effects of local inflammation on this sieving function of the synovium and in distinguishing between the fractions that are cleared into lymphatics, reflected back into the joint, or locally degraded. That work will be facilitated by Levick’s simple approach comparing half-lives of markers to that of albumin in order to quantify the extent of reflection [23].

III. CONCLUSION To date, studies of cartilaginous markers have been limited primarily to measurements of concentration in synovial fluid and in plasma. When such data are used to compare different conditions or to follow the same condition serially, the work necessarily assumes that the clearance does not vary between conditions, between individuals, or within the same individual over time. Unfortunately, those assumptions are demonstrably false. To fulfill the marker promise of information regarding “release” from cartilage, the kinetics of these molecules must be examined together with their concentration. Fortunately, relatively simple techniques, largely based on concurrent studies of serum albumin as a reference solute, may go a long way to remedy this situation.

References 1. Garnero, P., and Delmas, P. (2003). Biomarkers in osteoarthritis. Curr. Opin. Rheumatol. 15, 641–646. 2. Poole, A. R. (2003). Biochemical/immunochemical biomarkers of osteoarthritis: utility for prediction of incident or progressive osteoarthritis. Rheum. Dis. Clin. North Am. 29, 803–818. 3. Bruyere, O., Collette J. H., Ethgen, O., Rovati, L. C., Giacovelli, G., Henrotin, Y. E., Seidel, L., and Reginster, J. Y. (2003). Biochemical markers of bone and cartilage remodeling in prediction of longterm progression of knee osteoarthritis. J. Rheumatol. 30, 1043–1050. 4. Lohmander, L.S., and Felson, D. (2004). Can we identify a ‘high risk’ patient profile to determine who will experience rapid progression of osteoarthritis? Osteoarthritis Cartilage 12(Suppl A), S49–S52. 5. Simkin, P. A., and Bassett, J. E. (1995). Cartilage matrix molecules in serum and synovial fluid. Curr. Opin. Rheumatol. 7, 346–351.

6. Rayan, I., Thonar, E. J. M. A., Chen, L. M., Lenz, M. E., and Williams, J. M. (1998). Regional differences in the rise in blood levels of antigenic keratan sulfate and hyaluronan after chymopapain induced knee joint injury. J. Rheumatol. 25, 521–526. 7. Wallis, W. J., Simkin, P. A., Nelp, W. B., and Foster, D. M. (1985). Intraarticular volume and clearance in human synovial effusions. Arthritis Rheum. 28, 441–449. 8. Wallis, W. J., Simkin, P. A., and Nelp, W. B. (1987). Protein traffic in human synovial effusions. Arthritis Rheum. 30, 57–63. 9. Simkin, P. A., and Benedict, R. S. (1990). Iodide and albumin kinetics in normal canine wrists and knees. Arthritis Rheum. 33, 73–79. 10. Levick, J. R., and McDonald, J. N. (1995). Fluid movement across synovium in healthy joints: role of synovial fluid macromolecules. Ann. Rheum. Dis. 54, 417–423. 11. Rodnan, G. P., and MacLachlan, M. J. (1960). The absorption of serum albumin and gammaglobulin from the knee joint of man and rabbit. Arthritis Rheum. 3, 152–157. 12. Sliwinski, A. F., and Zvaifler, N. J. (1969). The removal of aggregated and nonaggregated autologous gamma globulin from rheumatoid joints. Arthritis Rheum. 12, 504–514. 13. Weinberger, A., and Simkin, P. A. (1989). Plasma proteins in synovial fluids of normal human joints. Semin. Arthritis Rheum. 19, 66–76. 14. Geborek, P., Saxne, T., Heinegard, D., and Wollheim, F. A. (1988). Measurement of synovial fluid volume using albumin dilution upon intraarticular saline injection. J. Rheumatol. 15, 91–94. 15. Delecrin, J., Oka, M., Kumar, P., Takahashi, S., Kotoura, Y., Yamamuro, T., and Daculsi, G. (1992). Measurement of synovial fluid volume: a new dilution method adapted to fluid permeation from the synovial cavity. J. Rheumatol. 19, 1746–1752. 16. Pitsillides, A. A., Will, R. K., Bayliss, M. T., and Edwards, J. C. W. (1994). Circulating and synovial fluid hyaluronan levels: effects of intraarticular corticosteroid on the concentration and the rate of turnover. Arthritis Rheum. 37, 1030–1038. 17. Myers, S. L., Brandt, K. D., and Eilam, O. (1995). Even low-grade synovitis significantly accelerates the clearance of protein from the canine knee. Implications for measurement of synovial fluid “markers” of osteoarthritis. Arthritis Rheum. 38, 1085–1091. 18. Myers, S. L., O’Connor, B. L., and Brandt, K. D. (1996). Accelerated clearance of albumin from the osteoarthritic knee: implications for interpretation of concentrations of “cartilage markers” in synovial fluid. J. Rheumatol. 23, 1744–1748. 19. Simkin, P. A., and Bassett, J. E. (unpublished). 20. Page-Thomas, D. P., Bard, D., King, B., and Dingle, J. T. (1987). Clearance of proteoglycan from joint cavities. Ann. Rheum. Dis. 46, 934–937. 21. Coleman, P. J., Scott, D., Ray, J., Mason, R. M., and Levick, J. R. (1997). Hyaluronan secretion into the synovial cavity of rabbit knees and comparison with albumin turnover. J. Physiol. 503(Pt 3), 645–656. 22. Levick, J. R. (1998). A method for estimating macromolecular reflection by human synovium, using measurements of intra-articular half lives. Ann. Rheum. Dis. 57, 339–344.

Chapter 27

Transgenic Models of Bone Disease Barbara E. Kream John R. Harrison

Departments of Medicine and Genetics and Developmental Biology, University of Connecticut Health Center, Farmington, CT 06030 Division of Orthodontics, University of Connecticut Health Center, Farmington, CT 06030

I. Introduction II. Generation of Mouse Models III. Transgenic Models in Bone Biology

IV. Perspectives and Future Directions References

I. INTRODUCTION

II. GENERATION OF MOUSE MODELS

The ability to manipulate the mouse genome has proven invaluable for the development of models to understand gene function in vivo [1]. Several excellent and comprehensive reviews have been published on the development and use of transgenic models of bone and mineral metabolism and diseases [2–5]. The intent of this chapter is to focus on recent work in this area, including a discussion of methods currently used to manipulate gene expression in the mouse. We will highlight selected examples of how transgenic models have provided insights into the roles of transcription factors and hormone signaling pathways in normal bone biology and the pathophysiology of bone diseases. Finally, new technological developments are discussed that will influence the future of functional genomics in the mouse.

A. Transgenesis

Dynamics of Bone and Cartilage Metabolism

The overexpression of genes in transgenic mice has become a widely used technique for understanding the role of genes in vivo. Transgenesis in the mouse provides a means of over-expressing genes of interest to generate gain- and loss-of-function models. Transgenesis has been traditionally accomplished by the microinjection of DNA constructs into the single-cell fertilized embryos [6]. Transgenes typically contain a fragment of a gene promoter fragment positioned upstream of a gene of interest, which can be a reporter, a functional gene, or a dominant negative molecule. The transgene can be epitope-tagged to distinguish it from its endogenous counterpart. Alternatively, the availability of species-specific antibodies to functionally 457

Copyright © 2006 by Academic Press. All rights of reproduction in any form reserved.

458 equivalent transgene products derived from a different species can allow their detection in mouse tissues and blood. Transgenes integrate randomly into the host genome, in one or a few different sites, usually as concatamers in which multiple copies are all oriented in the same direction. Transgene expression is typically dependent on the integration site but not on the copy number unless sequences are present in the transgenic construct to insulate it from the influence of its surrounding chromosomal environment. Both ubiquitous and tissue-specific promoters have been used for targeting transgenes to osteoblasts and osteoclasts. Transgenes driven by ubiquitous promoters may have global effects in the mouse model. Therefore, discernible bone phenotypes may be caused directly by transgene expression in bone cells or indirectly by effects on other tissues. Even with relatively tissue-specific promoters, low-level expression in other cell types may be sufficient to perturb normal function of various organs. Some transgenes, particularly those for secreted proteins, may have non-cell autonomous effects on neighboring nontargeted cells. The overexpression of an endogenous gene provides a gain-of-function model in which there is an exaggeration of the normal activity of a gene product. Expression of a dominant negative molecule, on the other hand, can be used to generate a loss-of-function model by disrupting the role of the normal gene product. The disadvantages of overexpression of genes by transgenesis include generating levels of a gene product that far exceeds its physiological level. This can cause phenotypes that are unexpected and/or do not represent normal physiological function. Moreover, transgenes may be misexpressed both temporally and spatially. Because the promoters used to target transgenes usually contain only a small fragment of the regulatory sequences in the gene, the transgene can be expressed in cells that normally do not express the endogenous gene. Ideally, to build a bone cell-specific transgenic model, expression of the promoter should closely mimic expression of the endogenous gene. Moreover, the transgene should be specifically expressed in the targeted cell and at a level at least as high as physiological. The promoters used to target transgene expression to bone cells have typically been those of matrix genes, which are selectively or specifically expressed in bone [5]. Type I collagen promoters have been used frequently to drive transgenes in cells of the osteoblast lineage. The endogenous Col1a1 gene is highly expressed in cells of the osteoblast lineage and serves as an early marker of osteoblast differentiation. The isolation and characterization of the rat [7] and mouse [8] Col1a1 promoters has provided the tools to drive transgenes in osteoblasts. Promoter fragments that have been commonly used include a rat 3.6-kb fragment of the rat Col1a1 gene and a 2.3-kb fragment of the rat or mouse

BARBARA E. KREAM AND JOHN R. HARRISON

Col1a1 genes [9, 10]. The rat 3.6-kb Col1a1 promoter, while highly expressed in cells of the osteoblast lineage, is not specific for bone since it is activated in a number of other tissues such as tendon, skin, and lung [11]. Studies of the spatial and temporal pattern of Col1a1 promoter– reporter constructs have provided crucial information for developing transgenic models in which genes are targeted to bone cells. These studies showed that a 3.6-kb Col1a1 promoter fragment (Col3.6) is highly expressed in osteoblasts. Deletion of the promoter to 2.3-kb (Col2.3) does not affect promoter activity in vivo, although there is a reduction in activity in vitro [11]. Recently, the green fluorescent protein (GFP) reporter has been cloned downstream of these promoter fragments to determine the spatial activation of the Col3.6 and Col2.3 promoters in vivo [9]. The Col3.6 promoter is broadly expressed in cells of the osteoblast lineage, while the Col2.3 promoter is expressed in mature osteoblasts both in vivo and in primary calvarial and bone marrow stromal cells. Both promoters drive strong transgene expression in osteoblasts and odontoblasts [12]. The Col3.6 promoter is expressed in tendon, lung, and aorta, while the Col2.3 promoter has a more restricted pattern of expression in other tissues. The advantages of these Col1a1 promoter fragments include strong expression in osteoblasts and the ability to distinguish subpopulations of osteoblast lineage cells. Because of the fidelity of its bone-specific expression, the osteocalcin promoter provides an excellent tool for targeting genes to mature osteoblasts [13]. The proximal osteocalcin promoter contains sequences that confer tissuespecific and developmental stage-specific expression in mature osteoblasts [14, 15]. The expression of osteocalcin in skeletal tissues is highly restricted to differentiated osteoblasts and is not seen in preosteoblasts or early osteoblast progenitors. Osteocalcin expression is activated late in development in the perinatal period. However, the strength and hormonal regulation of osteocalcin promoter fragments can differ due to species differences [16] and can be affected by age and sex of the animals [17]. Lowlevel transgene expression may be an advantage for some studies in which it is critical to limit the degree of overexpression. However, for other systems such as the Cre-loxP model for generating conditional knockouts, Cre expression must be driven above a threshold level that allows the rearrangement of a floxed gene [18]. Recently, the dental matrix protein-1 (DMP-1) promoter has been fused to GFP and introduced into transgenic mice [19]. The DMP-1 promoter is strongly expressed in late-stage osteoblasts and osteocytes and will provide a tool for targeting genes to these cells. There have been fewer transgenic models in which genes have been targeted to osteoclasts; this may be due

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Chapter 27 Transgenic Models of Bone Disease

to the relative paucity of cloned promoter fragments that specifically target osteoclasts and their progenitors. The tartrate-resistant acid phosphatase and cathepsin K promoters are expressed in osteoclasts and other hematopoietic cells and have been used to drive Cre recombinase in osteoclasts [20]. The CD11b promoter has also been used to target Cre recombinase to cells of the hematopoietic myeloid-osteoclast lineage [21]. The CD11b promoter is expressed in mature osteoclasts arising from bone marrow and spleen progenitors [21]. Both models may be useful tools for generating conditional knockout models in cells of the osteoclast lineage.

B. Gene Knockout Models Genes can be inactivated by homologous recombination of targeting vectors in embryonic stem cells [1]. A critical region of the gene is replaced or disrupted by a cassette containing both positive and negative selection markers for the selection of homologous and nonhomologous recombinants, respectively. At the advent of this technology, the positive selection marker (usually the neomycin resistance gene) was often left in the targeted allele. However, since the neo gene can sometimes interfere with the function of nearby genes [22], it is now customary to remove it by flanking it with sites that allow it to be removed by bacterial recombinases. The advantages of a global gene knockout are that the function of the gene can be demonstrated in the target cells of interest. However, major disadvantages include the early embryonic lethality that often accompany global gene knockouts, compensation by another gene product during development and alterations of other organ systems that may indirectly affect bone. Considering these issues, the ability to ablate genes both temporally and spatially has been an important tool for understanding tissue-specific gene function [22]. Conditional gene knockouts can be accomplished using Cre-loxP technology [18]. Two different mouse strains are required: a transgenic line expressing Cre recombinase in the tissue of interest and a line in which the gene of interest has been engineered with lox P sites flanking a critical region. The breeding of the two mouse lines will lead to progeny having gene rearrangement in the cells expressing Cre. Several criteria should be met when developing a conditional knockout. Cre expression should be above the minimum threshold to cause gene rearrangement in the tissue of interest. The efficiency of Cre-mediated gene rearrangement should be such that gene expression is knocked down enough to cause a phenotype. To ensure sufficient gene knockdown in a conditional model, mice

can be developed that contain one floxed allele and one null allele for a gene of interest. Thus, prior to Cre-mediated gene rearrangement, expression of the gene of interest should be decreased by 50%. The temporal and spatial pattern of gene rearrangement can be determined using several Cre reporter strains, such as ROSA26 [23], Z/EG [24], and Z/AP [25]. These reporter strains provide historical marking of Cre expression and are useful tools for characterizing a newly developed Cre transgenic line. Disadvantages with the currently available Cre transgenic lines are the early and widespread activation of Cre recombinase in the embryo and expression of Cre that is so low or sporadic that gene rearrangement is inefficient. To circumvent detrimental effects during embryonic development and to precisely time gene disruption, inducible Cre transgenic systems can be generated [22]. A popular model is the tamoxifen-inducible system in which Cre recombinase is fused to a mutated ligand-binding domain of the ER and then cloned downstream of a targeting promoter [26, 27]. The Cre fusion protein, which is initially sequestered in the cytoplasm, becomes activated and translocates to the nucleus when 4-hydroxytamoxifen is given the animal (or to cells if an ex vivo experiment is performed). An inducible model has recently developed in which the murine 2.3-kb Col1a1 promoter has been used to drive a tamoxifen-inducible Cre transgene for temporally regulated gene ablation in osteoblasts and odontoblasts [28]. The binary tetracycline-dependent systems have enjoyed widespread use [29]. With the Tet-Off system, removal of tetracycline induces Cre expression. By contrast, with the Tet-On system, addition of tetracycline induces Cre expression. There are several limitations of inducible systems. It is important to test several transgenic lines to find one that confers high and inducible Cre expression. It is critical to determine whether there is leaky expression of Cre in the uninduced state. Finally, it should be determined whether the inducer itself has any unexpected or adverse effects on bone (this can be tested in wild-type mice).

C. Phenotypic Analysis A careful phenotypic analysis is required to fully understand and interpret results in a transgenic mouse model. Because transgenes can integrate randomly into the genome, it is important to examine multiple lines of transgenic mice to exclude the possibility that the observed phenotype is not caused by disruption of an endogenous gene at a fortuitous transgene insertion site. Validity of the phenotype is strengthened if a transgene dose-dependent phenotype can be demonstrated. Often, transgenic mice are studied

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as hemizygotes. Subtle phenotypes can be enhanced if hemizygotes are bred together to produce homozygous progeny. Although producing a homozygous line reduces the need for frequent genotyping, transgene homozygosity can often lead to inbreeding and poor breeding performance. It is important to determine whether transgenic mice are obtained in Mendelian ratios. The under-representation of transgenic offspring may indicate embryonic lethality, while the converse may indicate multiple transgene insertion sites. Finally, it is important to consider the background strain used to generate transgenic mouse lines. Phenotypes may be altered among different inbred strains due to the presence of modifier genes. On the other hand, measurements of phenotypic traits may be more variable in an outbred strain, thus requiring a larger sample size. It may be advantageous to develop a transgenic in an inbred background for initial characterization; then, the line can be easily outbred to examine the impact of the transgene in a heterogeneous genetic background. Southern blot analysis of a transgenic line will serve to unequivocally identify the line.

III. TRANSGENIC MODELS IN BONE BIOLOGY A. Hormones that Regulate Bone Formation A number of hormonal signaling pathways are critical for bone formation including the BMP [30] and Wnt/LRP [31–33] pathways. Various components of their signaling pathways, including receptors, ligands, and downstream signaling molecules, have been genetically modified in the mouse and the reader is directed to the reviews mentioned above. We have chosen to discuss in detail the insulin like-growth factor-I (IGF-I) system, as manipulation of the system in mice has led to a greater understanding of how they may regulate bone mass acquisition in humans. The insulin/IGF system consists of the polypeptides insulin, IGF-I and IGF-II, their cognate receptors, six IGF-binding proteins (IGFBP 1–6) and IGFBP proteases [34, 35]. IGF-I and IGF-II signaling through tyrosine kinase type I IGF receptors (IGF1R) leads to enhanced cell proliferation, differentiation, and survival in many tissues. In postnatal life, IGF-I is produced in liver under the control of growth hormone and then secreted into the blood to provide an endocrine source of growth factor. In addition, most other tissues produce IGF-I, and this component plays an important local role in tissue homeostasis [35]. There has been much interest in elucidating the role of IGF-I in bone remodeling since many human and animal

studies show an association between circulating IGF-I and bone mineral density [36]. To delineate the role of IGF-I in embryonic development, the murine Igf1 gene has been inactivated by targeted mutagenesis [37, 38]. Fetal mice with a complete knockout of the Igf1 gene are nearly 40% smaller than wild-type littermates and display developmental abnormalities of muscle, lung, skeleton, and other organ systems [37, 38]. Mice with total IGF-I deficiency have varying degrees of perinatal lethality depending on the genetic background [39]. The few Igf1 knockout mice that survive into adulthood show severe growth impairment along with abnormalities in many organ systems. Trabecular bone mass in the proximal tibia of Igf1 knockout mice is increased [40], although this might be due to increased circulating growth hormone or the lack of IGF-I-mediated resorption. We have found that mice heterozygous null for the Igf1 gene (HET) in an outbred strain show seemingly normal perinatal survival and fertility. HET mice had reduced body weight, IGF-I levels, femur length, and bone mineral density compared to their sex-matched wild-type littermates between 1 and 18 months of age. Analysis by microcomputed tomography showed reduced cortical bone width and periosteal circumference at 2 and 8 months of age [40a]. Similar results in young HET mice have been published [41]. To distinguish between the endocrine and paracrine roles of IGF-I, liver-specific Igf1 knockout models (LID) have been generated using Cre-loxP technology [42, 43]. LID mice do not have detectable IGF-I expression in liver and at least 75% less circulating IGF-I than control littermates. Surprisingly, LID mice have apparently normal fetal development, do not show perinatal lethality and are similar in size to control littermates [42, 43]. In additional studies, a thorough inspection of the skeletons showed LID mice to have shorter femurs and reduced periosteal circumference, indicative of impaired longitudinal bone growth and cortical bone apposition [44]. Interestingly, these changes are similar to those in HET mice (Kream, unpublished), which have a more modest, yet global, reduction in circulating IGF-I. These data may imply that local as well as circulating IGF-I is important for bone acquisition. A particularly informative study was the conditional knockout of the type 1 IGF receptor gene (Igf1r) in osteoblasts using an OC-Cre transgene. These mice had increased osteoid surface indicating that IGF-I plays an important role in mineralization [45]. Several transgenic models have been generated in which IGF-I overexpression is targeted to osteoblasts. The osteocalcin promoter was first used to express a human IGF-I cDNA to mature osteoblasts [46]. Osteocalcin-IGF

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transgenic mice have increased matrix apposition rate at 3 weeks and increased trabecular bone volume at 6 weeks of age. There were no differences in cortical bone width, osteoblast number, or osteoclast number at these time points and no changes in static or dynamic bone histomorphometric parameters at 24 weeks. In another model developed by our group, the Col3.6 promoter (including most of the Col1a1 first intron) was used to drive a FLAGtagged murine IGF-I cDNA more broadly in osteoblast lineage cells (Kream, unpublished). Transgenic calvaria had increased width, marrow spaces, and osteoclast number and surface. Notably, the calvarial phenotype was dependent on transgene expression and persisted with age. One line of mice had increased circulating IGF-I levels, presumably due to spillover from bone production. These mice showed an increase in femur length and mid-diaphyseal cortical width and area. In Col3.6-driven IGF-I transgenic mice, trabecular bone volume was not increased, which may have been due to increased resorption. The phenotypic differences between the two models (Col3.6-IGF and osteocalcin-IGF mice) are likely due to the distinct temporal and spatial expression pattern of the two promoters. The osteocalcin promoter is expressed in the perinatal period and is restricted to mature osteoblasts [13], while the Col3.6 promoter is expressed early during embryonic life and more broadly both spatially and temporally in osteoblast lineage cells [9]. Recently, several models have been produced in which various IGFBPs have been targeted to osteoblasts. Some IGFBPs act to sequester and inactivate IGFs, while others, such as IGFBP-5, may have IGF-independent effects on bone. Mice with constitutive transgenic IGFBP-3 overexpression are smaller and have reduced bone mineral density, reduced mineral apposition rates in cortical bone, and increased bone resorption markers [47]. Ex vivo osteoblast cultures showed that the decrease in bone formation may have been due to impaired proliferation [47]. Transgenic mice with constitutive overexpression of IGFBP-5 have decreased bone mineral density, with males being more severely affected than females [48]. Disparate effects of the transgene on bone formation rates and mineralizing surfaces were found on the periosteal (decreased) and endosteal (increased) surfaces [48]. Likewise, transgenic mice with osteocalcin promoter-driven IGFBP-5 overexpression had a transient decrease in femoral trabecular bone volume and mineral apposition rate [49, 50]. Since osteoblast number was not changed, osteoblast function may have been impaired; osteoclast number was not changed. These studies indicate that overexpression of IGFBPs decreases bone formation by impairing osteoblast number and/or function most likely by sequestering and

461 inactivating IGFs. However, some effects of transgenic IGFBP-5 may have been IGF-independent [48].

B. Cytokines that Regulate Bone Resorption Murine transgenic and knockout models have been integral in unraveling the complex network of cytokine signaling that controls osteoclast differentiation and bone resorption. Receptor activator of NF-κΒ (RΑΝΚL) is a member of the TNF ligand family that is expressed on the surface of osteoblast lineage cells and signals through the receptor activator of NF-κB (RANK) on the surface of hematopoietic osteoclast progenitors via a direct cell–cell interaction. RANK, a member of the TNF receptor family, signals the nuclear translocation of NF-κB, which activates a number of target genes that ultimately lead to osteoclast progenitor fusion, osteoclast differentiation, and survival. To allow this process to be tightly controlled, a novel secreted glycoprotein called osteoprotegerin (OPG), a soluble member of the TNF receptor family, is expressed by cells of the osteoblast lineage and acts as a decoy receptor for RANKL. The balance of RANKL and OPG expression in osteoblasts is therefore a major determinant of osteoclast differentiation and bone resorption, and a number of factors known to modulate bone resorption regulate their expression. OPG was the first member of this network to be discovered. A transgenic mouse model engineered for hepatic overexpression of OPG produced a nonlethal osteopetrotic phenotype in which osteoclasts failed to undergo terminal differentiation [51]. Conversely, OPG-null mice exhibited osteoporosis due to excessive osteoclastogenesis, resulting in a severe loss of trabecular bone, increased cortical porosity, and diminished mechanical bone strength [52, 53]. Min et al. [54] demonstrated that this osteoporotic phenotype could be rescued by systemic transgenic overexpression of OPG, which restored osteoclastogenesis. Similar genetic approaches subsequently established the roles of RANKL and its cognate receptor, RANK, in this system. Both RANKL- and RANK-deficient mice were found to be severely osteopetrotic, with a complete absence of osteoclasts, leading to profound accumulation of bone and a failure of tooth eruption [55, 56]. After the major cytokines responsible for osteoclast differentiation had been identified, attention shifted to the downstream cellular signaling pathways mediating RANK signaling. As its name implies, RANK is capable of activating NF-κB by inducing degradation of cytoplasmic IκB, allowing translocation of NF-κB complexes to the nucleus. The essential nature of NF-κB signaling in osteoclasts was

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underscored by the observation that mice deficient in the p50 and p52 subunits of NF-κB exhibited osteopetrosis [57, 58]. Subsequent studies showed that RANK-expressing osteoclast progenitors are present in p50/52 doubleknockout mice, but terminal differentiation into TRAPpositive osteoclasts was blocked [59]. The RANK receptor is coupled to NF-κB signaling by the TNF receptorassociated factor (TRAF) family of adapter proteins. Not long after the observation that NF-κB signaling was essential for osteoclastogenesis, a requirement for TRAF6 in this process was demonstrated by the finding that TRAF5deficient mice likewise exhibited the osteopetrotic/tooth eruption phenotype [60]. There is, however, both longstanding and new evidence that other signaling pathways are essential for osteoclast differentiation. For example, it has been known for more than 10 years that c-fos-null mice are osteopetrotic secondary to a lack of differentiated osteoclasts. New studies have shown that a nuclear factor of activated T cells (NFAT) is a target for Fos and Jun signaling in the osteoclast progenitor, and that expression of an activated form of NFAT can rescue osteoclast differentiation in c-fos-null progenitor cells exposed to RANKL [61, 62]. While much progress has been made on understanding the regulation of osteoclastogenesis using global gene ablation and overexpression strategies, such studies are complicated by a lack of specificity of many target genes to the osteoclast lineage. Clearly, more sophisticated models are required for targeted overexpression and conditional gene knockout strategies in osteoclasts and their progenitors. Toward this end, transgenic mouse models with osteoclastspecific Cre recombinase are being developed. Chiu et al. [20] have recently generated mice driving Cre expression under control of the osteoclast-specific TRAP and cathepsin K promoters and demonstrated expression specifically in bones and cartilage of a Cre-reporter mouse strain.

C. Nuclear Receptors Many steroid hormones, including estrogens, vitamin D, glucocorticoids, and several orphan receptors affect bone development and remodeling. Transgenic approaches to understanding the role of these hormones have included knockout and overexpression of their cognate receptors. The functions of estrogen in bone have been highly studied due to their role in postmenopausal osteoporosis. To determine the role of estrogens in bone metabolism, the estrogen receptors, ERα and ERβ, have been targeted in mice by several laboratories producing either complete or incomplete knockouts [63, 64]. Although there have been some discrepant results, there appear to be complex gender- and

species-specific phenotypes [64]. It seems that ERα primarily mediates the effects on bone development in males and females and regulates the suppressive effect of estrogens on bone resorption [64]. ERβ may counteract some of the effects of ERα on bone formation. Since estrogens are formed by aromatization of androgens, genetic perturbations of the aromatase gene have been of particular interest. Aromatase knockout female and male mice have low bone mass, indicating that the gene is functional in both sexes [65]. By contrast, transgenic mice with human ubiquitin C promoter-driven aromatase expression have increased bone mass [66]. Male mice with transgenic expression of the androgen receptor driven by the rat 3.6-kb Col1a1 promoter had an altered skeleton characterized by shorter femurs, thickened calvaria, and increased trabecular bone volume. The phenotype was sexually dimorphic since females were not affected [67]. The vitamin D receptor (VDR) has been both ablated and overexpressed in mice. VDR knockout mice have rickets [68] and they can be cured by normalization of mineral homeostasis [69]. Interestingly, the rickets in VDR knockout mice appear to be secondary to the apoptosis of hypertrophic chondrocytes [70]. Human osteocalcin promoter-targeted VDR transgenic mice had increased bone formation and decreased bone resorption [71]. Although glucocorticoid excess has deleterious effects on bone, the endogenous function of glucocorticoids in bone has been elusive. There is unequivocal evidence that glucocorticoid excess in humans and most animal species leads to glucocorticoid-induced osteoporosis, characterized by a decrease in bone formation due to decreased matrix protein production, decreased preosteoblast replication and increased osteoblast apoptosis; yet, many in vitro studies indicate that glucocorticoids facilitate osteoblast differentiation, particularly in human and rat osteoblast cultures [72]. To determine the role of endogenous glucocorticoids in bone, transgenic ligand inactivation models employing 11β-hydroxysteroid dehydrogenase 2 (11β-HSD2) have been developed [73, 74]. This enzyme metabolizes active glucocorticoids to inactive metabolites, thereby producing a barrier to glucocorticoid action in target cells [75]. This approach was undertaken because global knockout of the glucocorticoid receptor (GR) results in perinatal lethality, precluding an analysis of the bone phenotype in postnatal life [76]. Moreover, while a global or conditional GR knockout would not preclude glucocorticoids from signaling through the mineralocorticoid receptor (MR), a ligand inactivation strategy would block glucocorticoid signaling through both the GR and MR. Both the osteocalcin [74] and the 2.3-kb rat Col1a1 [73] promoters have been used to drive 11β-HSD2 in osteoblasts. Transgenic mice with Col2.3-driven 11β-HSD2 showed a sexually dimorphic bone phenotype consisting of decreased vertebral bone

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mass in females, with no change in femoral trabecular bone in males or females [73]. However, both sexes have decreased femoral cortical bone mass and decreased latestage osteoblast differentiation in primary calvarial and bone marrow stromal cell cultures (Kream, unpublished). Interestingly, osteoid volume was increased in female vertebral bone [73]. An osteocalcin-driven 11β-HSD2 transgene blocked glucocorticoid-induced osteoblast and osteocyte apoptosis [74]. These data indicate that glucocorticoids are required for bone formation and mineralization. Recent data indicate that 11β-HSD1 expression is high in undifferentiated human osteoblast cultures and decreases as the cells [77]. Thus, it can be inferred that local glucocorticoid production in the bone microenvironment plays a role osteoblast differentiation.

D. Transcription Factors Identification of Runx2 (originally Cbfa1) as a master regulator of osteoblast differentiation and osteogenesis relied heavily on genetic mouse models and shed light on the etiology of a heritable human skeletal disorder. Homozygous mice with targeted deletion of the Runx2 gene die shortly after birth due to respiratory failure and show a complete absence of skeletal mineralization due to inhibition of osteoblast differentiation [78]. Expression of Runx2 is restricted to skeletal elements, and is both necessary and sufficient for expression of a number of osteoblast lineage-specific or enriched genes, including bone sialoprotein, type I collagen, and osteocalcin [79]. Heterozygous-null mice, on the other hand, show a nonlethal skeletal phenotype that includes hypoplasia of the clavicles and delayed development of the intramembranous cranial bones [80]. Strikingly, this phenotype parallels that seen in human patients with cleidocranial dysplasia (CCD), who suffer from short stature, supernumerary teeth, hypoor aplastic clavicles, and patent fontanelles. This autosomal dominant condition was shown to arise by heterozygous loss of Runx2 resulting from deletions, insertions, or mis-sense mutations within the Runx2 gene locus [80]. Similarly, a radiation-induced mouse mutant (Ccd), with a phenotype indistinguishable from the Runx2 heterozygous-null mouse, was found to have a deletion affecting the Runx2 gene. To determine whether Runx2 plays a role not only in development but in the postnatal skeleton as well, Ducy et al. [81] have used a targeted transgenic loss-of-function strategy. In this model, a dominant-negative version of Runx2 containing only the DNA-binding domain was overexpressed in postnatal osteoblasts using a 1.3-kb osteocalcin promoter fragment. These mice showed a normal skeleton at birth but developed a progressive osteopenia

463 due to a reduced rate of bone formation that was associated with reduced expression of osteoblast extracellular matrix components. In some cases, transgenic models have yielded important insights in the field of bone biology without bone-specific targeting of the transgene. For example, two members of the AP-1 transcription factor family that lack trans-activation domains, Fra-1, and deltaFosB, cause dramatic effects on bone mass in mice when overexpressed under the control of ubiquitous promoters. Fra-1 expression, driven by the major histocompatibility complex class I antigen H2-Kb promoter, caused increases in cortical width as well as trabecular number and thickness beginning at 4 weeks of age [82]. This osteosclerotic phenotype results from an increase in bone formation that is caused, in turn, by enhanced osteoblast differentiation and an increase in the osteoblast and mineralizing surfaces. No effects on osteoblast proliferation were observed in this model. While Fra-1 lacks transcription activation domains and is generally thought to be a dominant negative regulator of Fos function, it should be noted that a “knock-in” of Fra-1 into the c-fos locus (effectively replacing c-fos) rescued of some abnormalities observed in c-fos-null mice, including the lack of osteoclasts. These observations suggest that transcription factors devoid of transactivation domains may not act solely as dominant negative inhibitors of their transcriptionally active counterparts. A somewhat similar osteosclerotic phenotype was seen in transgenic mice expressing a naturally occurring truncated splice variant of the Fos family known as deltaFosB. Expression of this transcription factor, directed by the nonspecific enolase promoter, caused an increase in bone formation and a progressive osteosclerosis with no effects on bone resorption [83]. Unlike the Fra-1 model, the increase in bone mass seen in these mice is accompanied by a reduction in adipogenesis both in vivo and in vitro, indicating a cell autonomous effect on fat. To determine whether deltaFosB could regulate bone formation in an adult mouse independently of any developmental effects, a model was developed in which deltaFosB expression was regulated using the inducible Tet-Off expression system. In these mice, postnatal induction of deltaFosB expression by withdrawal of tetracycline causes a progressive increase in bone mass that was reversed by the subsequent re-administration of tetracycline and silencing of deltaFosB expression, demonstrating the ability of deltaFosB to regulate bone mass in the adult animal [84]. Since fat mass was reduced in deltaFosB transgenic mice, it is possible that some effects of the transgene on bone mass were secondary to reduced adipogenesis and low serum leptin levels. However, long-term administration of leptin to these mice did not rescue the bone phenotype [85].

464 Alternatively, since osteoblasts and adipocytes share a common progenitor cell in bone marrow, it is possible that deltaFosB was promoting osteoblast differentiation at the expense of adipogenesis. This question was answered by targeting deltaFosB expression specifically to bone using an osteocalcin promoter fragment, which is expressed only in mature osteoblasts. These mice showed enhanced bone mass with no effect on adiposity, indicating that deltaFosB exerts independent cell autonomous effects on both the osteoblast and adipocyte lineages [86]. It has long been recognized that a reciprocal relationship exists between bone marrow adiposity and trabecular bone volume, leading to a search for transcription factors that might act as a molecular switch between these closely related lineages. One such gene candidate is PPARg, a member of the peroxisome proliferator activated receptor family that is known to play a critical role in adipocyte differentiation. Its expression is stimulated by C/EBP transcription factors, including C/EBPα, with which it syergistically activates expression of terminal adipocyte marker genes. Akune et al. [87] recently reported that embryonic stem cells from PPARγ-deficient mice failed to differentiate into adipocytes, but rather demonstrated spontaneous differentiation into osteoblasts. Mice heterozygous for PPARγ showed increased osteoblastogenesis and exhibited high bone mass. The cell-autonomous nature of this haploinsufficiency was demonstrated using bone marrow stromal cell cultures, which displayed an increase in osteoblast differentiation at the expense of adipogenesis. Cock et al. [88], in a study of a lipodystrophic PPARγ hyp/hyp mouse model, reported a marked increase in bone mass and consequent reduction in the size of the bone-marrow cavity, resulting in compensatory extramedullary hematopoiesis in the spleen. These gene disruption studies in mice raise the possibility that polymorphisms in the PPARg gene, already the subject of intense scrutiny for their possible contribution to obesity and the metabolic syndrome, might also play a role in the genetic specification of bone mass. As mentioned previously, C/EBP transcription factors regulate adipocyte differentiation, and recent evidence suggests that osteoblasts and adipocytes share a common pluripotent progenitor in bone marrow. However, little is known about the role of C/EBP transcription factors in the control of osteoblast differentiation or function. To assess the role of C/EBP transcription factors in osteogenesis, we targeted a naturally occurring dominant negative C/EBP isoform, called p20C/EBPβ or LIP, using the 3.6-kb Col1a1 promoter/first intron construct [89]. All four transgenic lines generated showed osteopenia, ranging from mild to severe, as evidenced by reduced trabecular bone volume. Severely affected lines also showed reduced cortical width. Dynamic histomorphometry demonstrated a decrease in

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the mineral apposition and bone formation rates, which was associated with reduced Col1a1 and osteocalcin mRNA levels. These data, using a dominant negative overexpression strategy to block all C/EBP function, suggest that C/EBP transcription factors may be required for osteoblast differentiation or maintenance of osteoblast function, and may be important determinants of bone mass. The cAMP signaling pathway is particularly important to bone biology since cAMP production is stimulated by the osteotropic hormones parathyroid hormone (PTH) and parathyroid hormone-related peptide (PTHrP). The classical cAMP signaling pathway includes receptor-mediated cAMP production by adenylate cyclase, cAMP binding, and activation of protein kinase A, translocation of PKA into the nucleus, phosphorylation of the cAMP response element binding protein (CREB), and regulation of gene transcription. Various approaches have been used to understand the role of the cAMP pathway in bone cells. A transgenic approach to understanding the role of the cAMP signaling pathway in endochondral bone formation has involved driving expression of a dominant negative CREB molecule (A-CREB) in chondrocytes by the type II collagen promoter [90]. The A-CREB transgene led to a shortlimbed dwarfism due to a delay in chondrocyte hypertrophy [90]. We have used a similar strategy to explore the role of cAMP signaling in osteoblasts. The inducible cAMP early repressor (ICER) is a naturally occurring dominant negative isoform of the cAMP responsive element modulator (Crem) gene [91]. ICER is strongly and transiently induced by cAMP and PTH in osteoblasts [92]. To gain further insight into the role of ICER in bone, and to determine the importance of the cAMP signaling pathway in osteoblasts, we have used the 3.6-kb Col1a1 promoter to drive ICER overexpression in osteoblasts. Mice in the most affected transgenic lines were small and had almost no trabecular bone (Kream, unpublished). There was impairment of bone formation and an increase in osteoclast formation and resorption. Thus, the bone phenotype of ICER transgenic mice suggests the cAMP pathway is critical for bone formation and remodeling. Genetic mouse models have also been instrumental in the recent demonstration that ATF4, a member of the CREB/ATF transcription factor family, is another key regulator of osteoblast function and bone mass. Using an ATF4-null mouse model, Yang et al. [93] reported a delay in bone formation during embryogenesis and persistence of a reduced bone mass phenotype through adulthood. This osteopenia was associated with reduced expression of osteocalcin and an inhibition of type I collagen synthesis at a post-transcriptional level. Interestingly, these authors demonstrated that ATF4 is a phosphorylation target for Rsk2, a growth factor-regulated protein kinase. An Rsk2

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gene ablation model showed a reduction of femur length, hypoplasia of calvaria bones with failure to close the fontanelles, and a reduced rate of bone formation. This phenotype appears to mirror the bone phenotype seen in human patients with Coffin–Lowry syndrome, an X-linked genetic disorder characterized by mental retardation and skeletal dysplasia. ATF4 phosphorylation may also play a role in the neural control of bone remodeling mediated by the sympathetic nervous system. Sympathetic stimulation of preosteoblastic cells via the β2-adrenergic receptor stimulates expression of receptor activator of NF-κB ligand, or RANKL, thereby promoting osteoclast differentiation and bone remodeling [94].

IV. PERSPECTIVES AND FUTURE DIRECTIONS Many new tools have recently become available for studying gene expression in transgenic models. Visual markers based on GFP, used singly or in combination, will allow assessment of promoter activity in real time in vivo [95]. It is possible to follow in real time the intracellular trafficking of chimeric proteins containing GFP domains. The knock-in of GFP reporters into endogenous genes will enable investigators to observe the spatial and temporal pattern of transgene expression in vivo. GFP can be used as a reporter to examine the induction or repression of gene transcription in vivo in distinct populations of cells. Finally, it will be possible to use GFP-marked cells to demonstrate the homing of cells to different tissues in vivo [95]. Newly developed technologies should result in transgenic models in which the spatial and temporal pattern of expression of the gene of interest better recapitulates the endogenous gene. Bacterial artificial chromosomes (BAC) and BAC recombineering methodology have been used to develop transgenes that are expressed in a developmental stage-specific and tissue-specific manner [96, 97]. The large chromosomal segments in BACs contain multiple genes with their full spectrum of regulatory elements. A gene of interest within a BAC is engineered with a reporter or gene of interest and used for transgenesis. Finally, the use of small interfering RNAs (siRNA) will allow investigators to develop models with hypomorphic gene expression. High levels of siRNA can be driven with viral vectors to accomplish sustained and heritable knockdown of gene expression in vivo [98–100]. Since it is relatively easy to make siRNA vectors and they can be obtained commercially, this technology may become an efficient means of modulating gene expression that is easier and faster than producing a complex targeting construct for each gene. Thus, these recent technological advances provide

remarkable potential for the future of functional genomic studies in the mouse.

References 1. Capecchi, M. R. (2005). Gene targeting in mice: functional analysis of the mammalian genome for the twenty-first century. Nat. Rev. Genet. 6, 507–512. 2. Davey, R. A., MacLean, H. E., McManus, J. F., Findlay, D. M., and Zajac, J. D. (2004). Genetically modified animal models as tools for studying bone and mineral metabolism. J. Bone Miner. Res. 19, 882–892. 3. McCauley, L. K. (2001). Transgenic mouse models of metabolic bone disease. Curr. Opin. Rheumatol. 13, 316–325. 4. Schilling, A. F., Priemel, M., Timo Beil, F., Haberland, M., Holzmann, T., Catala-Lehnen, P., Pogoda, P., Blicharski, D., Muldner, C., Locherbach, C., Rueger, J. M., and Amling, M. (2001). Transgenic and knock out mice in skeletal research. Towards a molecular understanding of the mammalian skeleton. J. Musculoskelet. Neuronal. Interact. 1, 275–289. 5. Clark, S., and Rowe, D. (2002). Application of transgenic mice to problems of skeletal biology. In “Principles of Bone Biology” (J. P. Bilezikian, L. G. Raisz, and G. A. Rodan, eds.), Vol. 2, pp. 1491–1502. Academic Press, San Diego, CA. 6. Jaenisch, R. (1988). Transgenic animals. Science 240, 1468–1474. 7. Lichtler, A., Stover, M. L., Angilly, J., Kream, B., and Rowe, D. W. (1989). Isolation and characterization of the rat alpha 1(I) collagen promoter. Regulation by 1, 25-dihydroxyvitamin D. J. Biol. Chem. 264, 3072–3077. 8. Rossert, J., Eberspaecher, H., and de Crombrugghe, B. (1995). Separate cis-acting DNA elements of the mouse pro-a1(I) collagen promoter direct expression of reporter genes to different type I collagenproducing cells in transgenic mice. J. Cell Biol. 129, 1421–1432. 9. Kalajzic, I., Kalajzic, Z., Kaliterna, M., Gronowicz, G., Clark, S. H., Lichtler, A. C., and Rowe, D. (2002). Use of type I collagen green fluorescent protein transgenes to identify subpopulations of cells at different stages of the osteoblast lineage. J. Bone Miner. Res. 17, 15–25. 10. Dacquin, R., Starbuck, M., Schinke, T., and Karsenty, G. (2002). Mouse alpha1(I)-collagen promoter is the best known promoter to drive efficient Cre recombinase expression in osteoblast. Dev. Dyn. 224, 245–251. 11. Bogdanovic, Z., Bedalov, A., Krebsbach, P. H., Pavlin, D., Woody, C. O., Clark, S. H., Thomas, H. F., Rowe, D. W., Kream, B. E., and Lichtler, A. C. (1994). Upstream regulatory elements necessary for expression of the rat COL1A1 promoter in transgenic mice. J. Bone Miner. Res. 9, 285–292. 12. Braut, A., Kalajzic, I., Kalajzic, Z., Rowe, D. W., Kollar, E. J., and Mina, M. (2002). Col1a1-GFP transgene expression in developing incisors. Connect Tissue Res. 43, 216–219. 13. Kesterson, R. A., Stanley, L., DeMayo, F., Finegold, M., and Pike, J. W. (1993). The human osteocalcin promoter directs bone-specific vitamin D-regulatable gene expression in transgenic mice. Mol. Endocrinol. 7, 462–467. 14. Frendo, J. L., Xiao, G., Fuchs, S., Franceschi, R. T., Karsenty, G., and Ducy, P. (1998). Functional hierarchy between two OSE2 elements in the control of osteocalcin gene expression in vivo. J. Biol. Chem. 273, 30509–30516. 15. Schinke, T., and Karsenty, G. (1999). Characterization of Osf1, an osteoblast-specific transcription factor binding to a critical cis-acting element in the mouse Osteocalcin promoters. J. Biol. Chem. 274, 30182–30189. 16. Clemens, T. L., Tang, H., Maeda, S., Kesterson, R. A., Demayo, F., Pike, J. W., and Gundberg, C. M. (1997). Analysis of osteocalcin

466

17.

18. 19.

20.

21. 22. 23. 24.

25.

26.

27.

28.

29.

30. 31.

32. 33. 34.

35.

36.

37.

expression in transgenic mice reveals a species difference in vitamin D regulation of mouse and human osteocalcin genes. J. Bone Miner. Res. 12, 1570–1576. Frenkel, B., Capparelli, C., Van Auken, M., Baran, D., Bryan, J., Stein, J. L., Stein, G. S., and Lian, J. B. (1997). Activity of the osteocalcin promoter in skeletal sites of transgenic mice and during osteoblast differentiation in bone marrow-derived stromal cell cultures: effects of age and sex. Endocrinology 138, 2109–2116. Nagy, A. (2000). Cre recombinase: the universal reagent for genome tailoring. Genesis 26, 99–109. Kalajzic, I., Braut, A., Guo, D., Jiang, X., Kronenberg, M. S., Mina, M., Harris, M. A., Harris, S. E., and Rowe, D. W. (2004). Dentin matrix protein 1 expression during osteoblastic differentiation, generation of an osteocyte GFP-transgene. Bone 35, 74–82. Chiu, W. S., McManus, J. F., Notini, A. J., Cassady, A. I., Zajac, J. D., and Davey, R. A. (2004). Transgenic mice that express Cre recombinase in osteoclasts. Genesis 39, 178–185. Ferron, M., and Vacher, J. (2005). Targeted expression of Cre recombinase in macrophages and osteoclasts in transgenic mice. Genesis 41, 138–145. Lewandoski, M. (2001). Conditional control of gene expression in the mouse. Nat. Rev. Genet. 2, 743–755. Soriano, P. (1999). Generalized lacZ expression with the ROSA26 Cre reporter strain [letter]. Nat. Genet. 21, 70–71. Novak, A., Guo, C., Yang, W., Nagy, A., and Lobe, C. G. (2000). Z/EG, a double reporter mouse line that expresses enhanced green fluorescent protein upon Cre-mediated excision. Genesis 28, 147–155. Lobe, C. G., Koop, K. E., Kreppner, W., Lomeli, H., Gertsenstein, M., and Nagy, A. (1999). Z/AP, a double reporter for cre-mediated recombination. Dev. Biol. 208, 281–292. Indra, A. K., Warot, X., Brocard, J., Bornert, J. M., Xiao, J. H., Chambon, P., and Metzger, D. (1999). Temporally-controlled sitespecific mutagenesis in the basal layer of the epidermis: comparison of the recombinase activity of the tamoxifen-inducible Cre-ER(T) and Cre-ER(T2) recombinases. Nucleic Acids Res. 27, 4324–4327. Feil, R., Brocard, J., Mascrez, B., LeMeur, M., Metzger, D., and Chambon, P. (1996). Ligand-activated site-specific recombination in mice. Proc. Natl Acad. Sci. USA 93, 10887–10890. Kim, J. E., Nakashima, K., and de Crombrugghe, B. (2004). Transgenic mice expressing a ligand-inducible cre recombinase in osteoblasts and odontoblasts: a new tool to examine physiology and disease of postnatal bone and tooth. Am. J. Pathol. 165, 1875–1882. Gossen, M., Freundlieb, S., Bender, G., Muller, G., Hillen, W., and Bujard, H. (1995). Transcriptional activation by tetracyclines in mammalian cells. Science 268, 1766–1769. Wan, M., and Cao, X. (2005). BMP signaling in skeletal development. Biochem. Biophys. Res. Commun. 328, 651–657. Johnson, M. L. (2004). The high bone mass family – the role of Wnt/Lrp5 signaling in the regulation of bone mass. J. Musculoskelet. Neuronal. Interact. 4, 135–138. Johnson, M. L., Harnish, K., Nusse, R., and Van Hul, W. (2004). LRP5 and Wnt signaling: a union made for bone. J. Bone Miner. Res. 19, 1749–1757. Westendorf, J. J., Kahler, R. A., and Schroeder, T. M. (2004). Wnt signaling in osteoblasts and bone diseases. Gene 341, 19–39. Rosen, C. J., and Donahue, L. R. (1995). Insulin-like growth factorspotential therapeutic options for osteoporosis. Trends Endocrinol. Metab. 6, 235–240. Van Wyk, J. J., and Smith, E. P. (1999). Insulin-like growth factors and skeletal growth: possibilities for therapeutic interventions. J. Clin. Endocrinol. Metab. 84, 4349–4354. Rosen, C. J. (2004). Insulin-like growth factor I and bone mineral density: experience from animal models and human observational studies. Best Pract. Res. Clin. Endocrinol. Metab. 18, 423–435. Baker, J., Liu, J. P., Robertson, E. J., and Efstratiadis, A. (1993). Role of insulin-like growth factors in embryonic and postnatal growth. Cell 75, 73–82.

BARBARA E. KREAM AND JOHN R. HARRISON 38. Powell-Braxton, L., Hollingshead, P., Warburton, C., Dowd, M., Pitts-Meek, S., Dalton, D., Gillett, N., and Stewart, T. A. (1993). IGF-I is required for normal embryonic growth in mice. Genes Dev. 7, 2609–2617. 39. Liu, J. L., Yakar, S., and LeRoith, D. (2000). Mice deficient in liver production of insulin-like growth factor I display sexual dimorphism in growth hormone-stimulated postnatal growth. Endocrinology 141, 4436–4441. 40. Bikle, D., Majumdar, S., Laib, A., Powell-Braxton, L., Rosen, C., Beamer, W., Nauman, E., Leary, C., and Halloran, B. (2001). The skeletal structure of insulin-like growth factor I-deficient mice. J. Bone Miner. Res. 16, 2320-2329. 40a.He, J., Rosen, C. J., Adams, D. J., and Kream, B. E. (2006). Postnatal growth and bone mass in mice with IGF-I haploinsufficiency. Bone, Jan 18 [Epub ahead of print]. 41. Mohan, S., and Baylink, D. J. (2005). Impaired skeletal growth in mice with haploinsufficiency of IGF-I: genetic evidence that differences in IGF-I expression could contribute to peak bone mineral density differences. J. Endocrinol. 185, 415–420. 42. Sjogren, K., Liu, J. L., Blad, K., Skrtic, S., Vidal, O., Wallenius, V., LeRoith, D., Tornell, J., Isaksson, O. G., Jansson, J. O., and Ohlsson, C. (1999). Liver-derived insulin-like growth factor I (IGF-I) is the principal source of IGF-I in blood but is not required for postnatal body growth in mice. Proc. Natl Acad. Sci. USA 96, 7088–7092. 43. Yakar, S., Liu, J. L., Stannard, B., Butler, A., Accili, D., Sauer, B., and LeRoith, D. (1999). Normal growth and development in the absence of hepatic insulin-like growth factor I. Proc. Natl Acad. Sci. USA 96, 7324–7329. 44. Ueki, I., Ooi, G. T., Tremblay, M. L., Hurst, K. R., Bach, L. A., and Boisclair, Y. R. (2000). Inactivation of the acid labile subunit gene in mice results in mild retardation of postnatal growth despite profound disruptions in the circulating insulin-like growth factor system. Proc. Natl Acad. Sci. USA 97, 6868–6873. 45. Zhang, M., Xuan, S., Bouxsein, M. L., von Stechow, D., Akeno, N., Faugere, M. C., Malluche, H., Zhao, G., Rosen, C. J., Efstratiadis, A., and Clemens, T. L. (2002). Osteoblast-specific knockout of the insulin-like growth factor (IGF) receptor gene reveals an essential role of IGF signaling in bone matrix mineralization. J. Biol. Chem. 277, 44005–44012. 46. Zhao, G., Monier-Faugere, M. C., Langub, M. C., Geng, Z., Nakayama, T., Pike, J. W., Chernausek, S. D., Rosen, C. J., Donahue, L. R., Malluche, H. H., Fagin, J. A., and Clemens, T. L. (2000). Targeted overexpression of insulin-like growth factor I to osteoblasts of transgenic mice: increased trabecular bone volume without increased osteoblast proliferation. Endocrinology 141, 2674–2682 47. Silha, J. V., Mishra, S., Rosen, C. J., Beamer, W. G., Turner, R. T., Powell, D. R., and Murphy, L. J. (2003). Perturbations in bone formation and resorption in insulin-like growth factor binding protein-3 transgenic mice. J. Bone Miner. Res. 18, 1834–1841. 48. Salih, D. A., Mohan, S., Kasukawa, Y., Tripathi, G., Lovett, F. A., Anderson, N. F., Carter, E. J., Wergedal, J. E., Baylink, D. J., and Pell, J. M. (2005). Insulin-like growth factor-binding protein-5 induces a gender-related decrease in bone mineral density in transgenic mice. Endocrinology 146, 931–940. 49. Devlin, R. D., Du, Z., Buccilli, V., Jorgetti, V., and Canalis, E. (2002). Transgenic mice overexpressing insulin-like growth factor binding protein-5 display transiently decreased osteoblastic function and osteopenia. Endocrinology 143, 3955–3962. 50. Atti, E., Boskey, A. L., and Canalis, E. (2005). Overexpression of IGFbinding protein 5 alters mineral and matrix properties in mouse femora: an infrared imaging study. Calcif. Tissue Int. 76, 187–193. 51. Simonet, W. S., Lacey, D. L., Dunstan, C. R., Kelley, M., Chang, M. S., Luthy, R., Nguyen, H. Q., Wooden, S., Bennett, L., Boone, T., Shimamoto, G., DeRose, M., Elliott, R., Colombero, A., Tan, H. L., Trail, G., Sullivan, J., Davy, E., Bucay, N., Renshaw-Gegg, L., Hughes, T. M., Hill, D., Pattison, W., Campbell, P., Sander, S., Van, G.,

Chapter 27 Transgenic Models of Bone Disease

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53.

54.

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56.

57.

58.

59.

60.

61.

62.

63.

64.

Tarpley, J., Derby, P., Lee, R., and Boyle, W. J. (1997). Osteoprotegerin: a novel secreted protein involved in the regulation of bone density. Cell 89, 309–319. Bucay, N., Sarosi, I., Dunstan, C. R., Morony, S., Tarpley, J., Capparelli, C., Scully, S., Tan, H. L., Xu, W., Lacey, D. L., Boyle, W. J., and Simonet, W. S. (1998). Osteoprotegerin-deficient mice develop early onset osteoporosis and arterial calcification. Genes Dev. 12, 1260–1268. Mizuno, A., Amizuka, N., Irie, K., Murakami, A., Fujise, N., Kanno, T., Sato, Y., Nakagawa, N., Yasuda, H., Mochizuki, S., Gomibuchi, T., Yano, K., Shima, N., Washida, N., Tsuda, E., Morinaga, T., Higashio, K., and Ozawa, H. (1998). Severe osteoporosis in mice lacking osteoclastogenesis inhibitory factor/osteoprotegerin. Biochem. Biophys. Res. Commun. 247, 610–615. Min, H., Morony, S., Sarosi, I., Dunstan, C. R., Capparelli, C., Scully, S., Van, G., Kaufman, S., Kostenuik, P. J., Lacey, D. L., Boyle, W. J., and Simonet, W. S. (2000). Osteoprotegerin reverses osteoporosis by inhibiting endosteal osteoclasts and prevents vascular calcification by blocking a process resembling osteoclastogenesis. J. Exp. Med. 192, 463–474. Kong, Y. Y., Yoshida, H., Sarosi, I., Tan, H. L., Timms, E., Capparelli, C., Morony, S., Oliveira-dos-Santos, A. J., Van, G., Itie, A., Khoo, W., Wakeham, A., Dunstan, C. R., Lacey, D. L., Mak, T. W., Boyle, W. J., and Penninger, J. M. (1999). OPGL is a key regulator of osteoclastogenesis, lymphocyte development and lymph-node organogenesis. Nature 397, 315–323. Dougall, W. C., Glaccum, M., Charrier, K., Rohrbach, K., Brasel, K., De Smedt, T., Daro, E., Smith, J., Tometsko, M. E., Maliszewski, C. R., Armstrong, A., Shen, V., Bain, S., Cosman, D., Anderson, D., Morrissey, P. J., Peschon, J. J., and Schuh, J. (1999). RANK is essential for osteoclast and lymph node development. Genes Dev. 13, 2412–2424. Franzoso, G., Carlson, L., Xing, L., Poljak, L., Shores, E. W., Brown, K. D., Leonardi, A., Tran, T., Boyce, B. F., and Siebenlist, U. (1997). Requirement for NF-kappaB in osteoclast and B-cell development. Genes Dev. 11, 3482–3496. Iotsova, V., Caamano, J., Loy, J., Yang, Y., Lewin, A., and Bravo, R. (1997). Osteopetrosis in mice lacking NF-kappaB1 and NF-kappaB2. Nat. Med. 3, 1285–1289. Xing, L., Bushnell, T. P., Carlson, L., Tai, Z., Tondravi, M., Siebenlist, U., Young, F., and Boyce, B. F. (2002). NF-kappaB p50 and p52 expression is not required for RANK-expressing osteoclast progenitor formation but is essential for RANK- and cytokine-mediated osteoclastogenesis. J. Bone Miner. Res. 17, 1200–1210. Lomaga, M. A., Yeh, W. C., Sarosi, I., Duncan, G. S., Furlonger, C., Ho, A., Morony, S., Capparelli, C., Van, G., Kaufman, S., van der Heiden, A., Itie, A., Wakeham, A., Khoo, W., Sasaki, T., Cao, Z., Penninger, J. M., Paige, C. J., Lacey, D. L., Dunstan, C. R., Boyle, W. J., Goeddel, D. V., and Mak, T. W. (1999). TRAF6 deficiency results in osteopetrosis and defective interleukin-1, CD40, and LPS signaling. Genes Dev. 13, 1015–1024. Ikeda, F., Nishimura, R., Matsubara, T., Tanaka, S., Inoue, J., Reddy, S. V., Hata, K., Yamashita, K., Hiraga, T., Watanabe, T., Kukita, T., Yoshioka, K., Rao, A., and Yoneda, T. (2004). Critical roles of c-Jun signaling in regulation of NFAT family and RANKL-regulated osteoclast differentiation. J. Clin. Invest. 114, 475–484. Matsuo, K., Galson, D. L., Zhao, C., Peng, L., Laplace, C., Wang, K. Z., Bachler, M. A., Amano, H., Aburatani, H., Ishikawa, H., and Wagner, E. F. (2004). Nuclear factor of activated T-cells (NFAT) rescues osteoclastogenesis in precursors lacking c-Fos. J. Biol. Chem. 279, 26475–26480. McCauley, L. K., Tozum, T. F., and Rosol, T. J. (2002). Estrogen receptors in skeletal metabolism: lessons from genetically modified models of receptor function. Crit. Rev. Eukaryot. Gene. Expr. 12, 89–100. Windahl, S. H., Andersson, G., and Gustafsson, J. A. (2002). Elucidation of estrogen receptor function in bone with the use of mouse models. Trends Endocrinol. Metab. 13, 195–200.

467 65. Oz, O. K., Hirasawa, G., Lawson, J., Nanu, L., Constantinescu, A., Antich, P. P., Mason, R. P., Tsyganov, E., Parkey, R. W., Zerwekh, J. E., and Simpson, E. R. (2001). Bone phenotype of the aromatase deficient mouse. J. Steroid Biochem. Mol. Biol. 79, 49–59. 66. Peng, Z., Li, X., Makela, S., Vaananen, H. K., and Poutanen, M. (2004). Skeletal changes in transgenic male mice expressing human cytochrome p450 aromatase. J. Bone Miner. Res. 19, 1320–1328. 67. Wiren, K. M., Zhang, X. W., Toombs, A. R., Kasparcova, V., Gentile, M. A., Harada, S., and Jepsen, K. J. (2004). Targeted overexpression of androgen receptor in osteoblasts: unexpected complex bone phenotype in growing animals. Endocrinology 145, 3507–3522. 68. Kato, S., Takeyama, K., Kitanaka, S., Murayama, A., Sekine, K., and Yoshizawa, T. (1999). In vivo function of VDR in gene expressionVDR knock-out mice. J. Steroid Biochem. Mol. Biol. 69, 247–251. 69. Amling, M., Priemel, M., Holzmann, T., Chapin, K., Rueger, J. M., Baron, R., and Demay, M. B. (1999). Rescue of the skeletal phenotype of vitamin D receptor-ablated mice in the setting of normal mineral ion homeostasis: formal histomorphometric and biomechanical analyses. Endocrinology 140, 4982–4987. 70. Donohue, M. M., and Demay, M. B. (2002). Rickets in VDR null mice is secondary to decreased apoptosis of hypertrophic chondrocytes. Endocrinology 143, 3691–3694. 71. Gardiner, E. M., Baldock, P. A., Thomas, G. P., Sims, N. A., Henderson, N. K., Hollis, B., White, C. P., Sunn, K. L., Morrison, N. A., Walsh, W. R., and Eisman, J. A. (2000). Increased formation and decreased resorption of bone in mice with elevated vitamin D receptor in mature cells of the osteoblastic lineage. Faseb. J. 14, 1908–1916. 72. Harrison, J. R., Woitge, H. W., and Kream, B. E. (2002). Genetic approaches to determine the role of glucocorticoid signaling in osteoblasts. Endocrine 17, 37–42. 73. Sher, L. B., Woitge, H. W., Adams, D. J., Gronowicz, G. A., Krozowski, Z., Harrison, J. R., and Kream, B. E. (2004). Transgenic expression of 11beta-hydroxysteroid dehydrogenase type 2 in osteoblasts reveals an anabolic role for endogenous glucocorticoids in bone. Endocrinology 145, 922–929. 74. O’Brien, C. A., Jia, D., Plotkin, L. I., Bellido, T., Powers, C. C., Stewart, S. A., Manolagas, S. C., and Weinstein, R. S. (2004). Glucocorticoids act directly on osteoblasts and osteocytes to induce their apoptosis and reduce bone formation and strength. Endocrinology 145, 1835–1841. 75. Krozowski, Z., Li, K. X., Koyama, K., Smith, R. E., Obeyesekere, V. R., Stein-Oakley, A., Sasano, H., Coulter, C., Cole, T., and Sheppard, K. E. (1999). The type I and type II 11beta-hydroxysteroid dehydrogenase enzymes. J. Steroid Biochem. Mol. Biol. 69, 391–401. 76. Kellendonk, C., Tronche, F., Reichardt, H. M., and Schutz, G. (1999). Mutagenesis of the glucocorticoid receptor in mice. J. Steroid Biochem. Mol. Biol. 69, 253–259. 77. Eijken, M., Hewison, M., Cooper, M. S., de Jong, F. H., Chiba, H., Stewart, P. M., Uitterlinden, A. G., Pols, H. A., and van Leeuwen, J. P. (2005). 11beta-Hydroxysteroid dehydrogenase expression and glucocorticoid synthesis are directed by a molecular switch during osteoblast differentiation. Mol. Endocrinol. 19, 621–631. 78. Komori, T., Yagi, H., Nomura, S., Yamaguchi, A., Sasaki, K., Deguchi, K., Shimizu, Y., Bronson, R. T., Gao, Y. H., Inada, M., Sato, M., Okamoto, R., Kitamura, Y., Yoshiki, S., and Kishimoto, T. (1997). Targeted disruption of Cbfa1 results in a complete lack of bone formation owing to maturational arrest of osteoblasts. Cell 89, 755–764. 79. Ducy, P., Zhang, R., Geoffroy, V., Ridall, A. L., and Karsenty, G. (1997). Osf2/Cbfa1: a transcriptional activator of osteoblast differentiation. Cell 89, 747–754. 80. Mundlos, S., Otto, F., Mundlos, C., Mulliken, J. B., Aylsworth, A. S., Albright, S., Lindhout, D., Cole, W. G., Henn, W., Knoll, J. H., Owen, M. J., Mertelsmann, R., Zabel, B. U., and Olsen, B. R. (1997). Mutations involving the transcription factor CBFA1 cause cleidocranial dysplasia. Cell 89, 773–779.

468 81. Ducy, P., Starbuck, M., Priemel, M., Shen, J., Pinero, G., Geoffroy, V., Amling, M., and Karsenty, G. (1999). A Cbfa1-dependent genetic pathway controls bone formation beyond embryonic development. Genes Dev. 13, 1025–1036. 82. Jochum, W., David, J. P., Elliott, C., Wutz, A., Plenk, H., Jr., Matsuo, K., and Wagner, E. F. (2000). Increased bone formation and osteosclerosis in mice overexpressing the transcription factor Fra-1. Nat. Med. 6, 980–984. 83. Sabatakos, G., Sims, N. A., Chen, J., Aoki, K., Kelz, M. B., Amling, M., Bouali, Y., Mukhopadhyay, K., Ford, K., Nestler, E. J., and Baron, R. (2000). Overexpression of DeltaFosB transcription factor(s) increases bone formation and inhibits adipogenesis. Nat. Med. 6, 985–990. 84. Sims, N. A., Sabatakos, G., Chen, J. S., Kelz, M. B., Nestler, E. J., and Baron, R. (2002). Regulating DeltaFosB expression in adult Tet-OffDeltaFosB transgenic mice alters bone formation and bone mass. Bone 30, 32–39. 85. Kveiborg, M., Chiusaroli, R., Sims, N. A., Wu, M., Sabatakos, G., Horne, W. C., and Baron, R. (2002). The increased bone mass in deltaFosB transgenic mice is independent of circulating leptin levels. Endocrinology 143, 4304–4309. 86. Kveiborg, M., Sabatakos, G., Chiusaroli, R., Wu, M., Philbrick, W. M., Horne, W. C., and Baron, R. (2004). DeltaFosB induces osteosclerosis and decreases adipogenesis by two independent cell-autonomous mechanisms. Mol. Cell Biol. 24, 2820–2830. 87. Akune, T., Ohba, S., Kamekura, S., Yamaguchi, M., Chung, U. I., Kubota, N., Terauchi, Y., Harada, Y., Azuma, Y., Nakamura, K., Kadowaki, T., and Kawaguchi, H. (2004). PPARgamma insufficiency enhances osteogenesis through osteoblast formation from bone marrow progenitors. J. Clin. Invest. 113, 846–855. 88. Cock, T. A., Back, J., Elefteriou, F., Karsenty, G., Kastner, P., Chan, S., and Auwerx, J. (2004). Enhanced bone formation in lipodystrophic PPARgamma(hyp/hyp) mice relocates haematopoiesis to the spleen. EMBO Rep. 5, 1007–1012. 89. Harrison, J. R., Huang, Y. F., Wilson, K. A., Kelly, P. L., Adams, D. J., Gronowicz, G. A., and Clark, S. H. (2005). Col1a1 promoter-targeted expression of p20 CCAAT enhancer-binding protein beta (C/EBPbeta), a truncated C/EBPbeta isoform, causes osteopenia in transgenic mice. J. Biol. Chem. 280, 8117–8124.

BARBARA E. KREAM AND JOHN R. HARRISON 90. Long, F., Schipani, E., Asahara, H., Kronenberg, H., and Montminy, M. (2001). The CREB family of activators is required for endochondral bone development. Development 128, 541–550. 91. Molina, C. A., Foulkes, N. S., Lalli, E., and Sassone-Corsi, P. (1993). Inducibility and negative autoregulation of CREM: an alternative promoter directs the expression of ICER, an early response repressor. Cell 75, 875–886. 92. Tetradis, S., Nervina, J. M., Nemoto, K., and Kream, B. E. (1998). Parathyroid hormone induces expression of the inducible cAMP early repressor in osteoblastic MC3T3-E1 cells and mouse calvariae [In Process Citation]. J. Bone Miner. Res. 13, 1846–1851. 93. Yang, X., Matsuda, K., Bialek, P., Jacquot, S., Masuoka, H. C., Schinke, T., Li, L., Brancorsini, S., Sassone-Corsi, P., Townes, T. M., Hanauer, A., and Karsenty, G. (2004). ATF4 is a substrate of RSK2 and an essential regulator of osteoblast biology; implication for Coffin-Lowry Syndrome. Cell 117, 387–398. 94. Elefteriou, F., Ahn, J. D., Takeda, S., Starbuck, M., Yang, X., Liu, X., Kondo, H., Richards, W. G., Bannon, T. W., Noda, M., Clement, K., Vaisse, C., and Karsenty, G. (2005). Leptin regulation of bone resorption by the sympathetic nervous system and CART. Nature 434, 514–520. 95. Hadjantonakis, A. K., Dickinson, M. E., Fraser, S. E., and Papaioannou, V. E. (2003). Technicolour transgenics: imaging tools for functional genomics in the mouse. Nat. Rev. Genet. 4, 613–625. 96. Sparwasser, T., Gong, S., Li, J. Y., and Eberl, G. (2004). General method for the modification of different BAC types and the rapid generation of BAC transgenic mice. Genesis 38, 39–50. 97. Copeland, N. G., Jenkins, N. A., and Court, D. L. (2001). Recombineering: a powerful new tool for mouse functional genomics. Nat. Rev. Genet. 2, 769–779. 98. Carpenter, A. E., and Sabatini, D. M. (2004). Systematic genomewide screens of gene function. Nat. Rev. Genet. 5, 11–22. 99. Voinnet, O. (2005). Induction and suppression of RNA silencing: insights from viral infections. Nat. Rev. Genet. 6, 206–220. 100. Shashikant, C. S., and Ruddle, F. H. (2003). Impact of transgenic technologies on functional genomics. Curr. Issues Mol. Biol. 5, 75–98.

Part III

Markers of Bone and Cartilage Metabolism

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Chapter 28

The Role of Genetic Variation in Osteoporosis André G. Uitterlinden, Joyce B.J. van Meurs, Fernando Rivadeneira, Johannes P.T.M.van Leeuwen, Huibert A.P. Pols

I. II. III. IV. V.

Department of Internal Medicine, Department of Epidemiology & Biostatistics, Erasmus Medical Centre, Rotterdam, The Netherlands

Abstract Osteoporosis has Genetic Influences Genome-wide Approaches to Find the Genes Association Analysis of Candidate Gene Polymorphisms Haplotypes

VI. Meta-analyses VII. Osteoporosis Candidate Genes: Collagen Type Iα1 and the Vitamin D Receptor VIII. Summary References

I. ABSTRACT

association analysis has much better possibilities, as is illustrated by the successful identification of risk alleles for several complex diseases. Candidate gene association analysis followed by replication and prospective multicentered meta-analysis, is currently the best way forward to identify genetic markers for complex traits, such as osteoporosis. To accomplish this, we need large (global) collaborative studies using standardized methodology and definitions, to quantify by meta-analysis the subtle effects of the responsible gene variants and assess heterogeneity in these associations between populations.

Over the past decades epidemiological research of so-called “complex” diseases, i.e., common age-related disorders such as cancer, cardiovascular disease, diabetes, and osteoporosis, has identified anthropometric, behavioral, and serum parameters as risk factors. Recently, genetic polymorphisms have gained considerable interest, propelled by the Human Genome Project and its sequela that have identified most genes and uncovered a plethora of polymorphic variants, some of which embody the genetic risk factors. In all fields of complex disease genetics (including osteoporosis) progress in identifying these genetic factors has been hampered by often controversial results. Because of the small effect size for each individual risk polymorphism, this is mostly due to low statistical power and limitations of analytical methods. Genome-wide scanning approaches can be used to find the responsible genes. It is by now clear that linkage analysis is not suitable for this, but genome-wide Dynamics of Bone and Cartilage Metabolism

II. OSTEOPOROSIS HAS GENETIC INFLUENCES Certain aspects of osteoporosis have been found to have strong genetic influences. This can be derived, for example, from genetic epidemiological analyses which showed that, 471

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472 in women, a maternal family history of fracture is positively related to fracture risk [1, 2]. Most evidence, however, has come from twin studies on bone mineral density (BMD) [3–5]. For BMD the heritability has been estimated to be high: 50–80% [3–5]. Thus, although twin studies can overestimate the heritability, a considerable part of the variance in BMD values might be explained by genetic factors while the remaining part could be due to environmental factors and to gene–environment interactions. This also implicates that there are “bone density” genes, variants of which will result in BMD levels that are different between individuals. These genetic differences can become apparent in different ways, for example, as peak BMD or as differences in the rates of bone loss at advanced age. While this notion has resulted in much attention being paid to the genetics of BMD in the field of osteoporosis, it is likely that this attention is also due simply to the widespread availability of devices to measure BMD. At the same time it is important to realize that (low) BMD is but one of many risk factors for osteoporotic fracture, the clinically most relevant endpoint of the disease. In addition, several other phenotypes related to osteoporosis have been shown to have heritable components including biochemical markers, heel ultrasound measurements, and skeletal geometry. Heritability estimates of fracture risk have been – understandably – much more limited, due to the scarcity of good studies allowing somewhat precise estimates. Collecting large collections of related subjects with accurate standardized fracture data is notoriously difficult in view of the advanced age at which they occur. While documenting a fracture event is now possible in several longitudinal studies, excluding a fracture event in those who report no fracture is more difficult because they could still suffer a fracture later in life. One option to overcome this might be to take controls which are (much) older. In the case of hip fracture patients (with a mean age of 80 years) this would require control subjects of 90–100 years. It is questionable whether such healthy survivors are proper controls for fracture cases. In general, studies on fracture have shown low heritability in the range of 20–50% [6–9]. Initial studies on Colles’ fracture of the wrist in 2471 white midwestern USA proband women aged 65 years, 3803 sisters of the probands, and their mothers, reported a heritability of 25% [6]. A similar estimate for osteoporotic fractures was derived from data on 2308 Finnish monozygotic twins and 5241 dizygotic twins [7, 8]. Andrew et al. [9] recently studied 6570 white healthy UK female volunteer twins between 18 and 80 years of age, and identified and validated 220 nontraumatic wrist fracture cases. They estimated a heritability of 54% for the genetic contribution to liability of wrist fracture in

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these women. Interestingly, while BMD was also highly heritable, the statistical models showed very little overlap of shared genes between the two traits in this study. An additional difficulty in these studies is, of course, the multifactorial and age-related etiology of fracture. This could implicate that perhaps it is not so much the heritability of bone strength which underlies the fracture heritability, but rather, for example, heritability of falls which by themselves also appear to have a heritable component [10]. Finally, it can be assumed that expression of genetic predispositions will not be constant during life, further complicating a precise estimate of heritability of (a certain type of) fracture. While it might be difficult to demonstrate fracture risk is heritable, one can also argue that it follows from simple logic reasoning that aspects of osteoporosis, including fracture risk, must have a genetic influence. We know that DNA is the blueprint of life, and that the genotype differs between individuals, and that phenotypes differ between individuals. Thus, the difficulties in demonstrating heritability of fracture risk are probably due to limitations of our methods and approaches of measuring it. The heritability estimates of osteoporosis indicate a considerable influence of environmental factors which can modify the effect of genetic predisposition. Gene– environment interactions in this respect include diet, exercise, and exposure to sunlight (for vitamin D metabolism), for example. While genetic predisposition itself (in the form of DNA sequence variations) will be constant during life, environmental factors tend to change during the different periods of life, resulting in different “expression levels” of the genetic susceptibility. Aging is associated with a general functional decline resulting in, for example, less exercise, less time spent outdoors, changes in diet, etc. This can result in particular genetic susceptibilities being revealed only later on in life after a period when they went unnoticed due to sufficient exposure to one or more environmental factors. Taking all this into account it becomes evident that osteoporosis is, not very surprisingly, considered a truly “complex” genetic trait. This complex character is shared with other common and often age-related traits with genetic influences such as diabetes, schizophrenia, Alzheimer’s disease, osteoarthritis, cancer, etc. “Complex” means that a trait is multifactorial as well as multigenic. Thus, genetic risk factors (i.e., certain alleles or gene variants) will be transmitted from one generation to the next, but the expression of these genotype factors in the final phenotype will be dependent on interaction with other gene variants and with environmental factors. Given that the Human Genome Project has now resulted in the identification of nearly all genes in the

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Chapter 28 Genetic Variation in Osteoporosis Table I. Allele

Haplotype kbp Linkage

Linkage disequilibrium Locus Mbp Microsatellite

Minisatellite

Mutation Polymorphism QTL RFLP SNP VNTR

Brief Glossary of Genetic Terms One of several alternative forms of a DNA sequence at a specific chromosomal location (locus). At each autosomal chromosomal locus in a cell two alleles are present, one inherited from the mother, the other from the father A series of alleles found at linked loci on a single chromosome (phase) Kilobasepairs (1 × 103 bp) The tendency of DNA sequences to be inherited together as a consequence of their close proximity on a chromosome Nonrandom association of alleles at linked loci A unique chromosomal location defining the position of a particular DNA sequence Megabasepairs (1 × 106 bp) A locus consisting of tandemly repetitive sequence units the size of which is (arbitrarily) defined as 1–5 bp A locus consisting of tandemly repetitive sequence units the size of which is (arbitrarily) defined as 6 bp or more An alteration in the DNA sequence The existence of two or more alleles at a frequency of at least 1% in the population Quantitative trait locus; a gene that influences quantitative variation in a trait Restriction fragment length polymorphism Single Nucleotide Polymorphism Variable number of tandem repeats; a polymorphic micro- or minisatellite

human genome, it is not very surprising that most attention in the analysis of gene–environment interactions has gone to the genes, also referred to as the “genocentric” approach. The idea behind this is that, once we know which gene variants are involved, it will be more straightforward to analyze the contribution of environmental factors and their interplay with genetic factors. Table I lists a few terms frequently used when discussing genetics of complex diseases.

III. GENOME-WIDE APPROACHES TO FIND THE GENES To identify and disentangle the genetic factors underlying risk for osteoporosis, basically two approaches can be applied: the “genome-wide analysis” approach, which is free of any hypothesis regarding which gene(s) are involved, and the “candidate gene” approach, which requires an a priori hypothesis as to which gene is involved (see Fig. 1). The idea was and is that genome-wide approaches are preferred because they will identify true and major genetic effects (the “low hanging fruit”) while candidate gene approaches are prone to heavy bias and thus not to be pursued with great priority. While genome-wide approaches have of course great appeal the results obtained so far have been (very) disappointing. This is mainly due to methodological limitations of linkage analysis as indicated below. However, novel and much better possibilities are now offered by the so-called genome-wide association strategies, which are also discussed below.

Figure 1 Methods of dissection of a complex disease/trait to identify the “risk” alleles of candidate genes that explain the genetic contribution to the trait. Resolution refers to the size of the genomic area that can be identified by the approach.

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In the meantime, several of the more classical candidate genes have already been analyzed to establish what the role of polymorphisms in these genes is in conferring osteoporosis risk. Even in the absence of such genome-wide scans, this is a valid approach to determine their contribution to the genetic risk for osteoporosis. Indeed, candidate gene analyses have identified genetic risk factors for osteoporosis, albeit of modest effect size. In addition, the outcome of any genome-wide analysis is the subsequent study of a particular candidate gene, so this approach will be among us for the coming years in any case. A paradigm in human genetics is already for a long time to scan the whole genome, free of any hypothesis and to look for co-segregation of the disease gene(s) with a DNA marker, so-called linkage. With the advent of the Human Genome Project a multitude of single nucleotide polymorphism (SNP) markers has now become available which, coupled with the discovery of so-called “haplotype blocks”, in the genome, has led to a new genome-wide approach based on the old-fashioned association analysis in cases and controls: the genome-wide association analysis. Thus, two genome-wide approaches can now be distinguished: those based on linkage analysis and those based on association analysis.

A. Genome-wide Linkage Analysis

of relatives (siblings, pedigrees, etc.) are genotyped for hundreds of DNA markers (mostly microsatellites but now also thousands of SNPs can be used, e.g., the Affymetrix 10K SNP chip) evenly spread over the genome. Most genome searches focus on humans, although several mice genome searches have also been performed. Such genome searches are based on the assumption that relatives who share a certain phenotype will also share one or more chromosomal areas, identical-by-descent, containing one or more gene variants causing (to a certain extent) the phenotype of interest (e.g., low BMD). The gene is then said to be linked with the DNA marker used to “flag” a certain chromosomal region, but this area is usually several million base pairs in size. Upon positive linkage, subsequent research will then have to analyze dozens of genes in the chromosomal area to determine which one(s) is (are) the one(s) involved in bone metabolism, and then identify the particular sequence variant in that (those) gene(s) giving rise to (aspects of) osteoporosis. Up to the stage of finding linkage to one or more chromosomal regions this approach has been quite successful in the past and linkage results from several genome searches have indeed been published (e.g., reference [14]). However, so far the subsequent step to identify the gene variant causing the linkage peak has been difficult and has not resulted in many – if any – “osteoporosis risk genes”. Why is that?

Finding the responsible gene for monogenic disorders (caused by rare mutations in a single gene) is a straightforward routine exercise for specialized laboratories. This is based on linkage analysis in pedigrees (in which the disease is segregating according to Mendelian laws) whereby a standardized set of hundreds of well-characterized DNA markers are analyzed for co-segregation with a phenotypic endpoint. An example of such an approach is the identification of LRP5 gene mutations being responsible for osteoporosis pseudoglioma as well as for a trait called High Bone Mass [11, 12]. Many other examples in the bone field exist and so for single-gene diseases this approach works very well to identify “bone genes”. However, the complex (and nonMendelian) character of osteoporosis makes it quite resistant to the methods of analysis which in the past decades have worked so well for the monogenic diseases. Therefore different and often more cumbersome approaches have to be applied which can be broadly defined as top–down and bottom–up approaches (see, e.g. reference [13]). In top–down approaches whole-genome searches are performed which indicate which chromosomal areas might contain osteoporosis genes. For linkage analysis hundreds

1. Weak statistical linkage evidence. It has proven difficult to find statistically significant linkage with LOD scores above 3.6 for genome-wide significance. Typically only “suggestive” linkage is found with LOD scores of 1–3. This indicates that there are not a few major genes for osteoporosis but rather many subtle genes. 2. Lack of replication of linkage peaks. There is hardly any single region which has been identified convincingly (and statistically significant) by more than one genome search (with perhaps some possible exceptions such as 1p36). Replication has also proven difficult because the families/pedigrees/sibling pairs between different genome searches have differences in ethnicity, environmental factors, gender, age, etc. 3. Lack of power to find a gene. The effects per polymorphism are too weak to be detected with the typical number of sibling pairs available. It has therefore been difficult to go beyond “linkage” and to demonstrate that a certain gene variant is causing the linkage peak observed in the genome search. Chromosomal regions showing linkage are typically 1–10 million base pairs wide containing dozens of candidate genes. In these candidate genes hundreds of polymorphisms occur, some of which are organized in linkage disequilibrium blocks of 10–20 kb,

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making it virtually impossible (by statistical genetics alone) to pinpoint the causative variant using the linkage design. 4. Choice of endpoint. Most genome searches have focused on BMD as an endpoint. BMD, however, explains only a part of osteoporotic fracture risk. It is difficult to “switch” between major outcomes during or after the study because the families/sib-pairs are selected on the basis of such an endpoint. It is also noteworthy that all linkage scans have identified few linkage peaks, suggesting few genes to explain the genetics of osteoporosis. It is by now widely assumed however, that many, maybe hundreds of gene variants are implicated. Given the very limited population-attributable risk of the claimed genes identified so far by this approach, this probably also reflects the very limited power of genome-wide linkage scans. Recently, an example of the identification of an allegedly “major osteoporosis gene” through a genome search was published. It was the identification of BMP2 (20p12.3) as a risk factor for osteoporotic fracture by analysis of Icelandic pedigrees and a Danish cohort by the company Decode from Iceland [15]. Although one might interpret this as proof of the success of the genome search approach, several notions would preclude the following. 1. BMP2 was already well-known as an important gene for bone metabolism for several decades and as such represents a good candidate gene. So far, however, nobody had looked for polymorphisms in this gene in relation to osteoporosis. 2. The effect size of the BMP2 gene variants on fracture risk in the two samples (Icelandic and Danish) is modest and in line with what has been found for other candidate genes. It is rather premature to call this a “major” risk gene for osteoporosis and, especially given the low population frequency of a risk allele (37Ser, f<10%), the evidence presented would not support this view. 3. Only one “major” linkage peak was observed. Does this imply this is the only major osteoporosis gene? Clearly not. Perhaps then only so in Iceland and/or in Denmark? 4. The study identified a low-frequency amino acid variant (Ser37Ala) in the gene as being responsible for the effect but did not provide functional evidence. In addition, haplotypes were constructed that associated with osteoporosis (defined in many different ways). The very large haplotypes (up to 200 kb!), however, are ill-defined and encompass dozens of so-far unknown polymorphisms and this haplotype association could not be convincingly replicated in another, Danish, sample. While BMP2 was heralded as the first osteoporosis gene to be identified by way of a genome linkage search,

more research is needed, including multiple association studies in different populations and meta-analysis, to establish what the contribution of variants in this gene are to osteoporosis risk. So, the final proof of its involvement must come from candidate gene association studies and it is far from certain this will be a major important risk gene.

B. Genome-wide Association Analysis Since we now know that the human genome has a haplotype block structure, this has opened a novel approach to search the genome for genetic markers of disease: the genome-wide association (GWA) analysis (reviewed in references [16, 17]). In this approach many hundreds of thousand of SNPs are analyzed in sets of (usually) a few hundred unrelated cases and unrelated controls. The exact set of SNPs depends somewhat on the techniques used: Affymetrix (www.affymetrix.com) currently offers chips with SNPs that are detected using XbaI/HindIII digestion to reduce genomic complexity and that are more or less evenly spread across the genome. Densities have increased over time from 10 000 (10 k), to 100 K and now 500 K. Perlegen (www.perlegen.com) offers as a service their in-house developed Affymetrix chip technology containing >1 million haplotype tagging SNPs [18]. Illumina (www.illumina.com) is using glass arrays which are spotted at high density with very selected SNPs, such as coding SNPs (100 k), and will shortly offer such arrays with haplotype tagging SNPs. The first successful use of a genome-wide association analysis using such high numbers of SNPs was reported by Ozaki et al. [19] who, by means of a large-scale case (n = 94)- control (n = 658) association study using 92 788 gene-based SNPs, identified significant associations between myocardial infarction and two SNPs in LTA (encoding lymphotoxin-alpha): one SNP changed an aminoacid residue from threonine to asparagine (Thr26Asn) while another SNP in intron 1 influenced the transcription level of LTA. More recently, Klein et al. [20] reported a GWA study of 96 cases and 50 controls for polymorphisms associated with age-related macular degeneration (AMD), a major cause of blindness in the elderly. Among 116 204 singlenucleotide polymorphisms genotyped (using Affymetrix chips), a tyrosine–histidine change at amino acid position 402 (T402H) in the complement factor H gene (CFH) was strongly associated with AMD. This polymorphism is in a region of CFH that binds heparin and C-reactive protein. The CFH gene is located on chromosome 1 in a region repeatedly linked to AMD in family-based studies.

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The relatively low density of SNPs used in these studies, in combination with the limited genetic complexity of diseases studied (of AMD in particular), could explain why only a few associated regions were observed in these studies, while one would expect (many) more gene regions to show up. Indeed ,very recent press releases from Affymetrix (www.microarraybulletin.com; April 2005) indicated similar successes for multiple sclerosis (MS; Cohen et al.), graft vs. host disease (GVHD; Ogawa et al.), and cardiovascular disease (CVD; Salonen et al.), where more cases/ controls were analyzed (1800 for MS, 2000 for GVHD, and with higher density SNP arrays (100–500 K) and more gene regions were found to be associated with the disease (80 for multiple sclerosis, 400 for CVD). However, we will have to await the detailed report in the scientific literature of these studies to know the exact methods used and results found. Nevertheless, these results from GWA studies show the great potential they have for elucidating complex disease and it will only be a matter of time before similar approaches will be reported for osteoporosis. The only current limitation might be that they are quite expensive: they cost roughly 1000 euros per DNA sample. Yet, the same statistical requirements apply as in a given case-control or population-based study of a candidate gene polymorphism. This means that for truly complex traits a minimum of several hundred cases/controls, if not

Figure 2

thousands of subjects have to be studied, thus requiring 1–10 million euros per GWA study. For less complex traits, e.g. those with a few major genes such as might be the case for AMD, this might be less. It should be noted that the size of the haplotype blocks/ chromosomal regions identified through the GWA approach is much smaller (10–50 kb) than what is usually found in genome-wide linkage analyses (which is 1–10 million(!) base pairs). This offers major advantages for the subsequent research. Yet, even when one or more such haplotype blocks are found associated, these blocks need further scrutiny to identify the one or more polymorphisms driving the association and functionality has to be established. So this approach could (also) end up with a candidate gene analysis. After that, the associations have to be replicated in other populations and, finally, meta-analysis can be used to assess heterogeneity and quantify effect size (see later).

IV. ASSOCIATION ANALYSIS OF CANDIDATE GENE POLYMORPHISMS The bottom–up approach to identify genetic risk factors for osteoporosis builds upon biology, i.e., the known involvement of a particular gene in aspects of osteoporosis, e.g., bone metabolism (see Fig. 2). This gene is then referred

A schematic flow-diagram depicts the different steps in a candidate gene polymorphism analysis. On top genome-wide association analysis is indicated that will identify multiple areas across the genome as LD blocks within candidate genes. This is used in concordance with biological evidence based on three independent sources, to implicate a gene in the disease of interest.

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to as a “candidate gene”. The candidacy of such a gene can be established by several lines of evidence. 1. Cell biological and molecular biological experiments, indicating for example bone cell-specific expression of the gene. 2. Animal models in which a gene has been mutated (e.g., natural mouse mutants), overexpressed (transgenic mice), or deleted (knockout mice) and which result in a bone phenotype. 3. Naturally occurring mutations of the human gene resulting in monogenic Mendelian diseases with a bone phenotype. Subsequently, in the candidate gene frequently occurring sequence variants (polymorphisms) have to be identified which supposedly lead to subtle differences in function of the encoded protein. We distinguish mutations from polymorphisms purely on the basis of frequency: polymorphisms occur in at least 1% of the population, mutations in less. Sequence analysis of a “candidate” osteoporosis gene in a number of different individuals will identify sequence variants, but also several databases are now available which contain this information (e.g., NCBI, Celera, HapMap, and several more specialized databases). Some DNA sequence variations will be just polymorphic (anonymous polymorphisms) while others will have consequences for the activity of the protein encoded (functional polymorphisms). These can include, e.g., sequence variations leading to alterations in the amino acid composition of the protein, changes in the 5′ promoter region leading to differences in mRNA expression, and/or polymorphisms in the 3′ region leading to differences in mRNA degradation. Clearly, it depends on the gene how many and what kind of polymorphisms will be present in the population. Some genes will have been, for example, under more evolutionary pressure and will not display much variation. Other genes, however, might be part of a pathway with sufficient redundancy to allow for more genetic variation to occur. Polymorphisms of interest are usually first tested in population-based and/or case-control “association studies”, to evaluate their contribution to the phenotype of interest at the population level. However, association studies do not establish cause and effect; they just show correlation or co-occurrence of one with the other. Table II lists some of the pitfalls in interpreting candidate gene association studies. Yet, it is also important to realize in this respect that it is of uncertain value to test functionality of a certain polymorphism in the absence of an association at the population level. Cause and effect has to be established in truly functional cellular and molecular biological experiments involving,

Table II.

Pitfalls in Genetic Association Studies of Candidate Genes

Epidemiological/statistical 1. Sample size is too small, leading to chance findings 2. Population is biased due to selection, admixture, inbreeding, etc. 3. Environmental factors differ between populations 4. Lack of standardized phenotypes Genetic 1. Allelic heterogeneity: different alleles are associated in different populations 2. Locus heterogeneity: gene effects differ between populations due to genetic drift and founder effect 3. Linkage disequilibrium: one or more adjacent genes are the true susceptibility loci instead of the locus being tested Molecular genetic 1. Lack of standardized genotyping techniques 2. Ill-defined choice of polymorphisms: there is no known functional effect of the polymorphism to provide a direct biological explanation of the association Other 1. Publication bias

e.g. transfection of cell lines with allelic constructs and testing activities of the different alleles. This can occur at different levels of organization (see Fig. 3) and depends on the type of protein analyzed, e.g., enzymes vs. matrix molecules vs. transcription factors. Acknowledging these complexities will remain a challenge, once an association has been observed, to identify the correct test of functionality; and vice versa once functionality has been established, to identify the correct endpoint in an epidemiological study. Because functional polymorphisms lead to meaningful biological differences in function of the encoded “osteoporosis” protein this also makes the interpretation of association analyses using these variants quite straightforward. For example, for functional polymorphisms it is expected that the same allele will be associated with the same phenotype in different populations. This can even be extended to similar associations being present in different ethnic groups, although allele frequencies can of course differ by ethnicity [21]. Out of the three lines of evidence mentioned above, numerous candidate genes have emerged and Table III lists only a few of these. These include “classical” candidate genes such as collagen type I, the vitamin D receptor, and the estrogen receptors, but also more novel genes of which their involvement in bone biology has only recently become known. A good example of the latter is the identification of LRP5 gene mutations being responsible for osteoporosis pseudoglioma as well as for a trait called high bone mass [11, 12]. These studies have identified LRP5 as a candidate gene, but, of course, not established its role as a genetic risk factor for osteoporosis.

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Figure 3 Depiction of how “functional” DNA polymorphisms might affect physiological processes at different levels of organization, that ultimately result in an association that is seen after many years (for age-related disorders this can be 80 years) of “exposure” to the risk factor.

With this plethora of candidate genes it is difficult to decide where to start. Initially this happened somewhat randomly but current choices are guided by increasing insights in the metabolic pathways in which the genes play a pivotal role. For example, the identification of LRP5 as a candidate gene has put the complete Wnt-signaling pathway on the map as a target. It can therefore now be expected that multiple genes from this pathway will be tested as candidate genes for osteoporosis. The current focus in genetic studies of osteoporosis is quite strongly on common variants which are expected to explain a substantial portion of population variance, simply due to their frequency in the population (10–50%). However, also more rare variants (0.1–10% or even less frequent) can contribute to population variance with stronger effects, and perhaps can play an important role in certain populations but not in others. An example of this was described by Cohen et al. [22] who tested whether rare DNA sequence variants collectively contribute to variation in plasma levels of high-density lipoprotein cholesterol (HDL-C). They sequenced three candidate genes (ABCA1, APOA1, and LCAT) that cause Mendelian forms of low HDL-C levels in individuals from a population-based study. Nonsynonymous sequence variants were significantly more common (16% versus 2%) in individuals with low HDL-C (below the fifth percentile) than in those with high HDL-C (above the 95th percentile). Similar findings were obtained in an independent population, and biochemical studies indicated that most sequence variants in the low HDL-C group were functionally important. Thus, rare alleles with major phenotypic

effects contribute significantly to low plasma HDL-C levels in the general population. Similarly, such rare alleles of bone genes might contribute to variation in BMD and other bone parameters, and even fracture risk in the general population. Thus, when we compare the genome-wide linkage approach to the candidate gene approach, the latter approach is now clearly the more promising one [16–18, 23]. Genome-wide linkage searches are not designed and not statistically powered to detect the many subtle gene effects which underlie osteoporosis following the “common variant–common disease” hypothesis. The genome-wide association analysis seems to be a better alternative, but has not been used so far and has not identified any risk genes yet. Thus, the current way forward is to simply test individual candidate genes to establish what their contribution is to osteoporosis risk. Once that has been established, the interaction or multiplicative effects of several genes will be analyzed and, finally, gene–environment interactions can be studied.

V. HAPLOTYPES From resequencing studies for the HapMap project (www.hapmap.org) it has become evident that, on average, one out of every 300 base pairs is varying in the population. Given an average size of 100 kb of a gene this means there are hundreds of polymorphisms in a given gene. Thus, candidate gene analyses will have to focus on which of the many variant nucleotides are the ones that actually matter.

Table III. Selected Osteoporosis Candidate Genes Gene name Matrix protein molecules Osteocalcin α2HS Glycoprotein Osteopontin Osteonectin Collagen type Iα1 Collagen type Iα2 Matrix-associated enzymes Cathepsin K Alkaline phosphatase Carbonic anhydrase II Matrix metalloproteinase 3 Lysyl oxidase Lysine hydroxylase Lysine hydroxylase 2 Lysine hydroxylase 3 Calciotropic (steroid) hormone/receptors/enzymes Estrogen receptor α Estrogen receptor β Aromatase Androgen receptor Glucocorticoid receptor Vitamin D receptor Vitamin D binding protein B3-adrenergic receptor Peroxisome proliferator-activated receptor-gamma Calcium sensing receptor Calcitonin receptor Parathyroid hormone PTH receptor Epidermal growth factor Gonadotropin releasing hormone 1 Gonadotropin releasing hormone receptor Luteinizing hormone beta peptide LH-choriogonadotropin receptor Growth factors/cytokines/receptors Interleukin-1β receptor antagonist Interleukin-4 Interleukin-6 Transforming growth factor β1 Transforming growth factor β2 Growth hormone Growth hormone receptor Insulin-like growth factor I Insulin-like growth factor I receptor Insulin-like growth factor 2 Insulin-like growth factor II receptor IGF-binding protein 3 Tumor necrosis factor α TNF receptor superfamily/1β Bone morphogenetic protein 2 Bone morphogenetic protein 3 Sclerostin Osteoprotegerin RANK RANKL-ligand/OPG-ligand Wnt-signaling pathway Low density lipoprotein receptor-related protein 5* Homocystein pathway Methylene TetraHydroFolate reductase Cystathionine beta-synthase Methionine synthase reductase Methyltetrahydrofolate-homocysteine s-methyltransferase Thymidylate synthetase Miscellaneous Major histocompatibility complex Apolipoprotein E

Gene symbol

Chromosomal location

BGLAP AHSG SPP1 SPOCK COL1A1 COL1A2

1q25–q31 3q27 4q21–q25 5q31.3–q32 17q21.3–q22.1 7q22.1

CTSK ALPL CA2 MMP3 LOX PLOD PLOD2 PLOD3

1q21 1p36.1–p34 8q22 11q22.3 5q23.3–q31.2 1p36 3q23–q24 7q36

ESR1 ESR2 CYP19 AR GR/NR3C1 VDR DBP/GC ADRB3 PPARG CASR CALCR PTH PTHR1 EGF GNRH1 GNRHR LHB LHCGR

6q25.1 14q23 15q21.1 Xq11 5q31 12q13 19q13.3 8p12–p11.2 3p25 3q21–q24 7q21.3 11p15.3–p15.1 3p22–p21.1 4q25 8p21–p11.2 4q21.2 19q13.32 2p21

IL-1RN IL-4 IL-6 TGFB1 TGFB2 GH1 GHR IGF1 IGF1R IGF2 IGF2R IGFBP3 TNF TNFRG5 BMP2 BMP3 SOST OPG/TNFRSF11B RANK/TNFRSF11A RANKL/TNFSF11

2q14.2 5q31.3 7p21 19q13.2 1q41 17q22–q24 5p13–p12 12q22–q23 15q25–q26 11p15.5 6q26 7p14–p12 6p21.3 1p36.3–p36.2 20p12.3 4p14–q21 17q12–q21 8q24 18q22.1 13q14

LRP5

11q12

MTHFR CBS MTRR MTR TYMS

1p36.3 21q22.3 5p15.3–p15.2 1q43 18p11.32

MHC/HLA APOE

6p21.3 19q13.2

480 That is, which sequence variation is functionally relevant by changing expression levels, changing codons, etc. Given the average size of a gene and the relatively young age of human populations, it can be predicted that several sequence variations “that matter” will co-exist in a gene in a given number of subjects from a study population. A major challenge of fundamental research will therefore be to unravel the functionality of these variations and how they interact with each other within the gene. More recently, it has become clear that some of these neighboring polymorphisms are not independent from each other in genetic terms, that is to say some of them tend to “travel together” in so-called haplotypes. Haplotypes are strings of coupled or linked variants which occur, on average, over a distance of 10–30 kb in the human genome. With polymorphisms occurring in roughly one out of 300 base pairs this means there will be dozens of polymorphisms within these “haplotype blocks”. An important aspect of association analyses in this respect is then to establish which common haplotype alleles are occurring in the candidate gene, which has two important practical consequences. 1. If association is found of a particular allele of an individual polymorphism with a certain phenotype/disease, this can also be explained by an adjacent polymorphism within the haplotype block. Thus, one can never be sure what causes the association until the haplotype structure at that position within the gene has been resolved. 2. When, for example, 20 polymorphisms are located within a haplotype block only a fraction (typically only 30%) has to be genotyped to identify the haplotype alleles. This saves on time and money to perform the association analyses while obtaining maximal information relevant for point 1. Once such haplotype blocks have been identified, it becomes important how they are organized in the gene of interest. A typical gene can have one or several haplotype blocks covering the promoter region, another block covering the coding region and yet another block covering regulatory regions 3′ of the gene. For the functioning of a complete gene in a given cell of a given subject, it is then important to know which combination of haplotype alleles is present in that subject. In Figure 4 a hypothetical example is given, of the functional relevance of gene-wide combinations of genotypes (based on single SNPs or on haplotypes). The figure describes the situation when two subjects have identical genotypes for three adjacent polymorphic sites when analyzed independently. Yet, they differ in their combination of alleles on one chromosome, and this will result in different effects at the cellular level. This example illustrates that the effects of single polymorphisms might

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be difficult to interpret when ignoring the polymorphisms in the rest of the haplotype block and the other haplotype blocks in the gene. Alternatively, if an association is driven by a single SNP rather than by a haplotype, the association will be diluted if haplotyes are analyzed. In the case of high-density SNP analyses of a candidate gene both approaches, i.e., haplotypes and single SNP analysis, should therefore be compared to determine the driving force of the strongest association.

VI. META-ANALYSES In the coming years we can expect to see more and more association analyses to be performed of an ever-increasing list of candidate gene polymorphisms. It will therefore be necessary to put all these data in perspective by performing meta-analyses of the individual association analyses. Meta-analysis can quantify the results of various studies on the same topic and by assessing heterogeneity estimate and explain their diversity. Recent evidence indicates that a systematic meta-analysis approach can estimate populationwide effects of genetic risk factors for human disease [24] and that large studies are more conservative in these estimates and should preferably be used [25]. An analysis of 301 studies on genetic associations (on many different diseases) concluded that there are many common variants in the human genome with modest but real effects on common disease risk, and that studies using large samples will be able to convincingly identify such variants [26]. Several meta-analyses of osteoporosis candidate gene polymorphisms have been published in the past few years, in particular for VDR polymorphisms, the COL1A1 Sp1 polymorphism, and ESR1 polymorphisms. Although these have to some extent all confirmed association of these polymorphisms with bone-related endpoints, including fracture, considerable caution must be taken in interpreting these results. For example, these meta-analyses were all based on published data, thereby raising the possibility of publication bias influencing the outcome of the analysis, since some (perhaps mostly negative) studies were not included. Another important problem can be lack of standardization of methods to determine genotype and phenotype in each of the component populations. For example, there are differences in accuracy of genotyping technologies, BMD can be measured in different ways by different machines, and “osteoporotic fracture” is a quite heterogeneous phenotype. In addition, such meta-analysis treats studies to some extent as equal whereby subtle differences might not be properly accounted for, such as for example differences between populations in environmental factors. In addition, some alleles might be quite specific for some

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Figure 4

Hypothetical example of the importance of gene-wide genotype combinations. Three adjacent SNPs in different parts of a gene are shown for two individuals (A and B indicated at the bottom). The subjects A and B have identical genotypes, i.e., they are both heterozygous for all three SNPs. However, they have different allele combinations on the same chromosome (numbered 1–4): 1 + 2 for subject A and 3 + 4 for subject B. The promoter area regulates production of mRNA while the 3′ UTR is involved in degradation of mRNA and their interaction/combined effects regulate the net availability of the mRNA for translation into the protein. In this case the example is shown for a promoter polymorphism which has two alleles + and −, of which the + allele is the high-producer variant in certain target cells. Of the two different 3′ UTR variants + and −, the + is the more stable 3′ UTR resulting in more mRNA being maintained. Hence, a “good” promoter allele and a “good” 3′ UTR allele on the same chromosome, result in more protein being produced. The protein itself can occur in two variants: a less active “risk” form (−) and a more active form (+), and both A and B are again heterozygous for this polymorphism. The combined result of the particular allele combinations is that individual A has less of the “risk” protein than individual B in the target cell. This could not have been predicted by analyzing single SNPs and/or only looking at genotypes of individual SNPs, but is only evident upon analysis of the gene-wide genotype combinations.

populations but not for others (such as the lactose deficiency polymorphism LPH C-13170T) and show specific gene– environment interactions and gene–gene interactions. One might for these reasons therefore still prefer to perform a single, large, well-powered study on candidate gene associations and seek replication, rather than rely on meta-analysis results based on published data. To overcome the potential drawbacks of the classical meta-analysis approach in the field of osteoporosis, the EUsponsored GENOMOS (Genetic Markers for Osteoporosis) consortium attempts to perform prospective meta-analyses of osteoporosis candidate gene polymorphisms using standardized methods of genotyping and phenotyping. The GENOMOS project involves the large-scale study of several candidate gene polymorphisms in relation to osteoporosisrelated outcomes in subjects drawn from several European centers. Its main outcomes are femoral neck and lumbar spine BMD and fractures, and design details are described in the first meta-analysis of individual-level data on the

ESR1 gene performed by the GENOMOS team [27]. The GENOMOS meta-analysis of three polymorphisms in the ESR1 gene (intron 1 polymorphisms XbaI [dbSNP: rs9340799] and PvuII [dbSNP: rs2234693] and promoter TA repeats microsatellite) and haplotypes thereof, among 18 917 individuals in eight European centers, demonstrated no effects on BMD but a modest effect on fracture risk (19–35% risk reduction for XbaI homozygotes), independent of BMD [27]. An important aspect of this study is its prospective multicenter design. This means the genotype data are generated for all centers only after which the association analysis is done, thereby rendering it practically immune to possible publication bias. The targets of the study are polymorphisms for which some a priori evidence for involvement in osteoporosis is present already; it is not designed to be a risk gene-discovery tool and currently therefore cannot assess all genetic diversity across a gene. While fracture has been debated as an endpoint in genetics

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of osteoporosis studies, this was chosen in the GENOMOS study because it is clinically the most relevant endpoint. Statistical power of the GENOMOS study to detect genetic effects on fracture risk is high with >5000 fractures in the dataset. With such a diverse set of populations included in the GENOMOS study, possible population stratification could be a problem. This is not likely, however, because GENOMOS involves almost exclusively white Caucasians, who – in addition – come from relatively stable populations with little recent immigration/emigration. Indeed, so far the tested allele frequencies for ESR1 [27] and COL1A1 [27a] are remarkably similar between populations supporting the absence of major population stratification. Importantly, functional SNPs are expected to show similar effects across different ethnic groups in spite of the different genetic backgrounds of the ethnic groups. In this respect it has recently been demonstrated that genetic markers for proposed gene–disease associations can vary in frequency across populations, but their biological impact on the risk for common diseases may usually be consistent across traditional “racial” boundaries [21]. Thus, such a meta-analysis approach will identify individual genetic risk factors but it will probably also be instrumental in estimating presence and effect size of genetic interactions (gene–gene) and gene–environment interactions. This approach will be followed for genes in a certain pathway, for which we know that interaction is likely, and can be extended to explore unexpected interactions. However, even with large studies of, e.g., 20 000 subjects, this might be difficult to convincingly demonstrate. This stresses the need for even larger studies.

VII. OSTEOPOROSIS CANDIDATE GENES: COLLAGEN TYPE Ia1 AND THE VITAMIN D RECEPTOR The vitamin D receptor (VDR) is the candidate gene that initiated the molecular genetic studies on osteoporosis in 1994 by the work of Morrison, Eisman et al. [28]. They initially reported anonymous 3′ polymorphisms, detected as BsmI, ApaI, and TaqI RFLPs, to be associated with differences in BMD and claimed that up to 80% of the population variance in BMD could be explained by this single gene. Although some of their results were later retracted [29], it left behind the idea that there could be such a thing as a single osteoporosis gene. For a complex disease such as osteoporosis this is very unlikely and, indeed, by now several large population-based association analyses [30] and including several meta-analyses [31–33] have

indicated that – at most – the VDR variants explain only a few percent of the population variance of BMD, if any. Thus, for prediction of BMD the VDR does not seem to be a marker. Whether it is associated with fracture risk remains to be analyzed by meta-analysis of the scarce published data so far and by the prospective type of metaanalysis as has been discussed above. However, in view of its pleiotropic involvement in a number of biological pathways, the vitamin D receptor polymorphisms have also been analyzed in relation to a number of other diseases. For example, we have shown VDR polymorphisms to be associated with (bony) aspects of osteoarthritis, i.e., osteophytosis [34]. Yet, a major problem with the vitamin D receptor gene polymorphisms is still the lack of insight into the functional consequences of the polymorphisms used so far and which other polymorphisms there are in this gene. Therefore a major effort in our own laboratory is focused on finding additional polymorphic sites but now in functionally relevant areas of the gene, such as the 3′ untranslated region and the 5′ promotor area of this gene. One such functional VDR polymorphisms is the Cdx2 polymorphism in the 1a/1e promoter region of the VDR gene. The Cdx2 polymorphism has been well-characterized by the studies of Arai et al. [35] and Yamamoto et al. [36]. The G to A polymorphism is located in a Cdx2-binding site in the 1e promoter region and this site is suggested to play an important role in intestinal-specific transcription of the VDR gene. As this is the site where the calcium absorption predominantly takes place, the Cdx2 site is thought to influence the vitamin D regulation of calcium absorption. The A allele has been demonstrated to be more “active” than the G-allele by binding the Cdx2 transcription factor more strongly, and by having more transcriptional activity [35]. Thus, the A allele is thought to cause increased VDR expression in the intestine and, thereby, can increase the transcription of calcium transport proteins (such as calbindin 9K and -28K, TRPV5, TRPV6). This could thereby enhance the intestinal absorption of calcium and result in increased BMD. Although this increased BMD has indeed been demonstrated for Japanese women who carry the A-allele [35], this was not found in a much larger study of Dutch Caucasian women [37]. Yet, the A-allele of this polymorphism was indeed found to be associated with decreased fracture risk (as would be expected from having an increased BMD) in the large study of Dutch Caucasian women, but independently of BMD [37]. Therefore, although the functionality of this polymorphism has indeed been convincingly demonstrated, the exact mechanism whereby the A-allele would confer lower risk for fracture has not been elucidated yet and requires further study.

Chapter 28 Genetic Variation in Osteoporosis

Another example of a functional sequence variation in an osteoporosis candidate gene that has quickly shown promising associations, is the G to T substitution in the Sp1binding site in the first intron of the collagen type Iα1 gene. After its discovery and initial association analysis by Grant et al. [38], a large-scale population analysis in the Rotterdam Study showed the T allele to be associated with low BMD and increased fracture risk [39]. Interesting observations in this cohort of 1782 elderly women were the increase with age of the genotype-dependent differences and the fact that the genotype-dependent fracture risk was independent of the differences in BMD. Meanwhile, several other studies have confirmed these observations as is demonstrated in meta-analyses of published studies [40–42], making this a promising osteoporosis candidate gene polymorphism with consistent, albeit modest, effects. The molecular way in which this polymorphism is influencing BMD and fracture risk is becoming more and more clear. Work by Ralston and colleagues [40] has shown that the T-allele has increased affinity for the Sp1-binding factor, has increased mRNA production, increased protein production, and, in biomechanic experiments on bone biopsies, has been shown to be associated with decreased bone strength. It has been suggested that the over-representation of collagen type Iα1 homo-trimers might explain the decreased bone strength although this still has to be proven. The COLIA1 and VDR gene are “classical” osteoporosis candidate genes and indeed have been shown to be associated with real, albeit modest, effects. Therefore there must be other osteoporosis genes carrying risk alleles. An overview of possible candidate genes is given in Table III. These include more classical genes (such as the ESR1 gene) but also more unexpected genes (such as the IL genes) and more novel genes such as SOST, BMP2, and LRP5. Systematic analysis of these pathways, and the candidate genes in them, will provide an ever-increasing clearer picture of the individual components of the genetic risk factors. Association analysis in population-based studies is the most promising way forward in this respect.

VIII. SUMMARY So, in summary, if people were to embark on an association study of a candidate gene to identify genetic markers for osteoporosis, what would be the crucial issues to address? A few suggestions. 1. Take a large population. Bigger is better, to make your initial observations statistically robust. 2. Identify proper endpoints upfront. Fractures are clinically the most relevant but you need substantial

483 numbers to make your finding statistically robust. BMD is only one of the risk factors but it is a continuous trait and gives more statistical power. Population-based studies have the advantage of being able to switch phenotypes during analysis very easily, for case-controls this possibility is very limited. 3. Cover all relevant genetic variation within the gene. Focus on functionally relevant variants within a gene. A clear-cut functional variant can be analyzed in isolation, ignoring the rest of the genetic variation in the gene. However, determine the haplotype structure to understand how the complete gene is functioning. 4. P-values: rather seek replication of your finding. Simple adjustment for multiple testing is regarded as not appropriate (where to start and stop counting?). Rather, formulate a proper a priori hypothesis and seek replication(s) of the observed association in similar populations. 5. Perform a meta-analysis to quantify effect size and assess heterogeneity. Join consortia with your population and datasets to standardize genotype and phenotype definition and estimate effect size of polymorphisms and assess heterogeneity of the association in different populations, preferably by prospective meta-analysis rather than meta-analysis of published data. Although still in its infancy with respect to having clinical implications, the field of genetics of osteoporosis (or any complex disease for that matter) is expected to eventually find applications in three main areas. 1. Prediction of response-to-treatment (pharmacogenetics). Polymorphisms in, e.g., drug-metabolizing enzymes, will result in different efficiencies with which drugs can exert their effect. The same holds true for receptors of hormones and growth factors, analogs of which are currently prescribed as treatment. Genotype analysis can identify those subjects expected to profit most from a particular treatment or exclude those subjects which will suffer more from side-effects (personalized medicine). 2. Identification of subjects-at-risk. Subjects carrying risk alleles are more likely to develop osteoporosis. Genotype analysis will allow to take preventive measures, targeted at the individual at an early stage. 3. Identification of novel disease pathways and therapeutic targets. Such novel pathways can be targeted for drug development and investigated further for key components. So far only one polymorphism is currently being considered as an osteoporosis risk factor (the COL1A1 Sp1 polymorphism) and commercial parties have taken up interest in this genetic marker. Its utility in clinical practice has to be considered with considerable caution, however. For example, analyses in different ethnic populations have

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shown it to be present mostly in Caucasian subjects [43]. Furthermore, interaction of this variant with another polymorphism (the VDR 3′ variants) has been demonstrated [44]. This latter study also highlights the complex and multigenic nature of osteoporosis. It underlines the need to identify additional osteoporosis risk alleles to better understand how particular genetic markers are expressed and result in a phenotype. Another spin-off of genetic research of osteoporosis is the discovery of new and/or unexpected genes and pathways to be involved in determining, e.g., BMD. A good example is the recent discovery of LRP5 mutations to influence BMD [5, 6] and by this the identification of the Wnt-signaling pathway to be involved in bone metabolism. Such discoveries led to new possibilities to develop drugs to treat osteoporosis. In addition, such genes become candidate osteoporosis risk genes and will be searched for polymorphisms. Risk alleles resulting from such analyses can then be added to the still-growing list of osteoporosis gene variants. Thus, in spite of complicating factors, genetic research will contribute to a further understanding of complex diseases, including osteoporosis. The identification of new genes, or new roles of already-known genes, will allow insights in mechanistic pathways which might help in designing therapeutic protocols. Finally, the description of genetic variation underlying phenotypic variation can be used, in concert with existing easy-to-assess risk factors, in prediction of risk for aspects of osteoporosis. In this respect, novel therapeutic protocols but also insights in gene–environment interactions allow for ways to further improve treatment of patients

7.

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9.

10.

11.

12.

13. 14.

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References 1. Cummings, S. R., Nevitt, M. C., Browner, W. S., Stone, K., Fox, K. M., Ensrud, K. E., Cauley, J., Black, D., and Vogt, T. M. (1995). Risk factors for hip fracture in white women. New Engl. J. Med. 332, 767–773. 2. Kanis, J. A., Johansson, H., Oden, A., Johnell, O., de Laet, C., Eisman, J. A., McClsokey, E. V., Mellstrom, D., Melton III, L. J., Pols, H. A. P., Reeve, J., Silman, A. J., and Tenenhouse, A. (2004). A family history of fracture and fracture risk: a meta-analysis. Bone 35, 1029–1037. 3. Smith, D. M., Nance, W. E., Kang, K. W., Christian, J. C., and Johnston, C. C. (1973). Genetic factors in determining bone mass. J. Clin. Invest. 52, 2800–2808. 4. Pocock, N. A., Eisman, J. A., Hopper, J. L., Yeates, G. M., Sambrook, P. N., and Ebert, S. (1987). Genetic determinants of bone mass in adults: a twin study. J. Clin. Invest. 80, 706–710. 5. Flicker, L., Hopper, J. L., Rodgers, L., Kaymakci, B., Green, R. M., and Wark, J. D. (1995). Bone density determinants in elderly women: a twin study. J. Bone Miner. Res. 10, 1607–1613. 6. Deng, H. W., Chen, W. M., Recker, S., Stegman, M. R., Li, J. L., Davies, K. M., Zhou, Y., Deng, H., Heaney, R., and Recker, R. R. (2000). Genetic determination of Colles’ fracture and differential bone mass

18.

19.

20.

21.

22.

23.

in women with and without Colles’ fracture. J. Bone Miner. Res. 15, 1243–1252. Kannus, P., Palvanen, M., Kaprio, J., Parkkari, J., and Koskenvuo, M. (1999). Genetic factors and osteoporotic fractures in elderly people: prospective 25 year follow up of a nationwide cohort of elderly Finnish twins. BMJ 319, 1334–1337. MacGregor, A., Snieder, H., and Spector, T. D. (2000). Genetic factors and osteoporotic fractures in elderly people. Twin data support genetic contribution to risk of fracture. BMJ 320, 1669–1670. Andrew, T., Antioniades, L., Scurrah, K. J., Macgregor, A. J., and Spector, T. D. (2005). Risk of wrist fracture in women is heritable and is influenced by genes that are largely independent of those influencing BMD. J. Bone Miner. Res. 20, 67–74. Carmelli, D., Kelly-Hayes, M., Wolf, P. A., Swan, G. E., Jack, L. M., Reed, T., and Guralnik, J. M. (2000). The contribution of genetic influences to measures of lower-extremity function in older male twins. J. Gerontol. A Biol. Sci. Med. Sci. 55, B49–B53. Gong, Y., Slee, R. B., Fukai, N., Rawadi, G., Roman-Roman, S., Reginato, A. M., and 56 others. (2001). LDL Receptor-related protein 5 (LRP5) affects bone accrual and eye development. Cell 107, 513–523. Little, R. D., Carulli, J. P., Del Mastro, R. G., Dupuis, J., Osborne, M., Folz, C., and 29 others. (2002). A mutation in the LDL Receptor related protein 5 gene results in the autosomal dominant high bone mass trait. Am. J. Hum. Genet. 70, 11–19. Lander, E. S., and Schork, N. J. (1994). Genetic dissection of complex traits. Science 265, 2037–2048. Ralston, S. H., Galwey, N., MacKay, I., Albagha, O. M., Cardon, L., Compston, J. E., Cooper, C., Duncan, E., Keen, R., Langdahl, B., McLellan, A., O’Riordan, J., Pols, H. A., Reid, D. M., Uitterlinden, A. G., Wass, J., and Bennett, S. T. (2005). Loci for regulation of bone mineral density in men and women identified by genome wide linkage scan: the FAMOS study. Hum. Mol. Genet. 14, 943–951. Styrkarsdottir, U., Cazier, J. B., Kong, A., Rolfsson, O., Larsen, H., Bjarnadottir, E., Johannsdottir, V. D., Sigurdardottir, M. S., Bagger, Y., Christiansen, C., Reynisdottir, I., Grant, S. F., Jonasson, K., Frigge, M. L., Gulcher, J. R., Sigurdsson, G., and Stefansson, K. (2003). Linkage of Osteoporosis to Chromosome 20p12 and Association to BMP2. PLoS Biol. 1(3), E69. Hirschhorn, J. N., and Daly, M. J. (2005). Genome-wide association studies for common diseases and complex traits. Nat. Rev. Genet. 6, 95–108. Wang, W. Y., Barratt, B. J., Clayton, D. G., and Todd, J. A. (2005). Genome-wide association studies: theoretical and practical concerns. Nat. Rev. Genet. 6, 109–118. Hinds, D. A., Stuve, L. L., Nilsen, G. B., Halperin, E., Eskin, E., Ballinger, D. G., Frazer, K. A., and Cox, D. R. (2005). Whole-genome patterns of common DNA variation in three human populations. Science 307, 1072–1079. Ozaki, K., Ohnishi, Y., Iida, A., Sekine, A., Yamada, R., Tsunoda, T., Sato, H., Sato, H., Hori, M., Nakamura, Y., and Tanaka, T. (2002). Functional SNPs in the lymphotoxin-alpha gene that are associated with susceptibility to myocardial infarction. Nat. Genet. 32, 650–654. Klein, R. J., Zeiss, C., Chew, E. Y., Tsai, J. Y., Sackler, R. S., Haynes, C., Henning, A. K., Sangiovanni, J. P., Mane, S. M., Mayne, S. T., Bracken, M. B., Ferris, F. L., Ott, J., Barnstable, C., and Hoh, J. (2005). Complement factor H polymorphism in age-related macular degeneration. Science 308, 385–389. Ioannidis, J. P., Ntzani, E. E., and Trikalinos, T. A. (2004). ‘Racial’ differences in genetic effects for complex diseases. Nat. Genet. 36, 1312–1318. Cohen, J. C., Kiss, R. S., Pertsemlidis, A., Marcel, Y. L., McPherson, R., and Hobbs, H. H. (2004). Multiple rare alleles contribute to low plasma levels of HDL cholesterol. Science 305, 869–872. Risch, N., and Merikangas, K. (1999). The future of genetic studies of complex human diseases. Science 273, 1516–1517.

Chapter 28 Genetic Variation in Osteoporosis 24. Ioannidis, D. G., Ntzani, E. E., Trikalinos, T. A., and ContopoulosIoannidis, D. G. (2001) Replication validity of genetic association studies. Nat. Genet. 29, 306–309. 25. Ioannidis, J. P., Trikalinos, T. A., Ntzani, E. E., and ContopoulosIoannidis, D. G. (2003). Genetic associations in large versus small studies: an empirical assessment. Lancet 361, 567–571. 26. Lohmueller, K. E., Pearce, C. L., Pike, M., Lander, E. S., and Hirschhorn, J. N. (2003) Meta-analysis of genetic association studies supports a contribution of common variants to susceptibility to common disease. Nat. Genet. 33, 177–182. 27. Ioannidis, J. P., Ralston, S. H., Bennett, S. T., Brandi, M. L., Grinberg, D., Karassa, F. B., Langdahl, B., van Meurs, J. B., Mosekilde, L., Scollen, S., Albagha, O. M., Bustamante, M., Carey, A. H., Dunning, A. M., Enjuanes, A., van Leeuwen, J. P., Mavilia, C., Masi, L., McGuigan, F. E., Nogues, X., Pols, H. A., Reid, D. M., Schuit, S. C., Sherlock, R. E., and Uitterlinden, A. G. (2004) GENOMOS Study. Differential genetic effects of ESR1 gene polymorphisms on osteoporosis outcomes. JAMA 292, 2105–2114. 27a.Ralston, S. H., Uitterlinden, A. G., Brandi, M. L., Balcells, S., Langdahl, B. L., Lips, P., Lorenc, R., Obermayer-Pietsch, B., Scollen, S., Bustamante, M., Husted, L. B., Carey, A. H., Diez-Perez, A., Dunning, A. M., Falchetti, A., Karczmarewicz, E., Kruk, M., van Leeuwen, J. P. T. M., van Meurs, J. B. J., Mangion, J., McGuigan, F. E. A., Mellibovsky, L., del Monte, F., Pols, H. A. P., Reeve, J., Reid, D. M., Renner, W., Rivadeneira, F., van Schoor, N., Sherlock, R. E., Ioannidis, J. P. A., APOSS investigators, EPOS investigators, EPOLOS investigators, FAMOS investigators, LASA investigators, and Rotterdam Study investigators; for the GENOMOS Study. Large-scale evidence for the effect of the COL1A1 Sp1 polymorphism on osteoporosis outcomes: The GENOMOS Study. PloS Medicine 2006 (in press). 28. Morrison, N. A., Qi, J. C., Tokita, A., Kelly, P. J., Crofts, L., Nguyen, T. V., Sambrook, P. N., and Eisman, J. A. (1994). Prediction of bone density from vitamin D receptor alleles. Nature 367, 284–287. 29. Morrison, N. A., Qi, J. C., Tokita, A., Kelly, P. J., Crofts, L., Nguyen, T. V., Sambrook, P. N., and Eisman, J. A. (1997). Prediction of bone density from vitamin D receptor alleles (correction). Nature 387, 106. 30. Uitterlinden, A. G., Pols, H. A. P., Burger, H., Huang, Q., van Daele, P. L. A., van Duijn, C. M., Hofman, A., Birkenhäger, J. C., and van Leeuwen, J. P .T .M. (1996). A large scale population based study of the association of vitamin D receptor gene polymorphisms with bone mineral density. J. Bone Miner. Res. 11, 1242–1248. 31. Cooper, G. S., and Umbach, D. M. (1996). Are vitamin D receptor polymorphisms associated with bone mineral density? A meta-analysis. J. Bone Miner. Res. 11, 1841–1849. 32. Gong, G., Stern, H. S., Cheng, S. C., Fong, N., Mordeson, J., Deng, H. W., and Recker, R. R. (1999). The association of bone mineral density with vitamin D receptor gene polymorphisms. Osteoporos. Int. 9, 55–64.

485 33. Thakkinstian, A., D’Este, C., Eisman, J., Nguyen, T., and Attia, J. (2004). Meta-analysis of molecular association studies: vitamin D receptor gene polymorphisms and BMD as a case study. J. Bone Miner. Res. 19, 419–428. 34. Uitterlinden, A. G., Burger, H., Huang, Q., Odding, E., van Duijn, C. M., Hofman, A., Birkenhäger, J. C., van Leeuwen, J. P. T. M., and Pols, H. A. P. (1997). Vitamin D receptor genotype is associated with osteoarthritis. J. Clin. Invest. 100, 259–263. 35. Arai, H., Miyamoto, K. I., Yoshida, M., Yamamoto, H., Taketani, Y., Morita, K., Kubota, M., Yoshida, S., Ikeda, M., Watabe, F., Kanemasa, Y., and Takeda, E. (2001). The polymorphism in the caudal-related homeodomain protein Cdx-2 binding element in the human vitamin D receptor gene. J. Bone Miner. Res. 16, 1256–1264. 36. Yamamoto, H., Miyamoto, K-I., Bailing, L., Taketani, Y., Kitano, M., Inoue, Y., Morita, K., Pike, J. W., and Takeda, E. (1999). The caudalrelated homeodomain protein Cdx-2 regulates vitamin D receptor gene expression in the small intestine. J. Bone Miner. Res. 14, 240–247. 37. Fang, Y., van Meurs, J. B., Bergink, A. P., Hofman, A., van Duijn, C. M., van Leeuwen, J. P., Pols, H. A., and Uitterlinden, A. G. (2003). Cdx-2 polymorphism in the promoter region of the human vitamin D receptor gene determines susceptibility to fracture in the elderly. J. Bone Miner. Res. 18, 1632–1641. 38. Grant, S. F. A., Reid, D. M., Blake, G., Herd, R., Fogelman, I., and Ralston, S. H. (1996). Reduced bone density and osteoporotic vertebral fracture associated with a polymorphic Sp1 binding site in the collagen type Iα1 gene. Nature Genet. 14, 203–205. 39. Uitterlinden, A. G., Burger, H., Huang, Q., Yue, F., McGuigan, F. E. A., Grant, S. F. A., Hofman, A., van Leeuwen, J. P. T. M., Pols, H. A. P., and Ralston, S. H. (1998). Relation of alleles at the collagen type Iα1 gene to bone density and risk of osteoporotic fractures in postmenopausal women. New Engl. J. Med. 338, 1016–1021. 40. Mann, V., Hobson, E. E., Li, B., Stewart, T. L., Grant, S. F. A., Robins, S. P., Aspden, R. M., and Ralston, S. H. (2001). A COLIA1 Sp1 binding site polymorphism predisposes to osteoporotic fracture by affecting bone density and quality. J. Clin. Invest. 107, 899–907. 41. Efstathiadou, Z., Tsatsoulis, A., and Ioannidis, J. P. A. (2001). Association of collagen Iα1 Sp1 polymorphism with the risk of prevalent fractures: a meta-analysis. J. Bone Miner. Res. 16, 1586–1592. 42. Mann, V., and Ralston, S. H. (2003). Meta-analysis of COL1A1 Sp1 polymorphism in relation to bone mineral density and osteoporotic fracture. Bone 32, 711–717. 43. Beavan, S., Prentice, A., Bakary, D., Yan, L., Cooper, C., and Ralston, S. H. (1998). Polymorphism of the collagen type Iα1 gene and ethnic differences in hip fractures rates. N. Engl. J. Med. 339, 351–352. 44. Uitterlinden, A. G., Weel, A. E. A. M., Burger, H., Fang, Y., van Duijn, C. M., Hofman, A., van Leeuwen, J. P. T. M., and Pols, H. A. P. (2001). Interaction between the vitamin D receptor gene and collagen type Iα1 gene in susceptibility for fracture. J. Bone Miner. Res. 16, 379–385.

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Chapter 29

Measurement of Calcium, Phosphate and Magnesium Heinrich Schmidt-Gayk

Department of Endocrinology, Laboratory Group, Im Breitspiel 15, 69126 Heidelberg, Germany

I. Measurement of Calcium II. Measurement of Phosphate

III. Measurement of Magnesium References

I. MEASUREMENT OF CALCIUM

the ionized calcium concentration is increased. For each 0.1 decrease in pH, the ionized calcium rises by about 0.05 mmol/L. Fetuin-A prevents extraosseous precipitation of calcium–phosphate complexes. a. Choice of Analyte: Total Calcium or Ionized Calcium For routine purposes in not severely ill patients it is often sufficient to determine the concentration of total calcium together with the concentration of total protein and/or albumin. The concentration of total protein or albumin might be necessary for the calculation of corrected serum calcium. Therefore, the most frequently used methods for the determination of serum or plasma calcium measure total calcium, and less often ionized calcium is measured using an ion-selective electrode (see below). However, the corrections for total protein, albumin, and pH are in many cases poor substitutes for measuring ionized calcium. Therefore, as described below, in severely ill patients, in transfused patients and in patients with an unclear diagnosis, the ionized calcium should be used whenever possible to guide therapy. b. Reference Range The reference range of calcium in serum or plasma is given in Table I.

A. Calcium in Serum 1. INTRODUCTION

Calcium in serum (or plasma) is found in three fractions, namely the ionized fraction (Ca2+), the complexed fraction, and the protein-bound fraction. The sum of the three fractions is the total serum (or plasma) calcium, which is fairly constant in a healthy person. In a normal person, approximately 45–50% is ionized, 10–15% is complexed to anions such as bicarbonate, citrate, sulfate, phosphate, and lactate and 40% is protein-bound (about 32% to albumin, about 8% to globulins) [1]. Some calcium is bound as a calcium–phosphate complex to α2-Heremann-Schmid globulin (α2HSG, Fetuin-A) (see Fig. 1). In alkalosis, hydrogen ions dissociate from albumin and globulins, and calcium binding to albumin and globulins increases. In addition, in alkalosis there is an increase in calcium complex formation. As a result, the concentration of ionized calcium decreases and this may result in clinical symptoms of hypocalcemia, although total plasma calcium concentration is unchanged. On the contrary, in acidosis Dynamics of Bone and Cartilage Metabolism

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Figure 1

The equilibrium between protein-bound, complexed, and ionized calcium in blood is shown. Some calcium is bound as a calcium-phosphate complex to α2-HeremannSchmid globulin (α2HSG, Fetuin-A).

The reference range of ionized calcium in venous blood in mothers is as in adults (1.20 ± 0.035 (mean ± SD)), however, neonates show higher levels (1.48± 0.055 mmol/L, venous cord blood) [3]. Within 24 hours, the ionized calcium levels drop to 1.20 mmol/L. From day 3 to 11, the levels in newborns increase and approach 1.38 mmol/L after day 7 [3]. The so-called physiological hypocalcemia in the first 3 days is no true hypocalcemia, if compared to adults. It is suggested that it takes some days until the parathyroid glands are fully active, as these are suppressed by the fairly high calcium levels at birth.

Table I. Reference Range of Calcium in Serum (or Plasma, Ammonium Heparinate) [1, 2]) Age Total calcium Children <28 days 1–12 months 1–20 years Ionized calcium Adults [1] Adults [2]

mg/dl

mmol/L

7.1–10.8 8.2–10.8 8.6–10.6

1.75–2.70 2.05–2.70 2.14–2.65

4.6–5.4 4.6–5.3

1.15–1.35 1.16–1.32

The fraction of ionized calcium/total calcium is 47–57%. The relative molecular mass of calcium is 40.08. Conversion factors: mmol/L × 4.008 = mg/dl mg/dl × 0.2495 = mmol/L (mmol/L × 2 = mEq/L) (mEq/L × 0.500 = mmol/L).

2. CURRENT ASSAYS AND TECHNOLOGY FOR THE MEASUREMENT OF CALCIUM

a. Atomic Absorption Spectrometry (AAS) This is the reference method for the determination of total calcium [4, 5]. AAS relies on the principle that electrons in the outermost shell of a particular atom absorb radiant energy and become excited. The energy absorbed by the atom is generally in the form of a very narrow absorption line either within the ultraviolet or visible spectrum. The flame is primarily used to disperse and release the atoms from their molecular states. Calcium in serum and urine is diluted sufficiently with lanthanum chloride solution. Lanthanum, in the presence of HCl, eliminates the effects of phosphate in both aqueous and serum samples. In addition, lanthanum acts as a releasing agent enhancing the dissociation of calcium from serum proteins. When introduced into a flame, the dissociated free calcium atom absorbs light from the characteristic wavelengths (e.g. 422.7 nm) produced by a hollow cathode lamp with a calcium filament. The imprecision within the series is <1% and between series <2%. Under optimized conditions the linearity of the method is excellent (coefficients of correlation of 0.9999 are obtained in the range of 0–140 mg/L). With modern equipment, calcium, magnesium, and even more elements may be determined simultaneously. Producers of automated AAS equipment include Hitachi (Japan), Shimadzu (Japan), Perkin Elmer (Norwalk, Connecticut, USA), and Varian (Mulgrave, Victoria, Australia).

Chapter 29 Measurement of Calcium, Phosphate and Magnesium

b. Flame Atomic Emission Spectroscopy (FAES) For routine purposes, flame atomic emission spectroscopy and photometry (see below) are used in many clinical chemistry laboratories. Flame atomic emission spectroscopy: serum or plasma is diluted with distilled water (which may contain lithium), sprayed into a flame of acetylene/air (2300⬚C) and vaporized. Some interference of sodium is corrected by a sodium-containing solution (140 mmol/L) for calibration. Simultaneously, the emission of sodium, potassium, lithium, and calcium may be measured. Lithium is used as an internal standard correcting for variations in flame intensity. FAES is used for high-throughput automated multichannel clinical chemistry analyzers (e.g. Olympus Co., Japan) or high-throughput stand-alone electrolyte analyzers (sodium, potassium, calcium; e.g. Eppendorf, Hamburg, Germany; production of new analyzers has now been stopped). By technical improvement in recent years, highly precise measurements are possible with coefficients of variation within series below 0.5% and from series to series below 0.8%. In addition to serum calcium determination, we use FAES successfully for the determination of calcium in feces (see below). A comparison of FAES with AAS has been published [6]. FAES showed a mean deviation of the target values obtained in commercially available control sera of −0.1% from the reference method (AAS). c. Photometry Calcium complexes with o-cresolphthalein complexone (OCPC) [7] or arsenazo III [8]. At pH 10–12 calcium yields a red complex with OCPC, which is measured at 570–575 nm. The color intensity of the purple complex formed is directly proportional to the calcium concentration. The complex is stabilized by KCN, thereby eliminating interference from heavy metals. By adding 8-hydroxychinoline, interference of magnesium is eliminated [9]. Photometry is used in many high-throughput automated multichannel clinical chemistry analyzers (see Table II). Using photometry, the arsenazo III method increased calibration stability (from 12 hours Table II. Methods Commonly Employed for the Determination of Calcium in Routine Laboratories (in alphabetical order according to the producers) Producer, Analyzer Abbott Architect® Bayer ADVIA® BeckmanCoulter LX® DadeBehring Dimension® Olympus AU® Roche Diagnostics, Integra®, Modular®

Method for calcium determination arsenazo-III salt o-cresolphthalein complexone ISE, indirect o-cresolphthalein complexone o-cresolphthalein complexone o-cresolphthalein complexone

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using o-cresolphthalein complexone to 48 hours with arsenazo III [8]). Within assay coefficients of variation of 1% and between assay coefficients of variation of <2% are obtained. d. Interferences in the Photometric Methods d.1. Hemoglobin: Interference in the OCPC method has been reported by hemoglobin. The reported effects of hemoglobin on the calcium–OCPC reaction are variable. In general, no significant interference has been found, although some studies have shown a distinct positive interference and one investigation suggested a negative effect (for references see [10]). d.2. Gadolinium: Recently, several publications have appeared about spurious hypocalcemia with the OCPC method applied on sera of patients who had received gadolinium [11–14]. In an in vitro study differences of 1.9, 0.9, and 0.55 mg/dl at 0.5, 0.25, and 0.125 mmol/L gadolinium, respectively, were found [13]. In contrast, these authors observed in vitro falsely increased calcium values when samples were measured by the arsenazobased method: the differences were 0.6 and 0.3 mg/dl at 1.0 and 0.5 mmol/L gadolinium. However, in vivo falsely decreased calcium values after gadolinium were observed when samples were measured by the arsenazo-based method [15]. The effect of gadolinium formulations may last from 24 h up to 188 h, depending on the glomerular filtration rate. It is recommended to use the ionized calcium, AAS or FAES methods in these patients [15] (and personal recommendation). Gadolinium chelate formulations are administered as contrast agents in magnetic resonance imaging (see also the possible interference of gadolinium chelate in the assay of magnesium). d.3. Paraproteins: Pseudohypercalcemia has been reported in two patients with IgM paraproteinemia [16]. In these, elevated calcium levels of about 3.0 mmol/L (12 mg/dl) were observed by the arsenazo method, and normal results were obtained by the OCPC method, by atomic absorption spectrophotometry, and by measurement of ionized calcium. d.4. Citrate: Citrate interfered in the arsenazo III method, whereas there was no interference in the OCPC method [17]. Table II gives the methods commonly employed for the determination of calcium in routine laboratories. e. Specimen Collection and Preparation for the Methods Employed in Table II Collect serum using standard sampling tubes, or heparinized plasma (serum is the preferred specimen). Plasma must be assayed fresh, as some fibrin formation may occur after hours and may occlude pipetting tips (centrifuge samples containing a precipitate before performing the assay).

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Do not use EDTA, oxalate, or citrated plasma. f. Stability of Serum Samples for the Photometric Methods (Package Insert Roche Diagnostics) • 7 days at 20–25⬚C • 3 weeks at 4–8⬚C • 8 months at −20⬚C. g. Enzymatic Method for Measuring Calcium An enzymatic method of assaying calcium in serum and urine with porcine pancreatic α-amylase has been published [18]. In this assay, calcium activates α-amylase yielding a product which is monitored at 405 nm. Another enzymatic assay of calcium in serum using phospholipase D with good precision, analytical recovery, and correlation to AAS has been published [19]. h. Ion-selective Electrodes (ISE) for Ionized Calcium A typical Ca2+ ion-selective electrode (Figure 2, [20]) consists of an assembly in which a Ca2+-selective membrane encloses an inner reference solution of CaCl2 held within a stem of plastic or glass. An internal reference electrode, usually Ag/AgCl, is immersed in the inner reference solution. In its simplest form this consists of a standard aqueous solution of CaCl2, usually saturated with AgCl to prevent dissolution of the AgCl from the internal reference electrode, and containing NaCl and KCl in physiological concentrations. Calcium ISEs in routine use in clinical chemistry are based on so-called liquid membranes in which an ionselective electroactive substance or “ionophore” is dissolved

Figure 2

in an organic liquid phase which is trapped within a polymeric matrix, usually polyvinyl chloride (PVC). The cell is completed by an external reference electrode in junction with the analyte solution. The affinity of the ion exchanger for the free ionized calcium imparts the selective permeability to the membrane, though with a very high resistance to mass transport. According to the Nernst equation, the cell will develop a potential for a given free ionized calcium activity in the analyte solution. The commercialized equipment with thermostatcontrolled flow-through electrodes, automated calibration, and pH correction has made the determination of ionized calcium as easy as modern blood gas analysis [21]. Withinassay coefficients of variation of 1% and between-assay coefficients of variation of <2% are obtained [22]. The measurement of ionized calcium has been evaluated in anaerobic whole blood, plasma, and serum and the authors recommended anaerobic whole blood as the preferred specimen [23]. Equipment is available from (alphabetical order), e.g. Applied Medical Technology (Menlo Park, CA, USA); AVL (Graz, Austria; and Roswell, Georgia, USA); CHIRON DIAGNOSTICS (Walpole, USA); Corning, (see Chiron) (USA); KONE (Espoo, Finland); Nova (Waltham, MA, USA); Orion (Cambridge, MA, USA); Radiometer Copenhagen (Brønshøj, Denmark). i. Fluorometric Titration Fluorometric titration with ethylene glycol tetraacetic acid (EGTA) is used in the Corning 940 calcium analyzer (Corning, USA). This method has a wide linear range,

Schematic diagram of a typical flow-through ISE-reference electrode measuring cell ([20], with kind permission of the publisher).

Chapter 29 Measurement of Calcium, Phosphate and Magnesium

and good precision (coefficients of variation of 1–3% within series) and recovery [24]. However, this method is susceptible to interferences by copper, iron, zinc, chelating agents, and certain drugs such as sulfadiazine, heparin, and acetylsalicylic acid [2]. This method has been very successfully used for the determination of serum and urinary calcium, especially in the smaller laboratory and in pediatric laboratories. 3. PREANALYTICAL AND BIOLOGICAL VARIABILITY OF SERUM CALCIUM

Preanalytical variability is mainly caused by posture and prolonged venous occlusion if blood samples are taken using a tourniquet. The effect of venous occlusion on serum calcium, protein, phosphorus, and magnesium concentrations is shown in Table III. The correction of calcium to total protein in Table III was performed according to Husdan [26], which is a slight modification of a formula obtained by Parfitt [27]. There exist different formulas for the correction of serum calcium. 1. Correction of serum calcium to albumin according to Payne [28]: corrected Ca (mg/dl) = measured Ca (mg/dl) – albumin (g/dl) + 4.0 or, recalculated for the SI system of units: corrected Ca (mmol/L) = measured Ca (mmol/L) – 0.025 × albumin (g/L) + 1.0.

2. Correction of serum calcium to total protein according to Husdan [26]: corrected Ca (mg/dl) = measured Ca (mg/dl)/(0.6 + TP/19.4) where TP is the serum total protein concentration (g/dl), or, recalculated for the SI system of units: corrected Ca (mmol/L) = measured Ca (mmol/L)/(0.6 + TP/194) where TP is the serum total protein concentration (g/L).

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protein concentration of 77.6 g/L the corrected calcium is the same as the measured value. We compared the formula of Husdan [26] with the formula of Payne [28] by applying these to patients with primary hyperparathyroidism and found a better separation of these groups from the normal range by the formula of Husdan. It is assumed that the method of albumin determination influences the albumin correction method. In addition to effects of venous occlusion, physiological protein fluctuations in a standing or a recumbent position are mainly responsible for intraindividual serum calcium variability [29]. The effect of standing or recumbent position on the different fractions of serum calcium is shown in Figure 3. Furthermore, some false low-normal ranges have been published in healthy blood donors, as in these the connection to the blood bag is cut after the donation and an extra tube is then collected for the determination of calcium [30]. After blood donation, however, the serum protein concentration is significantly lower than before the donation. a. Circadian Rhythm and Seasonal Variation The circadian rhythm of total serum calcium and ionized calcium is shown in Figure 4A, of ionized serum calcium and intact parathyroid hormone in Figure 4B, and of protein-corrected serum calcium (according to Husdan [26]) and intact parathyroid hormone in Figure 4C. By applying the formula of Husdan [26], we observed nearly no circadian variation of serum calcium from 9.00 a.m. to 5.00 p.m. (only a minor decrease from 2.45 to 2.42 mmol/L as shown in Figure 4C, FAES method for calcium, biuret method for total protein, [31]). Seasonal variations were not observed for serum calcium in a health examination survey in more than 2000 men [32].

At a total protein concentration of 77.6 g/L divided by 194 a value of 0.4 is obtained, which means that at a

Table III. Effect of Venous Occlusion on Serum Calcium, Protein, Phosphorus, and Magnesium Concentrations [25] Constituent Total calcium (mg/dl) (mmol/L) Total protein (g/dl) Corrected calcium (mg/dl) (mmol/L) Phosphorus (mg/dl) Magnesium (mg/dl) Ionized calcium (mg/dl) (mmol/L)

Baseline

3-min tourniquet

P

9.64 2.41 6.9 10.11 2.528 3.22 2.09 4.62 1.155

9.87 2.47 7.3 10.09 2.523 3.19 2.13 4.62 1.155

<0.01 <0.001 n.s. n.s. n.s. n.s.

Figure 3

Physiological protein fluctuations in a standing or a recumbent position are mainly responsible for intraindividual serum calcium variability (• standing,
recumbent; [29], with kind permission of the publisher).

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the first hour. A statistically significant increase of serum calcium was noted after 30 min by use of the FAES method (Eppendorf “EFIX 5055”, Hamburg, Germany) and correction of calcium for total protein according to Husdan, and after 60 min by use of the ionized calcium determination (for details see reference [31]). The advantage of the FAES method may be due to the high precision. If in patients with hypercalcemia ionized calcium is compared with total calcium without correction for total protein or albumin, ionized calcium appears to be a slightly better indicator of elevated calcium states than total calcium [33]. This has also been found in primary hyperparathyroidism [34], as shown in Figure 5. Figure 5 shows that by measuring ionized serum calcium, patients with primary hyperparathyroidism are separated from the normal range, whereas there is some overlap with the normal range with total serum calcium. A case with severe hypercalcemia (total serum calcium of 3.5 mmol/L) was observed in a patient with multiple myeloma and markedly elevated serum IgA κ-paraprotein concentration. Symptoms of hypercalcemia were absent, and serum ionized calcium was normal. This pseudohypercalcemia was due to calcium binding by this paraprotein [35]. A similar case with a calcium-binding IgG myeloma globulin with persistently increased total serum calcium and normal ionized calcium had already been described in 1973 [36]. A summary of causes of hyper- and hypocalcemia is given in Table IV.

C Figure 4 The circadian rhythm of (A) total serum calcium and ionized calcium, (B) ionized serum calcium and intact parathyroid hormone and (C) protein-corrected serum calcium and intact parathyroid hormone [26].

b. Biases and Errors, or: is there a Preferred Method? We gave 1000 mg of calcium dissolved in water (Calcium Sandoz fortissimum, effervescent calcium) to 12 healthy young men at 9.00 a.m. and monitored serum calcium for 8 hours. Blood was taken every 15 min during

Figure 5 Correlation of serum-ionized calcium and serum total calcium in the preoperative sera of 48 patients with documented primary hyperparathyroidism. The crosshatched areas represent the normal range (mean ± 2 S.D.) ([34], with kind permission of the publisher).

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Chapter 29 Measurement of Calcium, Phosphate and Magnesium Table IV.

Causes of Hypercalcemia and Hypocalcemia

Hypercalcemia

Hypocalcemia

Hyperparathyroidism (HPT), primary Renal failure (tertiary HPT) Renal transplantation (persistent HPT) Malignancy Immobilization Thiazide diuretics Lithium Aluminium intoxication Vitamin D or A intoxication Granulomatous disease Thyrotoxicosis Milk-alkali syndrome Familial hypocalciuric hypercalcemia (FHH) Erroneous (see text)

Hypoparathyroidism, pseudohypo-PT after parathyroidectomy Chronic renal insufficiency (sec. HPT) Malignancy Extreme physical activity (ionized Ca++d) Calcimimetic drugs Vitamin D deficiency Septic shock, pancreatitis, Rhabdomyolysis After thyroidectomy Calcium deficiency rickets Autosomal dominant hypocalcemia (ADH) Erroneous (see text)

FHH and ADH are inborn errors with deactivating or activating mutations of the calcium-sensing receptor.

Electrolyte-balanced heparin used in syringes may produce a bias in the measurement of ionized calcium concentration in specimens with abnormally low protein concentration [37]. However, in patients with multiple myeloma or hyperlipidemia, ionized serum calcium is superior to FAES or photometry, since in the latter methods the volume of serum proteins or serum lipids decreases the volume of serum water, so that FAES or photometry yield falsely low values for electrolytes. In patients receiving large amounts of citrate (multiple transfusions, cardiopulmonary bypass surgery, liver transplantation) the monitoring of ionized calcium is of great importance [38, 39]. In critically ill multiple-trauma patients, the accuracy of methods to estimate protein or albumin corrected serum calcium concentrations has recently been investigated. In this population, corrected serum calcium concentrations had a poor sensitivity (about 25 ± 32%) for assessing hypocalcemia [40]. For this population, direct measurement of serum-ionized calcium is indicated for assessing the calcium status. c. Erroneous Low Results Obtained by all Methods Patients on long-term hemodialysis via arteriovenous fistula received heparin when the fistula needle was inserted, before a sample of blood was obtained for chemical analysis. The resultant release of lipoprotein lipase activity in vivo and continued lipolytic activity in vitro sometimes produced sufficient free fatty acid to precipitate calcium soaps. The consequent spurious hypocalcemia was most frequently observed when the patients had chylomicronemia. This cause of apparent hypocalcemia was eliminated either by immediate analysis of the

blood samples or by obtaining samples before systemic heparinization [41]. For routine laboratory services for hospitals or practicing physicians, FAES or photometry are clearly adequate. However, implausible results obtained by photometry should be checked by other methods. Some routine hospital laboratories have switched from total calcium to ionized calcium. For intensive care units with patients receiving multiple transfusions or large amounts of citrate, the determination of ionized calcium is necessary.

B. Urinary Calcium Different guidelines for the collection and treatment of urine samples for the determination of calcium are found in the literature, as shown in Table V. Weber (Table V) found in freshly collected urine samples a mean calcium concentration of 2.39 mmol/L with and without acidification by HCl. Therefore the following recommendation is given: if electrolytes (sodium, potassium, calcium, magnesium, chloride), glucose, uric acid, urea, creatinine, albumin, protein, α-amylase, or β-NAG are to be determined in urine samples with modern automated photometric equipment (e.g. see Table II), flame photometers or atomic absorption spectroscopy, the urine should be collected without additives and analyzed within hours after collection according to this literature survey. Acidification would decrease the concentration of uric acid and excludes pH determination in the collected sample. a. Reference Range of 24-hour Urinary Calcium Similar to Coe [43] we define a 24-hour urinary calcium excretion of 120–320 mg (3–8 mmol) as normal for men, and 100–280 mg (2.5–7 mmol) for women, or 2–4 mg/kg body weight (either sex). According to the literature [44–47], we regard in infants 1.1–9.0 mg/kg/day as normal, and in children from 1 to 6 years 1.0–9.0 mg/kg, from 6 to 12 years 1.0–6.0 mg/kg and after stopping growth 2–4 mg/kg or, as stated above, 120–320 mg (3–8 mmol) as normal for men, and 100–280 mg (2.5–7 mmol) for women. With increasing age, a decrease in the urinary calcium excretion calculated in mg/kg occurs [46]. This is understandable: at a weight of an adult man of 75 kg a calcium excretion of 4 mg/kg/day results in a total excretion of 300 mg/day, which is regarded as the upper limit of the normal calcium excretion. Therefore, the 8 mg/kg found as an upper limit in children [45, 46] must decrease with age to 4 mg/kg in the adult. We recommend excluding vitamin D deficiency by measuring 25-hydroxyvitamin D in all children and

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HEINRICH SCHMIDT-GAYK

Table V. Review of the Literature [42] for the Collection and Storage of Urines if Calcium Determination is Intended Author

Year

Recommendation

Documenta Geigy

1968

Doerner K

1992

Frings CS Gowans EMS

1975 1986

Greiling G

1995

Guder WG

1986

Gunn IR Hauptmann Ch Jahnen A

1992 1992 1989

Keller H

1991

Manz F Misselwitz J Ng RH

1984 1981 1984

Nordin BEC

1976

Paunier L

1970

Richterich R

1971

Thomas L

1998

Tietz NW

1987

Weber A

1989

Ca-urate is precipitated in acidified urine Add 5 ml concentrated HCl after collection No addition of HCl Add HCl after collection until pH 5–6 is obtained Add 5 ml HCl, 6 mol/L, to the container before collection is started, after collection acidify with HCl to pH 1 Add thymol to the container before collection is started Acidify after collection No addition for pediatric purposes No difference between acidified and untreated divided samples No addition: analyze as soon as possible after collection No addition for pediatric purposes No addition for pediatric purposes No difference between acidified and untreated samples if assayed within 6 hours Add something against bacterial growth (e.g. thymol) before collection is started, after collection fill 25 ml into a separate container and add two drops of concentrated HCl Add 10 ml of concentrated HCl/L urine after collection (pediatrics) Add thymol to the container before collection is started, after collection add 10 ml of concentrated HCl/L urine Add 10 ml of concentrated HCl after collection and raise the temperature of the sample Add 5 ml HCl, 6 mol/L, to the container before collection is started: after collection acidify with HCl to pH 3–4: do not use more than 10 ml of HCl, 6 mol/L Immediately after collection there was no difference between samples collected with and without acidification by HCl

adults with a urinary calcium below 1.5 mg/kg body weight. (Note: the most sensitive indicator of vitamin D deficiency is lowered urinary calcium excretion, and the most sensitive indicator of vitamin D excess is elevated urinary calcium excretion.)

b. Factors Influencing the Urinary Excretion of Calcium Increased urinary calcium excretion is observed in many patients with increased intestinal calcium absorption and/or increased bone resorption (hypercalcemia). However, since parathyroid hormone increases the tubular reabsorption of calcium, some patients with primary hyperparathyroidism present with normal urinary calcium excretion. Urinary calcium is often increased in recurrent stone formers and is significantly related to urinary sodium excretion (= dietary sodium consumption). In a recent review it was summarized that salt-loading studies and population observations indicate that for every 100 mmol of sodium (= 6 g NaCl) excreted there is approximately 1 mmol loss of urinary calcium in free-living, normocalciuric healthy populations [48]. In addition, urinary calcium is related to urinary urea excretion (dietary protein consumption). A relationship between protein overconsumption and up-regulation of calcitriol (1,25-dihydroxyvitamin D3) synthesis has been found [49]. Whereas a high dietary calcium intake decreases the risk of kidney stones, probably by inhibiting intestinal oxalate absorption, additional pharmaceutical calcium intake in large doses (1000 mg or more) may increase the risk of kidney stone formation by hypercalciuria, which is detectable for some hours following large oral doses. Therefore it is recommended that additional pharmaceutical calcium intake should be divided into two doses of 500 mg each (one taken in the morning, one in the evening) or should even be divided into three doses. In healthy persons, for every 1000 mg (25 mmol) of additional oral calcium intake, urinary calcium increases by about 60 mg (1.5 mmol). The metabolic evaluation of patients with recurrent idiopathic calcium nephrolithiasis yielded the following results: the most frequent risk factor was hypercalciuria (39%), followed by hyperoxaluria (32%) and low volume (32%), hypocitraturia (29%), hyperuricosuria (23%), and hypomagnesiuria (19%) [50]. After immobilization, e.g. after tetraplegia, urinary calcium increases to amounts of about 600 mg/24 h caused by excessive bone resorption. c. Diurnal and Seasonal Variation of Urinary Calcium Excretion In normal individuals, a diurnal variation of urinary calcium excretion as pronounced as in fasting individuals has been found [51]. The maximum of urinary calcium excretion is observed between 10.00 a.m. to 2.00 p.m. according to Min [51] and 10.00 a.m. to 6.00 p.m. according to Wisser [52]; fasting from 4.00 to 8.00 a.m. (Min), and the minimum from 6.00 to 12.00 p.m. (fasting and nonfasting) (Min). In contrast to Min, the minimum is

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observed from 4.00 to 7.00 a.m. according to Wisser. The differences between the Min and Wisser studies may be explained by different conditions: the subjects in the Min study were fed every 3 hours, whereas the subjects in the Wisser study had meals at 8.00 a.m., 11.00 a.m., 2.00 p.m., 5.00 p.m., and 8.00 p.m. (For the diurnal variation of calcium see this Chapter, 29/I/3). The diurnal variation of urinary calcium excretion in fasting persons is remarkably similar to the diurnal variation of sodium excretion [51]. For the detection of hypercalciuria, an overnight urine collection may be used for screening purposes. We did not observe a seasonal variation of urinary calcium in patients with calcium-containing kidney stones. In contrast, Robertson [53] found increased urinary calcium in the summer months compared to the winter months. The results of Robertson might be explained by a lesser vitamin D supply in the winter months in Leeds (England) than in Heidelberg (Germany). d. Detection of Increased Calcium Resorption from Bone The basal rate of calcium excretion can be assessed from the calcium/creatinine ratio in the urine after a 12-hour fast according to Nordin [54]. The normal range of this value is 0.02–0.16 if each concentration is expressed in mg/100 ml (note: a ratio of 0.02–0.16 means that 20–160 mg of calcium is excreted per gram creatinine). The calcium measured in this test comes mainly from bone and reflects net bone resorption, i.e. the difference between the rates of mineralization and resorption. A raised fasting urinary Ca/Cr indicates an increase in net bone resorption and is frequently found in otherwise normal postmenopausal women, in primary hyperparathyroidism, in hyperthyroidism, in multiple myeloma, and in most conditions associated with hypercalcemia unless renal failure is present [54]. In hypercalcemic states, relatively low urinary calcium is observed in thiazide medication, Addison’s disease (cortisol and mineralocorticoid deficiency) and familial hypocalciuric hypercalcemia (FHH). Low urinary calcium excretion is observed in renal insufficiency, vitamin D deficiency, and other hypocalcemic states. e. Test Procedure for the Determination of Increased Calcium Resorption from Bone The evening before the test no milk products are allowed. On the morning of the test the first morning void urine is discarded (e.g. at 7.00 a.m.), then 500 ml of distilled water (or otherwise water with a very low calcium content, i.e. less than 10 mg/L) is ingested and 2 hours later urine is collected and 10 ml are sent to the laboratory for the measurement of calcium and creatinine. (Note: we recommend collecting also the first morning void urine and to separate 10 ml for the measurement of total pyridinium

crosslinks to detect increased bone collagen resorption. The first morning void urine can, however, not be used for the determination of the basal calcium excretion rate, because some of the calcium in this urine is still derived from intestinal calcium absorption. The intestinal calcium absorption from the evening meal is finished at 7.00 a.m. The second morning void urine may, however, be less well suited to detect increased bone collagen resorption [55].)

II. MEASUREMENT OF PHOSPHATE A. Phosphate in Serum 1. INTRODUCTION

Phosphate is present in blood chiefly as inorganic and organic phosphates, nearly all of the latter residing in the erythrocytes. Determination is often referred to as inorganic phosphorus rather than inorganic phosphate, since the phosphate determined is usually reported in terms of phosphorus [2]. Only inorganic phosphate is measured in the routine clinical settings. In normal healthy subjects the total phosphate concentration in plasma is 3.9 mmol/L, of which only 0.8–1.4 mmol/L is inorganic phosphate, the remaining being phospholipids and other organic compounds (for review on plasma phosphate see reference [56]). Approximately 10% of plasma inorganic phosphate is nonfiltrable or protein-bound, and 6% is complexed with calcium or magnesium; 84% of plasma phosphate is free and present in form of the ions H2PO4− and HPO42−, the relative proportion depending on the pH: the ratio varies between 1:1 in acidosis, 1:4 at pH of 7.4, and 1:9 in alkalosis [2]. The extracellular phosphorus pool and therefore the serum phosphate level is influenced by phosphate intake, intestinal absorption, exchange with the bone reservoir, and renal excretion [57]. 2. FUNCTIONS OF PHOSPHATE IN VIVO

The serum level of phosphate has many functions in mineral and bone metabolism; it increases the production of parathyroid hormone (PTH), inhibits the renal 1α-hydroxylase (formation of calcitriol = 1α,25dihydroxyvitamin D) and influences urinary phosphate excretion, as shown in Figure 6. In addition to its functions shown in Figure 6, phosphate has important intracellular and extracellular functions. Phosphate is a constituent of nucleic acids, phospholipids, and phosphoproteins and therefore plays a structural role. Phosphate plays a central role in cellular metabolic pathways, including glycolysis and oxidative phosphorylation.

496

HEINRICH SCHMIDT-GAYK

Figure 6 The serum level of phosphate has many effects on calcium and bone metabolism: it increases the production of parathyroid hormone (PTH), inhibits the renal 1α-hydroxylase (formation of calcitriol = 1α,25-dihydroxyvitamin D) and influences urinary phosphate excretion (according to [3], with kind permission of the publisher).

Phosphate in the form of 2,3-diphosphoglycerate (2,3-DPG) is a regulator of dissociation of oxygen from oxyhemoglobin and thereby of the delivery of oxygen to the tissues. 2,3-DPG accounts for about 80% of organic phosphate in erythrocytes. Phosphate forms high-energy compounds (ATP) and cofactors (NADP) and is critical in maintaining many physiological functions such as muscle contractility, neurological function, and electrolyte transport. a. Reference Range The reference ranges for fasting plasma or serum phosphate vary greatly with the age of the patient. During the first 6 months of life, phosphate concentrations are 1.6–2.5 mmol/L (5.0–7.8 mg/dl). Thereafter the values

decrease significantly. At 4–8 years of age the phosphate concentration range is 1.2–1.8 mmol/L (3.7–5.4 mg/dl), at 10 years of age 1.1–1.7 mmol/L (3.5–5.3 mg/dl). No sex difference is observed in childhood, but during puberty the phosphate concentration decreases abruptly to the range observed in young adults: 0.8–1.45 mmol/L (2.5–4.5 mg/dl) in males and 0.9–1.55 mmol/l (2.8–4.7 mg/dl) in females [58]. The levels diminish slightly in adults of both sexes until middle age; thereafter they increase in elderly females and decrease slightly in elderly males. Increased bone resorption occurring in a substantial proportion of women after the menopause may be responsible for some increase in serum phosphate in this group. The reference range in children and adults is given in Table VI.

Table VI. Reference Range for Fasting Plasma or Serum Phosphate in Children [59] and Adults [60] Age

Male (n)

Male mg/dl

Male mmol/L

Female (n)

Female mg/dl

1–30 d 31–365 d 1–3 y 4–6 y 7–9 y 10–12 y 13–15 y 16–18 y 19–25 y 26–80 y

62 83 126 112 117 135 109 95 >30§ >100§

3.9–6.9 3.5–6.6 3.4–6.0$ 3.3–5.7$ 3.3–5.7$ 3.2–5.7 2.9–5.1 2.7–4.9 2.8–4.8 2.5–4.5

1.25–2.25 1.15–2.15 1.10–1.95 1.05–1.85 1.05–1.85 1.05–1.85 0.95–1.65 0.90–1.60$ 0.90–1.55 0.80–1.45

66 66 119 107 107 115 110 122 >30§ >80§

4.3–7.7 3.7–6.5 3.4–6.0 3.2–5.5 3.2–5.5$ 3.2–5.3$ 2.8–4.8 2.8–4.8$ 2.8–4.8 2.5–4.5

$mean

value of Soldin [59] and Greenberg [60] data. counted from Figure 1 in Greenberg [60]. Relative molecular mass of phosphate (as phosphorus, inorganic): 30.97. Conversion factors: mmol/L × 3.097 = mg/dl mg/dl × 0.3229 = mmol/L.

§number

Female mmol/L 1.40–2.50 1.20–2.10 1.10–1.95 1.05–1.80 1.05–1.80 1.05–1.70 0.90–1.55 0.90–1.55 0.90–1.55 0.80–1.45

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Chapter 29 Measurement of Calcium, Phosphate and Magnesium

Causes of hyperphosphatemia and hypophosphatemia are given in Table VII.

Table VII. Causes of Hyperphosphatemia and Hypophosphatemia (modified from Weisinger [61], additions have citations)

3. CURRENT ASSAYS AND TECHNOLOGY

a. Molybdate Method for the Determination of Inorganic Phosphate Most methods depend on the reaction of phosphate ions with molybdate to form a colorless phosphomolybdate complex such as (NH4)3[PO4(MoO3)12], which can be measured directly by UV absorption at 340 nm or after reduction to a complex heteropolymer, molybdenum blue, at 600–700 nm [2]. The former method is the most commonly used by discrete analyzers on account of its speed, good precision, accuracy, and reagent stability. To prevent protein precipitation, a solubilizing agent such as Tween 80 is necessary in the reagents. In the application of the Roche Diagnostics GmbH, Mannheim, Germany, for the Hitachi automated analyzers, e.g. Hitachi 717 or Modular, the final concentrations of the reagents in the assay are ammonium molybdate 1.0 mmol/L; sulfuric acid 0.35 mol/L; sodium chloride 44 mmol/L; and a detergent. Measurement is performed at 340 nm, with an additional reading at 660 nm to correct for interference. In another adaptation, a blank absorbance is measured at 340 nm and at 376 nm [69]. The analyzers commonly employed for the determination of phosphate in routine laboratories (in alphabetical order of the producers), such as Abbott Architect®, Bayer® ADVIA, BeckmanCoulter LX®, DadeBehring Dimension®, Olympus AU®, and Roche Diagnostics (Hitachi, Integra®, Modular®) use the phosphomolybdate method. a.1. Precision and Accuracy The intra-assay coefficient of variation (imprecision) is usually less than 2% for the phosphomolybdate method, and the interassay coefficient of variation is usually below 3% with automated analyzers. Recovery is in the range of 98% to 103%. a.2. Interferences Turbidity may interfere in the phosphomolybdate method, yielding falsely high values. In addition, high hemoglobin and bilirubin concentrations interfere. High citrate concentrations yield falsely low results since citrate competes with phosphate for binding at the molybdate ion. In addition, mannitol from infusion therapy [65] and phenothiazines form complexes with phosphomolybdate, yielding falsely low values [66]. a.3. Paraprotein Interference Pseudohyperphosphatemia has previously been reported in patients with paraproteins [70–72]. Some studies have shown that interference is due to turbidity caused by precipitation of the paraprotein by the acid reagent used in both visible and ultraviolet spectrophotometric methods [73–77]. Paraprotein precipitation in the reaction mixture occurred frequently in IgM and, in particular, IgG paraproteins [73]. One case has been

Hyperphosphatemia Increased exogenous load or increased intestinal absorption Intravenous infusion Oral supplementation Cow’s milk feeding to premature babies Vitamin D intoxication Phosphate-containing enemas Acute phosphorus poisoning Increased endogenous load Tumor lysis syndrome Rhabdomyolysis Bowel infarction Malignant hyperthermia Hemolysis Acid–base disorders (lactic acidosis, diabetic ketoacidosis, respiratory acidosis)

Reduced urinary excretion Renal failure Hypoparathyroidism Magnesium deficiency (r Hypoparathyr.) Acromegaly Tumoral calcinosis

Pseudohyperphosphatemia Multiple myeloma Hemolysis in vitro; serum not separated from erythrocytes within six hours Hypertriglyceridemia Sample drawn from indwelling catheters: contamination by phosphate-containing heparinized saline [67]

Hypophosphatemia Decreased exogenous load or decreased intestinal absorption Dietary phosphate restriction Antacid abuse (aluminum hydroxide) Chronic diarrhea Vitamin D deficiency, VDDR, VDRR§ Steatorrhea Alcohol abuse Altered internal redistribution Recovery from malnutrition Recovery from diabetic ketoacidosis Hormonal and other agents (insulin, glucagon, epinephrine, cortisol, glucose, fructose, lactate) Sepsis Hungry bone syndrome Respiratory alkalosis (pain, anxiety, salicylate poisoning, sepsis, heatstroke) Increased urinary excretion Hyperparathyroidism (HPT, primary) HPT, persistent after kidney transplantation Secondary HPT in gastrointestinal disease, e.g. malabsorption, vitamin D deficiency Volume expansion (e.g. high sodium load) Carbonic anhydrase inhibition Metabolic or respiratory acidosis FGF 23 excess: XLH, ADHR, TIO§§ Renal tubular defects (Fanconi’s syndrome) Pseudohypophosphatemia Multiple myeloma [62] Monoclonal gammopathy: excessive sample blankings result in hypophosphatemia [63] Bilirubin interference [64] Mannitol interference [65] Citrate interference [66] Phenothiazine interference [66]

§VDDR, Vitamin D-dependent rickets (1α-hydroxylase deficiency). VDRR, Vitamin D-resistant rickets (vitamin D receptor defect). §§FGF 23 is associated with a marked depression in proximal renal tubular reabsorption of phosphate and hypophosphatemia (review: [68]). Disorders with increased levels of FGF 23 are 1. XLH, X-linked hypophosphatemic rickets/osteomalacia (mutation of the PHEX gene, which encodes an endopeptidase and results in less degradation of FGF 23) 2. ADHR, autosomal-dominant hypophosphatemic rickets (mutation of a FGF 23 which is less well degraded) 3. TIO, tumor-induced osteomalacia (ectopic overproduction of FGF 23).

498 reported with pseudohyperphosphatemia in association with a monoclonal gammopathy of undetermined significance [78]. Diluting the sample to 40 g/L total protein reduced but did not always eliminate the interference. In some cases, paraprotein concentrations as low as 8 g/L falsely increased plasma phosphate results [79]. Precipitation of multiple myeloma serum in the phosphorus channel of automated analyzers may also yield erroneously low values [80]. Other investigators have identified the presence of a phosphate-binding immunoglobulin [72, 81]. Savory and Pearce [13] evaluated an improved methodology of the Roche (formerly Boehringer Mannheim Co., (BM UK Ltd., Sussex, UK)) containing a detergent-modified reagent, which was compared with the standard kit supplied by the same manufacturer. Using the original reagent pseudohyperphosphatemia was found in 27% (12/45) of the samples. The modified reagent was free from interference with no significant difference between results from serum and protein-free supernatants. a.4. Method for the Evaluation of Hyperphosphatemia in Patients with Paraproteins Savory and Pearce [69] precipitated proteins using equal volumes of serum and 0.61 mol/L trichloroacetic acid. After mixing and centrifugation, the supernatant is analyzed directly. The same procedure has been used by others [82, 83]. Sulfosalicylic acid has also been used for deproteination [84]. Alternatively, the enzymatic method described below may be used for paraproteinemic sera. a.5. Enzymatic Method for the Determination of Inorganic Phosphate An enzymatic method for the determination of phosphate in serum and urine has been described, is marketed by Bayer Diagnostics (Munich, Germany), and applications have been published for the Hitachi 717 and Bayer/Technicon RA-XT analyzers [85]. In this method, phosphate ions react with inosine in the presence of purine nucleoside phosphorylase to form hypoxanthine; this is oxidized by xanthin oxidase to uric acid with production of hydrogen peroxide. The latter is determined with the aid of a chromogen system, the colored product being measured at 555 nm. Enzymatic phosphate determination on the Hitachi 717 and Bayer/Technicon RA-XT analyzers gave within-run coefficients of variation of 2.2% and day-to-day coefficients of variation of 3%. The adapted method was linear up to 3.2 mmol/L phosphate. No interference was found for bilirubin up to 270 µmol/L and for hemoglobin up to 114 µmol/L. The correlation (r = 0.99) to the automated molybdenum blue method was very good. The values of control sera were on average 4.1% lower than the declared values for the molybdenum blue methods [29]. Further purine nucleoside phosphorylasebased photometric assays for inorganic phosphate with good precision and accuracy have been published [86–88].

HEINRICH SCHMIDT-GAYK

b. Circadian Rhythm and Preanalytical Considerations Specimens should be obtained from a fasting patient in the morning, e.g. from 7.00 to 9.00 a.m., as there exists a decrease of phosphate after food ingestion, presumably because of insulin release, and a circadian variation. Hyperglycemia and hyperinsulinemia, as occurring during standard oral glucose tolerance tests, yield a hypophosphatemia which is caused by a shift of phosphate from the extracellular fluid into the cells [89, 90]. Circadian rhythm: a mean level of about 1.0 mmol/L is observed in healthy volunteers in the morning (7.00–10.00 a.m.), rising to 1.2 mmol/L at 3.00–5.00 p.m. and 1.3 mmol/L at midnight [91]. The circadian variation of phosphate in serum (together with intact PTH in EDTA plasma) in nonfasting healthy subjects is shown in Figure 7. It can be seen from Figure 7 that the level of phosphate is closely paralleled by the level of parathyroid hormone. The circadian rhythm of phosphate in plasma is abolished in fasting subjects [92]. A similar circadian rhythm as in healthy persons with higher levels in the evening and during the night was observed in patients on hemodialysis, but the levels were generally higher than those of healthy persons [93]. From the results shown in Figure 7 it is understandable that for a correct diagnosis in disorders of bone metabolism a standardized blood sample has to be obtained from a fasting patient in the morning, best before 08.30. Otherwise, many diagnoses of less severe cases, as in mild primary hyperparathyroidism or other hypophosphatemic disorders, might be missed or misinterpreted. Serum or plasma should be separated from cells within 4 h after collection of the specimen, thereafter the erythrocytes release phosphate and the concentration in serum or

Figure 7 The circadian variation of phosphate in serum (together with intact PTH in EDTA plasma) in nonfasting healthy subjects is shown.

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Chapter 29 Measurement of Calcium, Phosphate and Magnesium

plasma increases [94]. Hemolysis must be avoided because phosphate may be split off from labile esters in the erythrocytes. Plasma or serum may be stored at 4⬚C for several days, or in the frozen state for several months. The concentration of phosphate in serum is 0.06–0.10 mmol/L higher than that in plasma, because intracellular phosphate is released from platelets and erythrocytes during clotting [95]. This is especially marked in patients with thrombocytosis [96]. Citrate, oxalate, and EDTA should not be used as anticoagulants because they interfere with the formation of the phosphomolybdate complex. c. Phosphate Threshold The tubular maximum of phosphate reabsorption related to the glomerular filtration rate (TmP/GFR) is the best method to calculate renal tubular reabsorption of phosphate. TmP/GFR can be easily determined using the fasting urinary and plasma concentrations of phosphate and creatinine and a nomogram of Walton and Bijvoet [97]. Increased parathyroid hormone, volume expansion (e.g. increased sodium intake), or a phosphaturic factor in oncogenic osteomalacia decrease tubular phosphate reabsorption (lower TmP/GFR). TmP/GFR displays a similar circadian rhythm as described above for the serum phosphate levels: TmP/GFR starts at about 0.85 mmol/L in the morning and approaches 1.0 mmol/L in the evening and 1.2 mmol/L at 3.00 a.m. during sleep [89]. d. The Chloride/Phosphate Ratio in Hypercalcemia Serum chloride concentrations tend to be above 100 mmol/L in patients with primary hyperparathyroidism and serum phosphate concentrations tend to be below 1.0 mmol/L in this disorder (ratio > 100), if the samples are taken from fasting patients in the morning. On the other hand, serum chloride concentrations tend to be below 100 mmol/L in patients with hypercalcemia from other causes and serum phosphate concentrations tend to be above 1.0 mmol/L in these disorders (ratio < 100). Palmer and colleagues [98] found in primary hyperparathyroidism 96% of the patients with a ratio above 100 and in patients with hypercalcemia of other causes 92% below 100 (if phosphate is given in mg/dl the cut-off is 33). We measured the chloride/phosphate ratio in 288 patients with primary hyperparathyroidism and in 19 patients with malignancyassociated hypercalcemia (results not shown). We found that this ratio gives a good first estimation. However, this ratio has to be supplemented with further measurements, e.g. intact parathyroid hormone in EDTA plasma, to separate these groups more reliably.

B. Urinary Phosphate In healthy subjects the net intestinal absorption of phosphate is linearly related to dietary intake; about 50–75%

of ingested phosphate is absorbed and excreted into the urine. The bulk of the absorbed amount occurs by diffusion; only a small percentage is mediated by vitamin D metabolites, mainly by calcitriol. Therefore urinary phosphate is a valuable indicator of nutrition (meat, dairy products) and is often above the normal range of 25–35 mmol/24 h in overnutrition (overweight people). There exists a circadian rhythm for phosphate excretion with higher levels in the afternoon and during sleep (about 6 mmol/4 h) than in the morning (about 4 mmol/4 h) [89]. In fasting individuals, the circadian variation of urinary phosphate excretion vanishes [99]. Untimed urinary phosphate and calcium concentrations expressed as creatinine ratios were not helpful in detecting babies with rickets [100]. Urinary phosphate determination with analyzers employing dry chemistry may show nonlinearity at different dilutions of urine since this technique has been developed for serum. To circumvent this problem, urine samples may be diluted with serum instead of water [101]. 1. URINARY PHOSPHATE IN CHILDREN

The 24-hour urinary excretion of phosphate in 132 healthy children (mean age 8.3 years) has been investigated. The mean excretion was 532 µmol/kg body weight, the range was 312–741 µmol/kg body weight (2.5–97.5 percentile) [45]. In conventional units, the mean excretion was 16.5 mg/kg body weight, the range was 9.7–23.0 mg/kg body weight (2.5–97.5 percentile) [47].

III. MEASUREMENT OF MAGNESIUM A. Magnesium in Serum 1. INTRODUCTION

Magnesium is the most prevalent intracellular divalent cation and the second most prevalent cation in the body. The total body magnesium (TBMg) of an adult is approximately 25 g, of which 50–60% is in bone, and the remaining 40–50% is in the soft tissues [2]. One-third of the skeletal magnesium resides on the surface of bone, is exchangeable and serves as a reservoir for maintaining a normal extracellular magnesium concentration. Extracellular magnesium accounts for only 1% of total body magnesium (TBMg). Magnesium in plasma or serum is found in three fractions, namely the free or ionized magnesium (Mg2+, about 55%), a fraction of about 30% bound to proteins (mainly albumin), and a fraction of about 15% complexed with phosphate, citrate, and other anions [2]. Because only 1% of TBMg resides in the extracellular fluid (ECF), measurements of serum or plasma magnesium levels may not accurately reflect the level of TBMg stores. Therefore, some authors recommended the determination of magnesium in

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erythrocytes, mononuclear blood cells or muscle biopsies [102]. The primary diagnostic approach remains the serum magnesium measurement. Whole blood magnesium could be a better choice; however, even blood cell magnesium is unreliable because the magnesium content of blood cells correlates poorly with the magnesium content of the striated muscle and myocardium [102]. The requirement for a good magnesium test is demanding. Repeatedly reduced serum magnesium values strongly suggest underlying magnesium deficiency. The determination of magnesium excretion in a 24-h urine sample is helpful in critical cases [103]. 2. FUNCTIONS OF MAGNESIUM

Mg2+

is essential for the function of more than 300 cellular enzymes, including those related to the transfer of phosphate groups, all reactions that require ATP, and every step related to the replication and transcription of DNA and the translation of mRNA. This cation is also required for cellular energy metabolism and has an important role in membrane stabilization, nerve conduction, ion transport, and calcium channel activity [103]. In addition, magnesium plays a critical role in the maintenance of intracellular potassium concentration by regulating potassium movement through the membranes of the myocardial cells. Thus, magnesium deficiency can result in a variety of metabolic abnormalities and clinical consequences; these include refractory plasma electrolyte abnormalities (especially depressed potassium) and cardiac arrhythmias most often observed after stress such as cardiac surgery [103]. Magnesium and calcium both regulate the secretion of parathyroid hormone; this is shown in Figure 8.

Figure 8

Schematic diagram showing the relationships between the extracellular Ca2+ and Mg2+ concentrations and PTH release. The two divalent cations resemble one another in that high concentrations of either inhibit hormonal secretion. They differ, however, in that low concentrations of Mg2+ also inhibit PTH release, while secretion persists even at vanishingly low extracellular Ca2+ concentrations ([104], with kind permission of the publisher).

3. MAGNESIUM HOMEOSTASIS

The dietary magnesium content normally ranges from 6–15 mmol/d (140–360 mg), of which 30–40% is absorbed, mainly in the jejunum and ileum. Intestinal magnesium absorptive efficiency is stimulated by calcitriol (1,25(OH)2D), and can reach 70% during magnesium deprivation. Urinary magnesium excretion normally matches net intestinal absorption and is approximately 4 mmol/d (100 mg/d). Regulation of serum magnesium concentrations is achieved mainly by control of renal magnesium reabsorption. Only 20% of filtered magnesium is reabsorbed in the proximal tubule, about 60% is reabsorbed in the thick ascending loop of Henle, and 5–10% is reabsorbed in the distal convoluted tubule. Many factors, both hormonal and nonhormonal (e.g. parathyroid hormone, insulin, magnesium restriction, acid–base changes, and potassium depletion), influence both loop of Henle and distal tubule reabsorption. The major regulator is the plasma Mg2+ itself. Hypermagnesemia inhibits loop transport while hypomagnesemia stimulates transport whether there is magnesium depletion or not. The mechanism appears to be regulated by the Ca2+Mg2+-sensing receptor, located on the capillary side of the thick ascending loop cells, which senses the changes in Mg2+. Other factors that could influence reabsorption are hypercalcemia and the rate of sodium chloride reabsorption [103, 105]. During magnesium deficiency, the initial step will be a rapid decrease in the serum Mg2+ concentration. This leads to a reduction of magnesium excretion in the urine. It takes several weeks for equilibration with the bone stores to take place. 4. CURRENT ASSAYS FOR THE MEASUREMENT OF MAGNESIUM

a. Total Magnesium Serum is preferred over plasma for magnesium determination because anticoagulant interferes with most procedures. Atomic absorption spectrophotometry (AAS) is the method of choice and the reference method for the determination of total magnesium. After deproteinization and removal of phosphate ions with a lanthanum salt, the diluted filtrate is aspirated into an air–acetylene flame, where magnesium ions are atomized and detected via a magnesium hollow cathode lamp. Absorption at 285.2 nm is directly proportional to the number of magnesium atoms in the flame. Despite its excellent accuracy and precision, AAS is not routinely used in clinical laboratories because it is time-consuming and requires careful maintenance, calibration, and sample preparation. Most clinical laboratories now routinely use photometric methods for magnesium determination on automated analyzers. These methods employ dyes that change color upon selective binding

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Chapter 29 Measurement of Calcium, Phosphate and Magnesium Table VIII. Methods Commonly Employed for the Determination of Total Magnesium in Serum in Routine Laboratories Producer, Analyzer

Method

Abbott Architect® Bayer ADVIA® Beckman Coulter LX® DadeBehring Dimension® Olympus AU® Ortho-Clinical Diagnostics, Vitros® Roche Hitachi®, Modular®

Arsenazo complex Xylidyl blue1 Calmagite2 Methylthymol blue3 Xylidyl blue1 Formazan dye4 Xylidyl blue1

1Xylidyl

blue, a diazonium salt, forms in alkaline solution a purple complex with magnesium. The concentration of magnesium is determined by decrease of the extinction of xylidyl blue [106]. 2Calmagite forms a colored complex with magnesium in alkaline solution. This complex is stable over 30 minutes, and its absorbance at 520 nm is directly proportional to the magnesium concentration in the specimen. 3Methylthymol blue forms a blue complex with magnesium. Calcium interference is minimized by forming a complex between calcium and Ba-EGTA (chelating agent). 4Formazan dye: colorimetric assay using a formazan dye.

of magnesium from the sample [2]. Table VIII gives the methods commonly employed for the determination of magnesium in routine laboratories. b. Reference Range The reference range for children and adults is given in Table IX. The reference range for normal total serum magnesium is a subject of ongoing debate. Concentrations of 0.70–1.10 mmol/L are generally accepted. However, epidemiological studies have revealed correlations between increased cardiovascular risk and low serum magnesium concentrations within this reference interval [109]. Similar results have recently been obtained in a prospective study of patients with advanced atherosclerosis [110]. Therefore we regard values of at least 0.80 mmol/L as desirable. Toxicity: clinical symptoms appear at a concentration

Table IX. Age 1–30 d 31–365 d 1–3 y 4–6 y 7–9 y 10–12 y 13–15 y 16–18 y > 18 y

of 6.08 mg/dl (2.5 mmol/L), and respiratory muscle paralysis occurs at concentrations of about 12.2 mg/dl (5 mmol/L) [103]. c. Ionized Magnesium Today, ionized magnesium can be measured with the newly introduced magnesium ion-selective electrodes (ISEs) developed by AVL Medical Instruments (Schaffhausen, Switzerland), Kone Instruments (Oulu, Finland), and Nova Biomedical (Waltham, MA, USA) and incorporated in their commercial clinical analyzers [111]. However, in a study comparing the ionized magnesium results with these three analyzers, significant differences have been observed [111]. This limits a comparison of results in clinical practice. Recently, the determination of ionized magnesium in erythrocytes seemed to show more patients with hypomagnesemia than the determination of total serum magnesium or of ionized serum magnesium [112]. The reference interval for ionized magnesium depends on the analyzer used for its measurement and is on average 0.44–0.60 mmol/L [2].

B. Urinary Magnesium Hypomagnesemia develops late in the course of magnesium deficiency and intracellular magnesium depletion may be present despite normal serum magnesium levels. Due to the kidney’s ability to sensitively adapt its magnesium transport rate to imminent deficiency, urinary magnesium excretion is of special importance in the assessment of the magnesium status. In patients with hypomagnesemia, urinary magnesium excretion levels help to differentiate between renal magnesium wasting and extrarenal losses. In the presence of hypomagnesemia, the 24-h magnesium excretion is expected to be below 1 mmol [113].

Reference Range for Magnesium in Serum and Plasma in Children [107] and Adults [108]

Male (n) 68 62 140 120 118 111 84 73

Magnesium, relative molecular mass: 24.31. Conversion factors: mmol/L × 2.431 = mg/dl mg/dl × 0.4114 = mmol/L.

Male mg/dl 1.7–2.4 1.6–2.5 1.7–2.4 1.7–2.4 1.7–2.3 1.6–2.2 1.6–2.3 1.5–2.2 1.8–2.6

Male mmol/L 0.70–0.99 0.66–1.03 0.70–0.99 0.70–0.99 0.70–0.95 0.66–0.91 0.66–0.95 0.62–0.91 0.73–1.06

Female (n) 18 37 103 63 102 74 113 92

Female mg/dl 1.7–2.5 1.9–2.4 1.7–2.4 1.7–2.2 1.6–2.3 1.6–2.2 1.6–2.3 1.5–2.2 1.9–2.5

Female mmol/L 0.70–1.03 0.78–0.99 0.70–0.99 0.70–0.91 0.66–0.95 0.66–0.91 0.66–0.95 0.62–0.91 0.77–1.03

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Table X. Causes of Hypermagnesemia and Hypomagnesemia [modified from 105, 123; for additions references are given] Hypermagnesemia

Hypomagnesemia

Excessive magnesium intake

Impaired intake or intestinal absorption Malnutrition Malabsorption syndromes Vitamin D deficiency Primary infantile hypomagnesemia Increased intestinal losses Prolonged vomiting, diarrhea Intestinal drainage, fistula Kidney: Impaired tubular reabsorption Genetic Mg wasting syndromes Gitelman’s syndrome Bartter’s syndrome Paracellin-1 mutations Autosomal dominant, with low bone mass Tubulointerstitial disease Acute tubular necrosis, postobstruction Renal transplantation Drugs and toxins acting on the kidney Ethanol (metabolic acidosis, see also below) Diuretics (loop, thiazide, osmotic) Cisplatin Pentamidine, foscarnet Cyclosporine Aminoglycosides, amphotericin B Endocrine, ECF volume expansion Diabetes mellitus Hypercalcemia Metabolic acidosis (starvation, ketoacidosis) Redistribution, rapid shifts from extracellular fluid Recovery from diabetic ketoacidosis Refeeding syndrome Correction of respiratory acidosis Increased bone formation after PTX Treatment of vitamin D deficiency Other: Citrate in blood transfusion§§ [125] Pregnancy (3rd trimester) and lactation Preeclampsia (lower than normal pregnancy) [126] Pancreatitis, burns, excessive sweating

Cathartics (Mg-containing enemas) Excessive intake of antacids Intestinal obstruction following Mg ingestion Parenteral Mg administration Magnesium-rich urologic irrigants Tocolytic therapy with Mg-sulfate [124] Kidney: Impaired Mg excretion Renal failure Familial hypocalciuric hypercalcemia

Endocrine Adrenal insufficiency Hypothyroidism

Redistribution, rapid Mg mobilization from soft tissues Trauma Extensive burns Shock Sepsis Post cardiac arrest

Other: Hypothermia Erroneous: Prolonged venous occlusion (tourniquet)§ Hemolysis (high intracellular Mg content)

§Prolonged

venous occlusion leads to falsely elevated levels since about 30% of the magnesium in blood is bound to protein. §§Citrate is also used in apheresis procedures, e.g. platelet apheresis.

Magnesium/creatinine ratios and fractional excretion of magnesium have also been advocated as indicators of evolving magnesium deficiency in normomagnesemic individuals [114, 115]. However, the interpretation of results seems to be limited due to intra- and interindividual variability [116, 117]. The parenteral magnesium loading test (MLT) remains the gold standard for the evaluation of body magnesium status [118, 119]. The major limitation is a decreased renal function. The aim is to calculate the retention of an intravenous magnesium load. Several protocols have been proposed. In one protocol, 1 mmol of elemental magnesium per kg body weight is infused over 4 h and a 24-h urine collection is started simultaneously [120]. A shortterm MLT with 0.1 mmol/kg body weight of magnesium aspartate given over 1 h yielded comparable results and might represent a useful alternative [121]. Excretion of less than 70% of the infused magnesium is considered indicative of functional magnesium deficiency [119]. Besides its role for the identification of magnesium deficiency in normomagnesemic individuals with normal renal magnesium handling, the parenteral MLT allows differentiation between renal and extrarenal magnesium losses in patients with inherited forms of magnesium wasting, which are overtly hypomagnesemic [122]. A summary of the causes of hypermagnesemia and hypomagnesemia is given in Table X.

References 1. Thomas, L. (2005). Calcium (Ca). In “Labor und Diagnose” (L. Thomas, ed.), pp. 333–347. TH-Books, Verlagsgesellschaft mbH, Frankfurt/Main. 2. Henry, J. B. (2001). Analytical techniques. In “Clinical Diagnosis and Management by Laboratory Methods” (J. B. Henry, ed.), 20th ed. 194–198. W.B. Saunders Company, Philadelphia, PA. 3. Wandrup, J. (1989). Critical analytical and clinical aspects of ionized calcium in neonates. Clin. Chem. 35, 2027–2033. 4. Cali, J. P., Bowers, G. N. Jr., and Young, D. S. (1973). A referee method for the determination of total calcium in serum. Clin. Chem. 19, 1208–1213. 5. Welch, M. W., Hamar, D. W., and Fettman, M. J. (1990). Method comparison for calcium determination by flame atomic absorption spectrophotometry in the presence of phosphate. Clin. Chem. 36, 351–354. 6. Külpmann, W. R., Buchholz, R., Dyrssen, C., and Ruschke, D. (1989). A comparison of reference method values for calcium, lithium and magnesium with method-dependent assigned values. J. Clin. Chem. Clin. Biochem. 27, 631–637. 7. Corns, C. M., and Ludman, C. J. (1987). Some observations on the nature of the calcium-cresolphthalein complexone reaction and its relevance to the clinical laboratory. Ann. Clin. Biochem. 24, 345–351. 8. Janssen, J. W., and Helbing, A. R. (1991). Arsenazo III: An improvement of the routine calcium determination in serum. Eur. J. Clin. Chem. Clin. Biochem. 34, 197–201.

Chapter 29 Measurement of Calcium, Phosphate and Magnesium 9. Walmsley, T. A., and Fowler, R. T. (1981). Optimum use of 8-hydroxyquinoline in plasma calcium determinations. Clin. Chem. 27, 1782. 10. Corns, C. M. (1990). Interference by haemoglobin with the cresolphthalein complexone method for serum calcium measurement. Ann. Clin. Biochem. 27, 152–155. 11. Proctor, K. A. S., Rao, L. V., and Roberts, W. L. (2004). Gadolinium magnetic resonance contrast agents produce analytic interference in multiple serum assays. Am. J. Clin. Pathol. 121, 282–292. 12. Kefalas, C. H., Murray, N. G. B., Aguanno, J. J., Dockery, W. D., Weinstein, J. S., Anderson, K., and Klintmalm, G. B. (2004). Pseudohypocalcemia after magnetic resonance imaging with gadolinium in patients with cirrhosis. Liver Transplantation 10, 136–140. 13. Kang, H. P., Scott, M. G., Joe, B. N., Narra, V., Heiken, J., and Parvin, C. A. (2004). Model for predicting the impact of gadolinium on plasma calcium measured by the o-cresolphthalein method. Clin. Chem. 50, 741–746. 14. Prince, M. R., Erel, H. E., Lent, R. W., Blumenfeld, J., Kent, K. C., Bush, H. L., and Wang, Y. (2003). Gadodiamide administration causes spurious hypocalcemia. Radiology 227, 639–646. 15. Normann, P. T., Froysa, A., and Svaland, M. (1995). Interference of gadodiamide injection (OMNISCAN®) on the colorimetric determination of serum calcium. Scand. J. Clin. Lab. Invest. 55, 421–426. 16. John, R., Oleesky, D., Issa, B., Scanlon, M. F., Williams, C. P., Harrison, C. B., and Child, D. F. (1997). Pseudohypercalcaemia in two patients with IgM paraproteinaemia. Ann. Clin. Biochem. 34, 694–696. 17. Gawosky, J. M., and Walsh, D. (1989). Citrate interference in assays of total calcium in serum. Clin. Chem. 35, 2140–2141. 18. Kayamori, Y., and Katayama, Y. (1994). Enzymatic method for assaying calcium in serum and urine with porcine pancreatic α-amylase. Clin. Chem. 40, 781–784. 19. Morishita, Y., Iinuma, Y., Nakashima, N., Kadota, A., Miike, A., and Tadano, T. (1999). Enzymatic assay of calcium in serum with phospholipase D. Clin. Chem. 45, 2280–2283. 20. Buckley, B. M., and Russell, L. J. (1988). The measurement of ionised calcium in blood plasma. Ann. Clin. Biochem. 25, 447–465. 21. Vanstapel, F. J., and Lissens, W. D. (1984). Free ionised calcium – a critical survey. Ann. Clin. Biochem. 21, 339–351. 22. Bowers, G. N. Jr., Brassard, C., and Sena, S. F. (1986). Measurement of ionized calcium in serum with ion-selective electrodes: A mature technology that can meet the daily service needs. Clin. Chem. 32, 1437–1447. 23. Wandrup, J., and Kvetny, J. (1985). Potentiometric measurements of ionized calcium in anaerobic whole blood, plasma, and serum evaluated. Clin. Chem. 31, 856–860. 24. Gowans, E. M. S., and Fraser, C. G. (1986). Five methods for determining urinary calcium compared. Clin. Chem. 32, 1560–1562. 25. Husdan, H., Rapoport, A., Locke, S., and Oreopoulos, D. (1974). Effect of venous occlusion of the arm on the concentration of calcium in serum, and methods for its compensation. Clin. Chem. 20, 529–532. 26. Husdan, H., Rapoport, A., and Locke, S. (1973). Influence of posture on the serum concentration of calcium. Metabolism 22, 788–797. 27. Parfitt, A. M. (1969). Chlorothiazide-induced hypercalcemia in juvenile osteoporosis and hyperparathyroidism. New Engl. J. Med. 281, 55–59. 28. Payne, R. B., Little, A. J., Williams, R. B., and Milner, J. R. (1973). Interpretation of serum calcium in patients with abnormal serum proteins. Brit. Med. J. IV, 643–646. 29. Pedersen, K. O. (1972). On the cause and degree of intraindividual serum calcium variability. Scand. J. Clin. Lab. Invest. 30, 191–199. 30. Kaiser, C., Müller, H. A. G., Schmitt, Y., Geisen, H. P., and Schott, F. J. (1994). Should reference values for calcium be revised? Klin. Lab. 40, 599–602. 31. Herfarth, K., Drechsler, S., Imhoff, W., Schlander, M., Engelbach, M., Maier, A., and Schmidt-Gayk, H. (1992). Calcium regulating hormones after oral and intravenous calcium administration. Eur. J. Clin. Chem. Clin. Biochem. 30, 815–822.

503

32. Ljunghall, S., Hedstrand, H., Hellsing, K., and Wibell, L. (1977). Calcium, phosphate and albumin in serum. Acta Med. Scand. 201, 23–30. 33. Ladenson, J. H., Lewis, J. W., McDonald, J. M., Slatopolsky, E., and Boyd, J. C. (1978). Relationship of free and total calcium in hypercalcemic conditions. J. Clin. Endocrinol. Metab. 48, 393–397. 34. Edmondson, J. W., and Li, T. K. (1976). The relationship of serum ionized and total calcium in primary hyperparathyroidism. J. Lab. Clin. Med. 87, 624–629. 35. Pearce, C. J., Hine, T. J., and Peek, K. (1991). Hypercalcaemia due to calcium binding by a polymeric IgA κ-paraprotein. Ann. Clin. Biochem. 28, 229–234. 36. Lindgärde, F., and Zettervall, O. (1973). Hypercalcemia and normal ionized serum calcium in a case of myelomatosis. Ann. Intern. Med. 78, 396–399. 37. Lyon, M. E., Guajardo, M., Laha, T., Malik, S., Henderson, P. J., and Kenny, M. A. (1995). Electrolyte balanced heparin may produce a bias in the measurement of ionized calcium concentration in specimens with abnormally low protein concentration. Clin. Chim. Acta 233, 105–113. 38. Mann, S. W., Buckley, B. M., Roberts, K. D., and Green, A. (1988). Changes in plasma ionised calcium concentration during pediatric cardiopulmonary bypass surgery. Ann. Clin. Biochem. 25, 226–227. 39. Kost, G. J., Jammal, M. A., Ward, R. E., and Safwat, A. M. (1986). Monitoring of ionized calcium during human hepatic transplantation. Am. J. Clin. Pathol. 86, 61–70. 40. Dickerson, R. N., Alexander, K. H., Minard, G., Croce, M. A., and Brown, R. O. (2004). Accuracy of methods to estimate ionized and “corrected“ serum calcium concentrations in critically ill multiple trauma patients receiving specialized nutrition support. J. Parenteral Enteral Nutr. 28, 133–141. 41. Godolphin, W., Cameron, E. C., Frolich, J., and Price, J. D. E. (1979). Spurious hypocalcemia in hemodialysis patients after heparinization. In-vitro formation of calcium soaps. Am. J. Clin. Pathol. 71, 215–218. 42. Schmidt-Gayk, H. (1992). Urinary calcium. Klin. Lab. 38, 614–615. 43. Coe, F. L., Parks, J. H., and Moore, E. S. (1979). Familial idiopathic hypercalciuria. New Engl. J. Med. 300, 337–340. 44. Dörner, K. (1989). In (S. Meites, ed.) Pediatric Clinical Chemistry, Reference (Normal Values). p. 86. AACC Press, Washington, DC. 45. Paunier, L., Borgeaud, M., and Wyss, M. (1970). Urinary excretion of magnesium and calcium in normal children. Helv. Paediat. Acta 25, 577–584. 46. Misselwitz, J., and Kauf, E. (1981). Die Kalziumausscheidung im Urin bei gesunden Kindern als Grundlage für die Suche nach einer Hyperkalziurie bei Kalziumnephrolithiasis. Kinderärztliche Praxis 49, 64–71. 47. Hauptmann, C., Priem, F., Glatzel, E., and Schneider, M. (1992). Reference values for the excretion of calcium (Ca), magnesium (Mg), phosphorus (P) and creatinine in children. Klin. Lab. 38, 211–216. 48. Massey, L. K., and Whiting, S. J. (1996). Dietary salt, urinary calcium, and bone loss. J. Bone Min. Res. 11, 731–736. 49. Hess, B., Ackermann, D., Essig, M., Takkinen, R., and Jaeger, P. (1995). Renal mass and serum calcitriol in male idiopathic calcium renal stone formers: Role of protein intake. J. Clin. Endocrinol. Metab. 80, 1916–1921. 50. Hess, B., Hasler-Strub, U., Ackermann, D., and Jaeger, P. (1997). Metabolic evaluation of patients with recurrent idiopathic calcium nephrolithiasis. Nephrol. Dial. Transplant. 12, 1362–1368. 51. Min, H. K., Jones, J. E., and Flink, E. B. (1966). Circadian variations in renal excretion of magnesium, calcium, phosphorus, sodium, and potassium during frequent feeding and fasting. Fed. Proc. 25, 917–921. 52. Wisser, H., Doerr, P., Stamm, D., Fatranska, M., Giedke, H., and Wever, R. (1973). Circadian rhythm of electrolytes, catecholamines and 17-hydroxycorticosteroids in urine. Klin. Wschr. 51, 242–246.

504 53. Robertson, W. G., Gallagher, J. C., Marshall, D. H., Peacock, M., and Nordin, B. E. C. (1974). Seasonal variations in urinary excretion of calcium. Brit. Med. J. IV, 436–437. 54. Nordin, B. E. C. (ed.) (1976). Calcium, Phosphate and Magnesium Metabolism. Clinical Physiology and Diagnostic Procedures. Churchill Livingstone, Edinburgh and New York. 55. Meyer-Lüerssen, B., Traber, L., Knörzer, T., and Schmidt-Gayk, H. (2000). Bone resorption markers in pre- and postmenopausal females. Clin. Lab. 46, 285–290. 56. Crook, M., and Swaminathan, R. (1996). Disorders of plasma phosphate and indications for its measurement. Ann. Clin. Biochem. 33, 376–396. 57. Schiavi, S. C., and Kumar, R. (2004). The phosphatonin pathway: New insights in phosphate homeostasis. Kidney Int. 65, 1–14. 58. Cherian, A. G., and Hill, J. G. (1978). Percentile estimates of reference values for fourteen chemical constituents in sera of children and adolescents. Am. J. Clin. Pathol. 69, 24–31. 59. Soldin, S. J., Hicks, J. M., Bailey, J., Cook, J. F., and Beatey, J. (1997). Pediatric reference ranges for phosphate on the Hitachi 747 analyzer. Clin. Chem. 43, No. 6, S198 (Abstract). 60. Greenberg, B. G., Winters, R. W., and Graham, J. B. (1960). The normal range of serum inorganic phosphorus and its utility as a discriminant in the diagnosis of congenital hypophosphatemia. J. Clin. Endocrinol. Metab. 20, 364–379. 61. Weisinger, J. R., and Bellorin-Font, E. (1998). Magnesium and phosphorus. Lancet 352, 391–396. 62. Loghman-Adham, M., Walton, D., Iverius, P.-H., Deiss, A., Knight, J. A., and Cheung, A. K. (1997). Spurious hypophosphatemia in a patient with multiple myeloma. Am. J. Kidney Dis. 30, 571–575. 63. Tokmakjian, S., Moses, G., and Haines, M. (1990). Excessive sample blankings in two analyzers generate reports of apparent hypoglycemia and hypophosphatemia in patients with macroglobulinemia. Clin. Chem. 36, 1261–1262. 64. Duncanson, G. O., and Worth, H. G. J. (1990). Pseudohypophosphataemia as a result of bilirubin interference. Ann. Clin. Biochem. 27, 253–257. 65. Donhowe, J. M., Freier, E. F., Wong, E. T., and Steffes, M. W. (1981). Factitious hypophosphatemia related to mannitol therapy. Clin. Chem. 27, 1765–1769. 66. Keller, H. (ed.). (1991). Phosphat. In: “Klinisch-chemische Labordiagnostik für die Praxis”, pp. 217–218. Georg Thieme Verlag, Stuttgart, New York. 67. Ball, C. L., Tobler, K., Ross, B. C., Connors, M. R., and Lyon, M. E. (2004). Spurious hyperphosphatemia due to sample contamination with heparinized saline from an indwelling catheter. Clin. Chem. Lab. Med. 42, 107–108. 68. Takeda, E., Yamamoto, H., Nashiki, K., Sato, T., Arai, H., and Taketani, Y. (2004). Inorganic phosphate homeostasis and the role of dietary phosphorus. J. Cell. Mol. Med. 8, 191–200. 69. Savory, D. J., and Pearce, C. J. (1995). Paraprotein interference causing pseudohyperphosphataemia: evaluation of an improved methodology. Ann. Clin. Biochem. 32, 498–501. 70. Cohen, A. M., Magazanik, A., van-der Lijn, E., Shaked, P., and Levinsky, H. (1994). Pseudohyperphosphataemia incidence in an automatic analyzer. Eur. J. Clin. Chem. Clin. Biochem. 32, 559–561. 71. Sonnenblick, M., Eylath, U., Brisk, R., Eldad, C., and Hershko, C. (1986). Paraprotein interference with colorimetry of phosphate in serum of some patients with multiple myeloma. Clin. Chem. 32, 1537–1539. 72. Petterson, T., Hortling, L., Teppo, A. M., Tötterman, K. J., and Fyhrquist, F. (1987). Phosphate binding by a myeloma protein. Acta Med. Scand. 222, 89–91. 73. Bowles, S. A., Tait, R. C., Jefferson, S. G., Gilleece, M. H., and Haeney, M. R. (1994). Characteristics of monoclonal immunoglobulins that interfere with serum inorganic phosphate measurement. Ann. Clin. Biochem. 31, 249–254. 74. Landowne, R. A. (1979). Immunoglobulin interference with phosphorus and chloride determinations with the Coulter Chemistry. Clin. Chem. 25, 1189–1190.

HEINRICH SCHMIDT-GAYK 75. Weinberg, J., and Adler, A. J. (1989). Spurious hyperphosphatemia in patients with dysglobulinemia. Miner. Electrolyte Metab. 15, 185–186. 76. Bakker, A. J., Bosma, H., and Christen, P. J. (1990). Influence of monoclonal immunoglobulins in three different methods for inorganic phosphorus. Ann. Clin. Biochem. 27, 227–231. 77. Hawkins, R. C. W. (1991). Pseudohyperphosphataemia in multiple myeloma. Ann. Clin. Biochem. 28, 226–228. 78. Larner, A. J. (1994). Pseudohyperphosphataemia and monoclonal gammopathy. Brit. J. Clin. Pathol. 48, 272–274. 79. Zaman, Z., Sneyers, L., van Orshoven, A., Blanckaert, N., and Marien, G. (1995). Elimination of paraprotein interference in determination of plasma inorganic phosphate by ammonium molybdate method. Clin. Chem. 41, 609–614. 80. Tipper, P. L., and Miyada, D. S. (1986). Precipitation of multiple myeloma serum in the phosphorus channel of the Beckman Astra. Clin. Chem. 32, 1597. 81. Mandry, J. M., Posner, M. R., Tucci, J. R., and Eil, C. (1991). Hyperphosphatemia in multiple myeloma due to a phosphate-binding immunoglobulin. Cancer 68, 1092–1094. 82. Sinclair, D., Smith, H., and Woodhead, P. (2004). Spurious hyperphosphataemia caused by an IgA paraprotein: a topic revisited. Ann. Clin. Biochem. 41, 119–124. 83. McClure, D., Lai, L. C., and Cornell, C. (1992). Pseudohyperphosphataemia in patients with multiple myeloma. J. Clin. Pathol. 45, 731–732. 84. Marcu, C. B., and Hotchkiss, M. (2004). Pseudohyperphosphatemia in a patient with multiple myeloma. Connecticut Medicine 68, 71–72. 85. Zimmermann, K., Jenckel-Loß, G., Fiedler, H., and Weber, R. (1993). Evaluation of enzymatic phosphate determination in comparison with the molybdenum blue method. Klin. Lab. 39, 777–780. 86. Berti, G., Fossati, P., Tarenghi, G., and Musitelli, C. (1988). Enzymatic colorimetric method for the determination of inorganic phosphorus in serum and urine. J. Clin. Chem. Clin. Biochem. 26, 399–404. 87. de Castro, M. D. L., Quiles, R., Fernández-Romero, J. M., and Fernández, E. (1995). Continuous-flow assay with immobilized enzymes for determining of inorganic phosphate in serum. Clin. Chem. 41, 99–102. 88. Ghetta, A., Matus-Ortega, M., Garcia-Mena, J., Dehò, G., Tortora, P., and Regonesi, M. E. (2004). Polynucleotide phosphorylasebased photometric assay for inorganic phosphate. Anal. Biochem. 37, 209–214. 89. Kemp, G. J., Blumsohn, A., and Morris, B. W. (1992). Circadian changes in plasma phosphate concentration, urinary phosphate excretion, and cellular phosphate shifts. Clin. Chem. 38, 400–402. 90. Kemp, G. J., Blumsohn, A., and Morris, B. W. (1992). Cellular phosphate shifts during oral glucose loading. Clin. Chem. 38, 2338–2339. 91. Markowitz, M., Rotkin, L., Rosen, J. F. (1981). Circadian rhythms of blood minerals in humans. Science 213, 672–674. 92. Fraser, W. D., Logue, F. C., Christie, J. P., Cameron, D. A., O’Reilly, D. St. J., and Beastall, G. H. (1994). Alteration of the circadian rhythm of intact parathyroid hormone following a 96-hour fast. Clin. Endocrinol. 40, 523–528. 93. Ring, T., Sanden, A. K., Hansen, H. H. T., Halkier, P., Nielsen, C., and Fog, L. (1995). Ultradian variation in serum phosphate concentration in patients on haemodialysis. Nephrol. Dial. Transplant. 10, 59–63. 94. Ono, T., Kitaguchi, K., Takehara, M., Shiiba, M., and Hayami, K. (1981). Serum-constituents analyses: effect of duration and temperature of storage of clotted blood. Clin. Chem. 27, 35–38. 95. Ladenson, J. H., Tsai, L. M. B., Michael, J. M., Kessler, G., and Joist, J. H. (1974). Serum versus heparinized plasma for eighteen common chemistry tests. Is serum the appropriate specimen? Am. J. Clin. Pathol. 62, 545–552. 96. Lutomski, D. M., and Bower, R. H. (1994). The effect of thrombocytosis on serum potassium and phosphorus concentrations. Am. J. Med. Sci. 307, 255–258.

Chapter 29 Measurement of Calcium, Phosphate and Magnesium 97. Walton, R. J., and Bijvoet, O. L. M. (1975). Nomogram for derivation of renal threshold phosphate concentration. Lancet II, 309–310. 98. Palmer, F. J., Nelson, J. C., and Bacchus, H. (1974). The chloridephosphate ratio in hypercalcemia. Ann. Intern. Med. 80, 200–204. 99. Min, H. K., Jones, J. E., and Flink, E. B. (1966). Circadian variations in renal excretion of magnesium, calcium, phosphorus, sodium, and potassium during frequent feeding and fasting. Fed. Proc. 25, 917–921. 100. Aiken, C. G. A., Sherwood, R. A., and Lenney, W. (1993). Role of plasma phosphate measurements in detecting rickets of prematurity and in monitoring treatment. Ann. Clin. Biochem. 30, 469–475. 101. Soloni, F. G., Bandin, R., and Amazon, K. (1986). Influence of sample processing on urine phosphorus results with the Ektachem® 400 and 700 analyzers. Clin. Chem. 32, 1594–1595. 102. Tovay, J. A., Shyam Sundar, A., Ikram, S., Smith, S., Penny, W. J. (1992). Human cardiac muscle magnesium and potassium concentrations: can skeletal muscle, mononuclear blood cells, erythrocyte and plasma concentrations provide a surrogate measure? Ann. Clin. Biochem. 29, 461–462. 103. Dörner, K. (1998). Magnesium. In “Clinical Laboratory Diagnostics” (L. Thomas, ed.), pp. 339–340. TH-Books, Frankfurt/Main, Germany. 104. Brown, E. M., and Chen, C. J. (1989). Calcium, magnesium and the control of PTH secretion. Bone Mineral 5, 249–257. 105. Bringhurst, F. R., Demay, M. B., Krane, S. M., and Kronenberg, H. M. (2005). Bone and mineral metabolism in health and disease. Magnesium metabolism. In: “Harrison’s Principles of Internal Medicine”, D. L., Kasper, et al., 16th edition, pp. 2244–2245. McGraw-Hill, New York. 106. Mann, C. K., and Yoe, J. H. (1956). Spectrophotometric determination of magnesium with sodium 1-azo-2-hydroxy-3-(2,4-dimethylcarboxanilido)-naphthalene-1′-(2-hydroxy-benzene-5-sulfonate). Anal. Chem. 28, 202–205. 107. Hicks, J. M., Bailey, J., Bjorn, S., et al. (1995). Pediatric reference ranges for plasma magnesium. Clin. Chem. 41, S93 (abstract). 108. Speich, M., Bousquet, B., and Nicolas, G. (1981). Reference values for ionized, complexed, and protein-bound plasma magnesium in men and women. Clin. Chem. 27, 246–248. 109. Tsuji, H., Venditti, F. J. Jr., Evans, J. C., Larson, M. G., and Levy, D. (1994). The association of levels of serum potassium and magnesium with ventricular premature complexes (the Framingham Heart Study). Am. J. Cardiol. 74, 232–235. 110. Amighi, J., Sabeti, S., Schlager, O., Mlekusch, W., Exner, M., Lalouschek, W., Ahmadi, R., Minar, E., and Schillinger, M. (2004). Low serum magnesium predicts neurological events in patients with advanced atherosclerosis. Stroke 35, 22–27. 111. Huijgen, H. J., Sanders, R., Cecco, S. A., Rehak, N. N., Sanders, G. T., and Elin, R. J. (1999). Serum ionized magnesium: comparison of

112.

113. 114.

115.

116. 117.

118. 119.

120.

121.

122.

123. 124.

125.

126.

505 results obtained with three ion-selective analyzers. Clin. Chem. Lab. Med. 37, 465–470. Malon, A., Brockmann, C., Fijalkowska-Morawska, J., Rob, P., and Maj-Zurawska, M. (2004). Ionized magnesium in erythrocytes – the best magnesium parameter to observe hypo- or hypermagnesemia. Clin. Chim. Acta 349, 67–73. Sutton, R. A., and Domrongkitchaiporn, S. (1993). Abnormal renal magnesium handling. Miner. Electrolyte Metab. 19, 232–240. Elisaf, M., Panteli, K., Theodorou, J., and Siamopoulos, K. C. (1997). Fractional excretion of magnesium in normal subjects and in patients with hypomagnesemia. Magnes. Res. 10, 315–320. Tang, N. L., Cran, Y. K., Hui, E., and Woo, J. (2000). Application of urine magnesium/creatinine ratio as an indicator for insufficient magnesium intake. Clin. Biochem. 33, 675–678. Nicoll, G. W., Struthers, A. D., and Fraser, C. G. (1991). Biological variation of urinary magnesium. Clin. Chem. 37, 1794–1795. Djurhuus, M. S., Gram, J., Petersen, P. H., Klitgaard, N. A., Bollerslev, J., and Beck-Nielsen, H. (1995). Biological variation of serum and urinary magnesium in apparently healthy males. Scand. J. Clin. Lab. Invest. 55, 549–558. Elin, R. J. (1994). Magnesium: the fifth but forgotten electrolyte. Am. J. Clin. Pathol. 102, 616–622. Hebert, P., Mehta, N., Wang, J., Hindmarsh, T., Jones, G., and Cardinal, P. (1997). Functional magnesium deficiency in critically ill patients identified using a magnesium-loading test. Crit. Care Med. 25, 749–755. Ryzen, E., Elbaum, N., Singer, F. R., and Rude, K. (1985). Parenteral magnesium tolerance testing in the evaluation of magnesium deficiency. Magnesium 4, 137–147. Rob, P. M., Dick, K., Bley, N., Seyfert, T., Brinckmann, C., Hollriegel, V., Friedrich, H. J., Dibbelt, L., and Seelig, M. S. (1999). Can one really measure magnesium deficiency using the short-term magnesium loading test? J. Intern. Med. 246, 373–378. Schlingmann, K. P., Konrad, M., and Seyberth, H. W. (2004). Genetics of hereditary disorders of magnesium homeostasis. Pediatr. Nephrol. 19, 13–25. Rude, R. K. (1998). Magnesium deficiency: a cause of heterogenous disease in humans. J. Bone Miner. Res. 13, 749–758. Koontz, S. L., Friedman, S. A., and Schwartz, M. L. (2004). Symptomatic hypocalcemia after tocolytic therapy with magnesium sulfate and nifedipine. Am. J. Ostet. Gynecol. 190, 1773–1776. Inoue, S., Akazawa, S., Nakaigawa, Y., Shimizu, R., and Seo, N. (2004). Changes in plasma total and ionized magnesium concentrations and factors affecting magnesium concentrations during cardiac surgery. J. Anesth. 18, 216–219. Sukonpan, K., and Phupong, V. (2004). Serum calcium and serum magnesium in normal and preeclamptic pregnancy. Arch. Gynecol. Obstet. (Epub ahead of print).

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Chapter 30

Measurement of Parathyroid Hormone Harald Jüppner Ghada El-Hajj Fuleihan

I. II. III. IV.

Endocrine and Pediatric Nephrology Units, Massachusetts General Hospital and Harvard Medical School, Boston, MA, USA Calcium Metabolism and Osteoporosis Program, American University of Beirut-Medical Center, Beirut, Lebanon

V. Hyperparathyroidism in Renal Osteodystrophy VI. Pseudohypoparathyroidism (PHP) VII. Conclusion References

Abstract Background Different Immunometric Assays for the Detection of PTH Primary Hyperparathyroidism

I. ABSTRACT

and ntPTH(1-84)) with those obtained with newer, secondgeneration assays (detection of PTH(1-84) alone). While most immunometric PTH assay systems are appropriate for the diagnosis of primary hyperparathyroidism, it appears plausible that immunometric assays designed to detect only PTH(1-84) will be more useful for certain diagnostic studies, i.e. for monitoring PTH concentrations intraoperatively and for assessing the pulsatility of PTH secretion. In addition, the ability to distinguish between the relative concentrations of ntPTH(1-84) and PTH(1-84) may reveal previously unsuspected biological roles for ntPTH(1-84) fragments in patients with end-stage renal disease and/or other disorders involving PTH.

Biologically active parathyroid hormone (PTH) in humans with normal renal function circulates predominantly as an 84 amino acid peptide. However, large aminoterminally truncated PTH(1-84) fragments (ntPTH(1-84)) that are readily detectable with most first-generation immunometric PTH assays may also affect bone metabolism and calcium homeostasis. Some of these ntPTH(1-84) fragments exhibit an elution profile on high-performance liquid chromatography (HPLC), which is indistinguishable from that of synthetic PTH(7-84). This hormonal fragment was recently shown through in vitro experiments to be the predominant form of PTH secreted by the parathyroid glands. Furthermore, it appears to have hypocalcemic properties in vivo and to inhibit osteoclastic bone resorption and the formation of mature osteoclasts in vitro. At present, it is uncertain whether important differences in disease states can be revealed by comparing results obtained with firstgeneration immunometric assays (detection of PTH(1-84) Dynamics of Bone and Cartilage Metabolism

II. BACKGROUND Parathyroid hormone secretion, fragments and action: PTH(1-84), an important regulator of mineral ion homeostasis. It mediates its actions through the PTH/PTHrP receptor, 507

Copyright © 2006 by Academic Press. All rights of reproduction in any form reserved.

508

HARALD JÜPPNER AND GHADA EL-HAJJ FULEIHAN

Figure 1

Two distinct receptors appear to mediate the actions of parathyroid hormone (PTH). The PTH/PTHrP receptor mediates most of the known biological actions of PTH in target tissues by initiating intracellular signal transduction through protein kinase A and protein kinase C. This receptor is activated with similar efficacy and efficiency by intact PTH(1-84), by PTH(1-34), and by the parathyroid hormone-related peptide (PTHrP). At least in humans, PTH(1-84) and PTH(1-34) activate also a second receptor, the PTH2-receptor. In contrast to these G protein-coupled receptors, the putative C-PTH receptor interacts with portions of PTH(1-84) that are located further toward the carboxy-terminus; the signal transduction pathways for this receptor have yet to be determined.

a G protein-coupled receptor that is abundantly expressed in kidney and bone (Fig. 1) [1]. PTH(1-84) is cleaved within the parathyroid glands and in peripheral tissues, such as liver, into different carboxyl-terminal fragments, most of which are considered to be biologically inactive and are eliminated mainly by glomerular filtration and subsequent tubular degradation [2]. Larger aminoterminally truncated forms of PTH(1-84) (ntPTH(1-84)) are also generated by the parathyroid glands and recent studies have suggested that the predominant form of ntPTH(1-84) is PTH(7-84) [2a]. PTH(1-84) interacts not only with the PTH/PTHrP receptor, but also with a distinct, yet uncloned receptor, referred to as the C-PTH receptor [3]. At least in humans, PTH(1-84) mediates its actions furthermore through another G protein-coupled receptor, the PTH2-receptor, which serves primarily, however, as a receptor for the tuberoinfundibular peptide of 39 residues (TIP39) [4, 5]. Circulating ntPTH(1-84) fragments, particularly PTH(7-84), have gained considerable interest, since this latter peptide was recently shown to have hypocalcemic properties when tested in vivo [6, 7]. PTH(1-34) or PTH(1-84) given parentally rapidly corrects the hypocalcemia induced by parathyroidectomy. These calcemic actions of either PTH molecule are mediated through the PTH/PTHrP receptor. However, although synthetic PTH(7-84) is unable to efficiently antagonize the actions of PTH(1-34) at this G protein-coupled receptor in vitro [8, 9], the peptide efficiently and dose-dependently reduced

the calcemic response to PTH(1-84) and PTH(1-34) [6, 7]. Hence, PTH(7-84) and potentially other large ntPTH(1-84) fragments detected by the first generation immunometric PTH assays may have biological activities in vivo that are not mediated through the PTH/PTHrP receptor, but instead through the C-PTH receptor [2, 3, 8, 9]. C-PTH receptors were detected in several different tissues and in clonal cell lines [10-12], including ROS 17/2.8 cells [3, 13]; the most abundant concentrations of this receptor were detected on osteocytic cells [8]. Using in vitro bioassays such as calvarial resorption assays, synthetic PTH(7-84), but not PTH(3-34) or PTHrP(7-36), was shown to inhibit basal 45Ca-release from pre-labeled calvarial bones [9]. Furthermore, activation of bone resorption and the formation of TRAP+ multinucleated cells through a variety of osteotropic agents such 1,25(OH)2D3, prostaglandin E2, and IL-11, was inhibited by PTH(7-84), but not by PTH(3-34) or PTHrP(7-36). Consistent with these findings, Langub et al. showed that PTH(7-84) decreases PTH-stimulated bone turnover in rats with moderate renal failure [14]. Since the predominant form of ntPTH(1-84) fragments [15–20] secreted by the parathyroid glands appears to be PTH(7-84) [2a], naturally occurring hormonal fragments could display hypocalcemic properties in vivo by affecting the activity and/or maturation of bone-resorbing osteoclasts. The measurement of intact PTH(1-84) without interference by PTH(7-84) and other ntPTH(1-84) fragments, and vice versa, may therefore be of considerable diagnostic importance.

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Chapter 30 Measurement of Parathyroid Hormone

III. DIFFERENT IMMUNOMETRIC ASSAYS FOR THE DETECTION OF PTH PTH was first measured by radioimmunoassays using polyclonal antibodies directed against epitopes that proved to be located mostly within the mid- or carboxyl-terminal portion of the PTH molecule [21, 22] (Fig. 2). Such assays, which detected predominantly carboxyl-terminal PTH fragments that do not activate the PTH/PTHrP receptor, were therefore an unreliable predictor of biologically active PTH in the circulation. To detect intact PTH(1-84) rather than hormonal fragments, radioimmunoassays have now been largely replaced by immunometric assays [23]. Immunometric assays utilize two different affinity purified antibodies that are directed against distinct epitopes within the PTH molecule [23–26]. One of these antibodies, directed against a region within the carboxyl-terminus, is usually immobilized onto a plastic surface and captures the circulating intact PTH (as well as some PTH fragments). The second antibody, directed against an amino-terminal epitope, is either radiolabeled, biotinylated, or covalently bound to an enzyme, thus readily allowing its quantification. Due to the use of two antibodies that recognize distinct epitopes and due to amplification of the detection signal, immunometric assays are more sensitive and specific than displacement-type assays [27], and they are less influenced by nonspecific serum effects; the normal range for first generation assays is usually about 10-65 pg/ml [23]. However, most first-generation immunometric assays detect not only the intact PTH(1-84), but also ntPTH(1-84)

Figure 2

fragments, which co-elute from reverse-phase HPLC columns with synthetic PTH(7-84) [15–18, 28]. In fact, it was determined that the detection antibodies used in most first-generation immunometric assays are directed against epitopes that do not include the first six amino acids and, consequently, such assays detect PTH(1-84) and PTH(7-84) equivalently [6, 20, 29–31]. In contrast, several more recently developed second-generation immunometric assays that use detection antibodies directed against epitopes within the first few amino-terminal amino acids of PTH detect the intact hormone, but show little or no crossreactivity with PTH(7-84) [6, 20, 29, 30, 32]. Consistent with these assay characteristics, second-generation immunometric PTH assays detect mostly the intact hormone in HPLC-fractionated serum/plasma samples from patients with primary and secondary hyperparathyroidism, but not PTH(7-84) and other forms of ntPTH(1-84) [15–18, 20, 28, 31]. Second-generation PTH immunometric assays therefore have the advantage of detecting almost exclusively the intact PTH(1-84) without interference by ntPTH(1-84) fragments [6, 18, 20, 29, 30, 32], but it remains uncertain whether this improvement in specificity increases their diagnostic utility. In healthy individuals, the circulating levels of PTH are 5-36 pg/ml, when measured with one of the secondgeneration immunometric assays that uses a detection antibody, which requires the presence of the first few amino acid of intact PTH(1-84) [20, 29, 33, 34]. Other secondgeneration PTH immunometric assays that use detection antibodies, which require the presence of the first three to

Schematic presentation of PTH(1-84) and the approximate location of the peptide regions recognized by antibodies used in first- and second-generation “two-site” immunometric PTH assays (modified from [25]). Both assays usually employ capture antibodies directed toward epitopes located within the carboxyl-terminal region of the peptide that are immobilized on a solid phase. Detection antibodies are targeted to epitopes located closer to the amino-terminal end of the peptide. In first-generation immunometric PTH assays, detection antibodies were directed to a region between amino acid residues 13 and 34. By contrast, detection antibodies in second generation immunometric PTH assays are directed toward epitopes in the most amino-terminal portion of the molecule. Synthetic PTH peptides that are truncated at their amino-terminal end such as PTH(7-84) are much less likely to be detected in second-generation assays, but commonly cross-react with the detection antibodies used in first-generation assays.

510 five amino acids of PTH(1-84) have similar normal ranges [30, 31]. Thus second-generation immunometric PTH assay detect only about half of the PTH concentration that is detected with first-generation assays [20, 29–31, 33, 34]. A “reverse sandwich” assay which utilizes regionspecific antibodies in reverse order, i.e. the antibody against an amino-terminal epitope for capture and an antibody against a mid-regional fragment of PTH(1-84) for detection [35], was also reported to lack cross-reactivity with human PTH(7-84) [36]. This indicates that the immobilized capture antibody used in this assay has a specificity similar to that of the detection antibody used in the recently described second-generation immunometric PTH assays [6, 20, 29–32].

IV. PRIMARY HYPERPARATHYROIDISM First-generation immunometric PTH assays provide excellent sensitivity and specificity for the diagnosis of even mild forms of primary hyperparathyroidism, and the concentrations of PTH measured by these assays are usually below the normal range in most patients with humoral hypercalcemia of malignancy and with different forms of hypoparathyroidism (20, 22, 23, 33). However, since these immunometric assays detect not only PTH(1-84), but also significant amounts of ntPTH(1-84), several studies were recently conducted to re-assess their clinical performance in comparison to second-generation assays. Direct comparison of first- and second-generation PTH assays revealed an excellent correlation (r = 0.92-0.95, p < 0.0001) between both assays for a large number of patients with primary hyperparathyroidism [20, 33, 37]. In one study, the sensitivity of a second-generation immunometric PTH assay for the diagnosis of primary hyperparathyroidism was estimated to be 94%, compared to 92% sensitivity of a first generation assay [20], while another study found increased PTH levels in 73% and 96% of the patients using first- and second-generation assays, respectively [33]. Additional studies are, however, required to determine whether second-generation immunometric PTH assays provide also an improved diagnostic sensitivity for patients with mild forms of primary hyperparathyroidism [38]. The rapid measurement of PTH levels is now frequently used intraoperatively to confirm that an adenoma has been successfully removed. This technique is used particularly during less invasive procedures involving limited neck exploration after preoperative localization of the putative adenoma. PTH levels in patients with primary hyperparathyroidism that were measured 5, 10, and 15 minutes after excision of the adenoma, revealed that PTH levels

HARALD JÜPPNER AND GHADA EL-HAJJ FULEIHAN

decreased slightly faster postexcision when measured with a second-generation assay. At 5 minutes, PTH levels had dropped to less than 50% of the level before parathyroidectomy in all patients using the second-generation assay, but levels remained above that cut-off in six (8%) of the patients using the first-generation assay [34]. It is therefore conceivable that rapid second-generation assays, which measure only PTH(1-84), will provide more useful intraoperative PTH measurements during parathyroidectomy than currently employed first-generation assays.

V. HYPERPARATHYROIDISM IN RENAL OSTEODYSTROPHY An excellent correlation between first- and secondgeneration PTH assays (r = 0.977; p < 0.0001) was observed for most patients with end-stage renal disease on maintenance dialysis [20, 29, 32]. Mean PTH levels in patients with end-stage renal disease were, when assessed by secondgeneration assays, about 50% lower than those assessed by first-generation assays [20, 29, 32, 39, 40]. Some patients with renal failure were shown to have an abnormal ratio of PTH(1-84) to ntPTH(1-84), which was thought to provide a better prediction of bone turnover than each measurement alone [39]. Although similar studies by others were thus far unable to confirm these findings [25, 30, 32, 40–42], it is conceivable that an independent assessment of intact PTH and ntPTH(1-84) fragments may predict the form of bone disease in patients with end-stage renal disease with higher accuracy than is currently possible [43, 44]. As for patients with primary hyperparathyroidism, plasma PTH levels decreased more rapidly after parathyroidectomy for secondary (or tertiary) hyperparathyroidism when measured with a second-generation assay than with a first-generation assay [34, 45].

VI. PSEUDOHYPOPARATHYROIDISM (PHP) PHP type I (PHP-Ia) is characterized, besides various developmental abnormalities, by resistance to several hormones that mediate their effects through G proteincoupled receptors. However, most prominent is the resistance to PTH in proximal renal tubules, which leads to hypocalcemia and hyperphosphatemia [46-48]. PHP-Ia is caused by inactivating mutations on the maternal allele affecting those exons of the GNAS locus that encode the stimulatory G protein. Similar resistance to PTH and consequently secondary hyperparathyroidism is also observed in patients with PHP type Ib (PHP-Ib), an imprinted disorder characterized by resistance to the actions of PTH

Chapter 30 Measurement of Parathyroid Hormone

in the proximal, but not the distal renal tubules; PHP-Ib appears to be caused in most patients by deletions up-stream of GNAS or within this gene locus [49–52]. It was recently determined that the ratio of ntPTH(1-84) to PTH(1-84) is higher in patients with PHP-Ia or PHP-Ib than in healthy controls (0.56 ± 0.07 vs. 0.30 ± 0.11; P < 0.01) [31], but it remains uncertain whether the apparent increase in the levels of PTH fragments contributes to the pathophysiology of PTH-resistant hypocalcemia in patients with PHP.

VII. CONCLUSION Because some ntPTH(1-84) fragments, particularly PTH(7-84), appear to have biological activity affecting calcium homeostasis, it will be important to further evaluate the clinical utility of first- and second-generation immunometric PTH assays in different groups of patients. It is conceivable that future fragment-specific assays will allow the selective quantification of PTH(7-84) and possibly other ntPTH(1-84) fragments, and that the separate measurement of PTH(1-84) and different PTH fragments will provide further insights into the regulation of mineral ion homeostasis in normal and pathophysiological conditions.

References 1. Jüppner, H., Gardella, T., Brown, E., Kronenberg, H., and Potts, J., Jr. (2000). Parathyroid hormone and parathyroid hormone-related peptide in the regulation of calcium homeostasis and bone development. In: “Endocrinology” (L. DeGroot, and J. Jameson, eds.), pp. 969–998, Philadelphia, PA: W.B. Saunders Company. 2. Murray, T. M., Rao, L. G., Divieti, P., and Bringhurst, F. R. (2005). Parathyroid hormone secretion and action: evidence for discrete receptors for the carboxyl-terminal region and related biological actions of carboxyl-terminal ligands. Endocr Rev 26, 78–113. 2a.D’Amour, P., Brossard, J., Rousseau, L., Nguyen-Yamamoto, L., Nassif, E., Lazure, C., Gauthier, D., Lavigne, J., and Zahradnik, R. (2005). Structure of non-(1-84) PTH fragments secreted by parathyroid glands in primary and secondary hyperparathyroidism. Kidney Int. 68, 998–1007. 3. Inomata, N., Akiyama, M., Kubota, N., and Jüppner, H. (1995). Characterization of a novel PTH-receptor with specificity for the carboxyl-terminal region of PTH(1-84). Endocrinology 136, 4732–4740. 4. Usdin, T. B., Gruber, C., and Bonner, T. I. (1995). Identification and functional expression of a receptor selectively recognizing parathyroid hormone, the PTH2 receptor. J. Biol. Chem. 270, 15455–15458. 5. Usdin, T. B., Wang, T., Hoare, S. R. J., Mezey, E., and Palkovits, M. (2000). New members of the parathyroid hormone/parathyroid hormone receptor family: the parathyroid hormone 2 receptor and tuberinfundibular peptide of 39 residues. Frontiers in Neuroendocrinology 21, 349–383. 6. Slatopolsky, E., Finch, J., Clay, P., Martin, D., Sicard, G., Singer, G., Gao, P., Cantor, T., and Dusso, A. (2000). A novel mechanism for skeletal resistance in uremia. Kidney Int. 58, 753–761.

511 7. Nguyen-Yamamoto, L., Rousseau, L., Brossard, J. H., Lepage, R., and D’Amour, P. (2001). Synthetic carboxyl-terminal fragments of parathyroid hormone (PTH) decrease ionized calcium concentration in rats by acting on a receptor different from the PTH/PTH-related peptide receptor. Endocrinology 142, 1386–1392. 8. Divieti, P., Inomata, N., Chapin, K., Singh, R., Jüppner, H., and Bringhurst, F. (2001). Receptors for the carboxyl-terminal region of PTH(1-84) are highly expressed in osteocytic cells. Endocrinology 142, 916–925. 9. Divieti, P., John, M., Jüppner, H., and Bringhurst, F. (2002). Human PTH-(7-84) inhibits bone resorption in vitro via actions independent of the type 1 PTH/PTHrP receptor. Endocrinology 143, 171–176. 10. Demay, M., Mitchell, J., and Goltzman, D. (1985). Comparison of renal and osseous binding of parathyroid hormone and hormonal fragments. Am. J. Phyisol. 249, E437–E446. 11. McKee, M. D., and Murray, T. M. (1985). Binding of intact parathyroid hormone to chicken renal membranes: evidence for a second binding site with carboxyl-terminal specificity. Endocrinology 117, 1930–1939. 12. Murray, T. M., Rao, L. G., and Muzaffar, S. A. (1991). Dexamethasonetreated ROS 17/2.8 rat osteosarcoma cells are responsive to human carboxylterminal parathyroid hormone peptide hPTH(53-84): stimulation of alkaline phosphatase. Calcif. Tissue Int. 49, 120–123. 13. Rao, L. G., and Murray, T. M. (1985). Binding of intact parathyroid hormone to rat osteosarcoma cells: major contribution of binding sites for the carboxyl-terminal region of the hormone. Endocrinology 117, 1632–1638. 14. Langub, M., Monier-Faugere, M., Wang, G., Williams, J., Koszewski, N., and Malluche, H. (2003). Administration of PTH-(7-84) antagonizes the effects of PTH-(1-84) on bone in rats with moderate renal failure. Endocrinology 144, 1135–1138. 15. Brossard, J. H., Clouthier, M., Roy, L., Lepage, R., Gascon-Barre, M., and D’Amour, P. (1996). Accumulation of a non-(1-84) molecular form of parathyroid hormone (PTH) detected by intact PTH assay in renal failure: importance in the interpretation of PTH values. J. Clin. Endocrinol. Metab. 81, 3923–3929. 16. Lepage, R., Roy, L., Brossard, J. H., Rousseau, L., Dorais, C., Lazure, C., and D’Amour, P. (1998). A non-(1-84) circulating parathyroid hormone (PTH) fragment interferes significantly with intact PTH commercial assay measurements in uremic samples. Clin. Chem. 44, 805–809. 17. Brossard, J., Lepage, R., Cardinal, H., Roy, L., Rousseau, L., Dorais, C., and D’Amour, P. (2000). Influence of glomerular filtration rate on non(1-84) parathyroid hormone (PTH) detected by intact PTH assays. Clin. Chem. 46, 697–703. 18. D’Amour, P., Brossard, J. H., Rakel, A., Rousseau, L., Albert, C., and Cantor, T. (2005). Evidence that the amino-terminal composition of non-(1-84) parathyroid hormone fragments starts before position 19. Clin. Chem. 51, 169–176. 19. Kunii, I., and Vieira, J. (2001). Circulating forms of parathyroid hormone detected with an immunofluorometric assay in patients with primary hyperparathyroidism and in hyperparathyroidism secondary to chronic renal failure. Braz. J. Med. Biol. Res. 34, 1547–1550. 20. Gao, P., Scheibel, S., D’Amour, P., John, M., Rao, S., Schmidt-Gayk, H., and Cantor, T. (2001). Development of a novel immunoradiometric assay exclusively for biologically active whole parathyroid hormone 1-84: implications for improvement of accurate assessment of parathyroid function. J. Bone Miner. Res. 16, 605–614. 21. Berson, S. A., Yalow, R. S., Aurbach, G., and Potts, J. T., Jr. (1963). Immunassay of bovine and human parathyroid hormone. Proc. Natl. Acad. Sci. USA 49, 613–617. 22. Segre, G. V., and Potts, J. T., Jr. (1995). Differential Diagnosis of Hypercalcemia. In: “Endocrinolgy” (L. J. DeGroot, ed.), pp. 1075–1093, Philadelphia: W. B. Saunders. 23. Nussbaum, S. R., Zahradnik, R. J., Lavigne, J. R., Brennan, G. L., Nozawa-Ung, K., Kim, L. Y., Keutmann, H. T., Wang, C. A., Potts, J. T., Jr.,

512

24. 25.

26. 27. 28.

29.

30.

31.

32.

33.

34.

35.

36.

37.

38.

and Segre, G. V. (1987). Highly sensitive two-site immunoradiometric assay of parathyrin, and its clinical utility in evaluating patients with hypercalcemia. Clin. Chem. 33, 1364–1367. Jüppner, H., and Potts, J. T., Jr. (2002). Immunoassays for the detection of parathyroid hormone. J. Bone Miner. Res. 17 Suppl 2:N81–86. Goodman, W. G., Jüppner, H., Salusky, I. B., and Sherrard, D. J. (2003). Parathyroid hormone (PTH), PTH-derived peptides, and new PTH assays in renal osteodystrophy. Kidney Int. 63, 1–11. Martin, K. J., Akhtar, I., and Gonzalez, E. A. (2004). Parathyroid hormone: new assays, new receptors. Semin. Nephrol. 24, 3–9. Ekins, R. (1980). More sensitive immunoassays. Nature 284, 14–15. D’Amour, P., Brossard, J. H., Rousseau, L., Roy, L., Gao, P., and Cantor, T. (2003). Amino-terminal form of parathyroid hormone (PTH) with immunologic similarities to hPTH(1-84) is overproduced in primary and secondary hyperparathyroidism. Clin. Chem. 49, 2037– 2044. John, M., Goodman, W., Gao, P., Cantor, T., Salusky, I., and Jüppner, H. (1999). A novel immunoradiometric assay detects full-length human PTH but not amino-terminally truncated fragments: implications for PTH measurements in renal failure. J. Clin. Endocrinol. Metab. 84, 4287–4290. Salusky, I. B., Goodman, W. G., Kuizon, B. D., Lavigne, J. R., Zahranik, R. J., Gales, B., Wang, H. J., Elashoff, R. M., and Juppner, H. (2003). Similar predictive value of bone turnover using first- and second-generation immunometric PTH assays in pediatric patients treated with peritoneal dialysis. Kidney Int. 63, 1801–1808. Hatakeyama, Y., Mizunashi, K., Furukawa, Y., Yabuki, S., Sato, Y., and Igarashi, T. (2003). Plasma levels of parathyroid hormone (1-84) whole molecule and parathyroid hormone (7-84)-like fragments in pseudohypoparathyroidism type I. J. Clin. Endocrinol. Metab. 88, 2250–2255. Savoca, R., Bock, A., Kraenzlin, M. E., Schmid, H. R., and Huber, A. R. (2004). An automated ‘bio-intact’ PTH assay: a step towards standardisation and improved correlation with parathyroid function in renal disease. Clin. Chim. Acta 343, 167–171. Silverberg, S. J., Gao, P., Brown, I., LoGerfo, P., Cantor, T. L., and Bilezikian, J. P. (2003). Clinical utility of an immunoradiometric assay for parathyroid hormone (1-84) in primary hyperparathyroidism. J. Clin. Endocrinol. Metab. 88, 4725–4730. Yamashita, H., Gao, P., Cantor, T., Noguchi, S., Uchino, S., Watanabe, S., Ogawa, T., Kawamoto, H., and Fukagawa, M. (2004). Comparison of parathyroid hormone levels from the intact and whole parathyroid hormone assays after parathyroidectomy for primary and secondary hyperparathyroidism. Surgery 135, 149–156. Bouillon, R., Coopmans, W., Degroote, D., Radoux, D., and Eliard, P. (1990). Immunoradiometric assay of parathyrin with polyclonal and monoclonal region-specific antibodies. Clin. Chem. 36, 271–276. Fournier, A., Solal, M., Oprisiu, R., Mazouz, H., Morinire, P., Choukroun, G., and Bouillon, R. (2001). Optimal range of plasma concentration of true 1-84 parathyroid hormone in patients on maintenance dialysis. J. Clin. Endocrinol. Metab. 86, 1840–1842. Carnevale, V., Dionisi, S., Nofroni, I., Romagnoli, E., Paglia, F., De Geronimo, S., Pepe, J., Clemente, G., Tonnarini, G., and Minisola, S. (2004). Potential clinical utility of a new IRMA for parathyroid hormone in postmenopausal patients with primary hyperparathyroidism. Clin. Chem. 50, 626–631. Silverberg, S., and Bilezikian, J. (2003). “Incipient” primary hyperparathyroidism: a “forme fruste” of an old disease. J. Clin. Endocrinol. Metab. 88, 5348–5352.

HARALD JÜPPNER AND GHADA EL-HAJJ FULEIHAN 39. Monier-Faugere, M., Geng, Z., Mawad, H., Friedler, R., Gao, P., Cantor, T., and Malluche, H. (2001). Improved assessment of bone turnover by the PTH-(1-84)/large C-PTH fragments ratio in ESRD patients. Kidney Int. 60, 1460–1468. 40. Kazama, J. J., Omori, T., Ei, I., Ei, K., Oda, M., Maruyama, H., Narita, I., Gejyo, F., Shigematsu, T., and Fukagawa, M. (2003). Circulating 1-84 PTH and large C-terminal PTH fragment levels in uremia. Clin. Exp. Nephrol. 7, 144–149. 41. Coen, G., Bonucci, E., Ballanti, P., Balducci, A., Calabria, S., Nicolai, G., Fischer, M., Lifrieri, F., Manni, M., Morosetti, M., Moscaritolo, E., and Sardella, D. (2002). PTH 1-84 and PTH “7-84” in the noninvasive diagnosis of renal bone disease. Am. J. Kidney Dis. 40, 348–354. 42. Inaba, M., Okuno, S., Imanishi, Y., Ueda, M., Yamakawa, T., Ishimura, E., and Nishizawa, Y. (2004). Significance of Bio-intact PTH(1-84) assay in hemodialysis patients. Osteoporos Int. 16, 517–525. 43. Sherrard, D., Hercz, G., Pei, Y., Maloney, N., Greenwood, C., Manuel, A., Saiphoo, C., Fenton, S., and Segre, G. (1993). The spectrum of bone disease in end-stage renal failure—an evolving disorder. Kidney Int. 43, 436–442. 44. Salusky, I., Coburn, J., Brill, J., Foley, J., Slatopolsky, E., Fine, R., and Goodman, W. (1988). Bone disease in pediatric patients undergoing dialysis with CAPD or CCPD. Kidney Int. 33, 975–982. 45. Yamashita, H., Cantor, T., Uchino, S., Watanabe, S., Ogawa, T., Moriyama, T., Takamatsu, Y., Fukagawa, M., and Noguchi, S. (2005). Sequential changes in plasma intact and whole parathyroid hormone levels during parathyroidectomy for secondary hyperparathyroidism. World J. Surg. 29, 169–173. 46. Weinstein, L., Yu, S., Warner, D., and Liu, J. (2001). Endocrine Manifestations of Stimulatory G Protein alpha-Subunit Mutations and the Role of Genomic Imprinting. Endocr. Rev. 22, 675–705. 47. Jan de Beur, S. M., and Levine, M. A. (2001). Pseudohypoparathyroidism: Clinical, Biochemical, and Molecular Features. In: “The Parathyroids: Basic and Clinical Concepts” (J.P. Bilezikian, R. Markus, and M.A. Levine, eds.), pp. 807–825, New York: Academic Press. 48. Bastepe, M., and Jüppner, H. (2005). The GNAS locus and pseudohypoparathyroidism. Hormone Research 63, 65–74. 49. Bastepe, M., Fröhlich, L., Hendy, G., Indridason, O., Josse, R., H, K., Körkkö, J., Nakamoto, J., Rosenbloom, A., Slyper, A., Sugimoto, T., Tsatsoulis, A., Crawford, J., and Jüppner, H. (2003). Autosomal dominant pseudohypoparathyroidism type-Ib is associated with a heterozygous microdeletion that likely disrupts a putative imprinting control element of GNAS. J. Clin. Invest. 112, 1255–1263. 50. Laspa, E., Bastepe, M., Jüppner, H., and Tsatsoulis, A. (2004). Phenotypic and molecular genetic aspects of pseudohypoparathyroidism type Ib in a Greek kindred: evidence for enhanced uric acid excretion due to parathyroid hormone resistance. J. Clin. Endocrinol. Metab. 89, 5942–5947. 51. Mahmud, F., Linglart, A., Bastepe, M., Jüppner, H., and Lteif, A. (2005). Molecular diagnosis of pseudohypoparathyroidism type Ib in a family with presumed paroxysmal dyskinesia. Pediatrics 115, e242–244. 52. Bastepe, M., Fröhlich, L., Linglart, A., Abu-Zahra, H., Tojo, K., Ward, L., and Jüppner, H. (2005). Deletion of the NESP55 differentially methylated region causes loss of maternal GNAS imprints and pseudohypoparathyroidism type Ib. Nat. Genet. 37, 25–27.

Chapter 31

New Horizons for Assessment of Vitamin D Status in Man Gary L. Lensmeyer, Neil Binkley, and Marc K. Drezner

III. Measurement of 1,25-Dihydroxyvitamin D References

I. Measurement of Vitamin D3 (Cholecalciferol) and Vitamin D2 (Ergocalciferol) II. Measurement of 25-Hydroxyvitamin D

I. MEASUREMENT OF VITAMIN D3 (CHOLECALCIFEROL) AND VITAMIN D2 (ERGOCALCIFEROL)

II. MEASUREMENT OF 25-HYDROXYVITAMIN D Over the past several decades the measurement of 25-hydroxyvitamin D (25(OH)D) has emerged as the presumed ideal index to assess vitamin D status in humans. The utility of this measurement is predicated on several factors, which include: (1) the rapid transformation of vitamin D to 25(OH)D; (2) the confinement of 25(OH)D in large measure to the extracellular compartment, where it is bound to vitamin D binding protein; (3) the long half-life of 25(OH)D in the blood; and (4) the relatively high concentration of 25(OH)D in the circulation. Moreover, discovery that the circulating level of 1,25-dihyroxyvitamin D, the active form of vitamin D, is regulated as a hormone and provides no information regarding the nutritional vitamin D status or cutaneous production of vitamin D reaffirmed the utility of the 25(OH)D measurement. Thus, in 1997

Measurement of the circulating concentrations of the native vitamins D3 and D2 provides information about recent exposure to either nutritional vitamin D or to UVB radiation. However, vitamin D is removed from the circulation within hours and does not provide an integrated view of vitamin D stores. Moreover, measurement of vitamin D is difficult, since it is by far the most lipophilic of the antirachitic sterols and cannot be extracted from serum utilizing solid-phase techniques [1]. Therefore, assay of the native forms of vitamin D is not useful for assessing human vitamin D status and is not clinically available. Nevertheless, vitamin D measurements have proved useful for research, most notably in assessment of intestinal lipid-absorptive capacity and response to UVB exposure [2]. Dynamics of Bone and Cartilage Metabolism

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the Food and Nutrition Board of the Institute of Medicine accepted serum 25(OH)D concentration as the functional indicator of vitamin D status [3]. Recognition of the potential value that measurements of 25(OH)D would afford in identifying previously unappreciated widespread vitamin D insufficiency/deficiency spawned an increased demand for the measurement, creating the need for clinically feasible, validated assay procedures. As a result, over the past 10 years measurement of serum 25(OH)D concentrations has moved into the clinical laboratory setting, resulting in the development of multiple commercially available 25(OH)D assays, which have become frequently used tools of physicians worldwide. Despite this apparent progress, however, attempts to assess the vitamin D status in humans have been hampered by the considerable interlaboratory variation observed in the measurements of 25(OH)D (Table I) [4, 5]. Although such variability is undoubtedly due, in part, to the use of fundamentally different methodologies by various laboratories, the failure of several attempts to standardize, or at least harmonize, results from different laboratories indicates a more complex problem as the root source of the variability [19]. Moreover, the recently recognized difficulties that result from deriving normal ranges for the various assays employed from ostensibly normal populations has further confounded the ability to employ the 25(OH)D assay for a clinically useful purpose. In this regard, since there is a growing consensus that vitamin D insufficiency is more common than previously recognized and use of “normal” populations to define reference ranges in many countries has skewed the values to inappropriately low levels [20, 21]. These limitations have prompted a reappraisal of the methodologies available for measurement of 25(OH)D and the techniques to define the normal range

Table I.

Variability in the Reference Range for 25(OH)D

Range

Method

11–68 ng/ml 8–95 ng/ml 11–56 ng/ml 9–47 ng/ml 10–50 ng/ml 14–75 ng/ml 10–65 ng/ml 15–50 ng/ml 20–135 ng/ml 10–68 ng/ml 18–36 ng/ml 20–100 ng/ml 10–52 ng/ml

CPBA CPBA CPBA RIA CPBA RIA RIA RIA CPBA CLIA CPBA CPBA CLIA

Citation [6] [7] [8] [9] [10] [11] [12] [13] [14] [15] [16] [17] DiaSorin Corp.

Note: The fact that the “normal” range varies widely is well demonstrated by Carter et al. [18].

for this metabolite. Following is a review of these issues and insight to the appropriate use of 25(OH)D measurements.

A. What is the Normal Circulating Concentration of 25-Hydroxyvitamin D? Serum 25(OH)D measurements have been used to assess vitamin D status in patients and populations over the past 25 years. However, attempts to define the normal range for serum 25(OH)D have been complicated over the years by the impact of geography, seasonal variation, limited sun exposure, and skin color on the circulating levels of this metabolite. In this regard, it has long been recognized in populations with some sun exposure that the 25(OH)D concentration is higher in summer and lower in winter [7, 22], with the wintertime decrease accompanied by an elevation in circulating parathyroid hormone [23, 24]. The contribution of nutrition and sun exposure to the circulating 25(OH)D concentration varies with latitude, culture, and local dietary habits [25–28]. To this end, skin serves as a photoendocrine organ in which previtamin D3 is produced [29–31]. Upon skin exposure to ultraviolet B (UVB) radiation (290–315 nm) 7-dehydrocholesterol is converted to precholecalciferol in the basal layer of the epidermis. Precholecalciferol spontaneously isomerizes to form cholecalciferol, which exits the skin and is bound by vitamin D-binding protein in the circulation [32]. Since few foods contain appreciable amounts of vitamin D, in the absence of supplementation, cutaneous synthesis provides the majority of human vitamin D. Therefore, season and latitude (surrogates for UVB exposure) profoundly impact the “normal” 25(OH)D status, as does skin color, since melanin pigment is effective in scattering and absorbing the UVB wavelengths that cause vitamin D synthesis. As such, high concentrations of melanin in the skin reduce conversion of 7-dehydrocholesterol to previtamin D3 [33]. In fact, it has been suggested that early humans (approximately 1.2 million years ago) had darkly pigmented skin and that skin depigmentation evolved in humans living at higher latitudes because of the necessity to maintain adequate vitamin D3 production for as long as possible throughout the year [33]. Given the above, it is not surprising that individuals with dark skin living at northern latitudes and those with low sun exposure are at most risk for seasonal vitamin D inadequacy [34]. As a result of these observations, it has been assumed that use of “normal” populations from all geographic sites to define the normal 25(OH)D range is flawed and that it is more logical to utilize individuals who are highly sunexposed to define normal human vitamin D status [35]. When such an approach is taken, the mean serum 25(OH)D

Chapter 31 New Horizons for Assessment of Vitamin D Status in Man

Figure 1 Mean serum 25OHD concentration in highly sun-exposed adult humans. The bars on the left represent individual values from ten highly sun-exposed young adults in Honolulu, HI (latitude 21⬚N). These individual values are similar to those previously reported in comparable populations.

concentration among groups of such individuals ranges from approximately 45–65 ng/ml (Fig. 1) [8, 36–38]. Despite the above noted variability in 25(OH)D status, physicians have sought to overcome such impediments by developing site- and season-specific normal ranges. For example, Schmidt-Gayk [39] has reported developing normal ranges for the southwest of Germany (Heidelberg) 20–100 ng/ml for the summer and 10–50 ng/ml for the winter. However, the patient populations in which samples have been collected have compromised such efforts. In this regard, a reference range for an assay has been determined from samples obtained in population studies or from ostensibly healthy subjects of variable age [40–43]. Since the clinical and biochemical phenotype of people with mild vitamin D inadequacy or excess is not always immediately apparent, subjects with “abnormal” vitamin D status are often included amongst those sampled to obtain a normal reference range. Indeed, such sampling problems are compounded when the greater part of a population studied is vitamin D insufficient/deficient [20, 44], or manifest vitamin D abundance, due to common use of vitamin D supplements, ingestion of foods fortified with vitamin D, or abundant sunshine exposure [45, 46]. An alternative more appropriate approach to define the normal 25(OH)D range is to establish the physiological optimal range for this metabolite. Since vitamin D is a nutrient, which exhibits ‘threshold behavior’ (i.e. the values for the physiological responses change directly with intake up to some threshold value, above which the response does not change with further increases in intake) [21], several endpoints exist to define the physiological optimal range for 25(OH)D. These include measurements of intestinal calcium absorption [21], serum parathyroid hormone [40, 47, 48], serum 1,25(OH)2D [49], bone mineral density [50], and fracture risk [51, 52].

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In several studies 25(OH)D values have been determined that optimize intestinal calcium absorption and serve as the threshold below which calcium absorptive efficiency decreases [21, 53]. Similarly, threshold values have been identified in relatively large patient populations below which the mean serum parathyroid hormone level increases [40, 47, 48] and the serum 1,25(OH)2D level decreases [49]. Less well established is the relationship between the circulating 25(OH)D level and bone mineral density and fracture risk. Regardless, in population-based studies BischoffFerrari et al. [54] have documented in younger and older subjects a positive association between 25(OH)D levels and bone mineral density and Trivedi et al. [52], showed in older adults that treatment with vitamin D increased the 25(OH)D concentration to a level which decreased fracture risk at the hip, forearm, or spine. Since calcium absorption, as well as circulating levels of parathyroid hormone and 1,25(OH)2D, are dependent upon variables other than the serum 25(OH)D level, defining the threshold value of 25(OH)D using these endpoints is less precise than desired. As an example, calcium absorption occurs by both vitamin D-dependent gastrointestinal absorption and passive diffusion, a process dependent upon dietary calcium intake. Thus, an individual with high dietary calcium intake may meet the demand for calcium absorption largely by passive diffusion and relatively independent of 25(OH)D. Similarly, calcium directly influences serum parathyroid hormone and 1,25(OH)2D levels. Hence, the 25(OH)D threshold level for each of these analytes may vary with the availability of dietary calcium. Despite these recognized limitations, studies of calcium and vitamin Drelated functions has led to classification of vitamin D status as follows [55–57]: • Vitamin D replete
Serum 25(OH) D levels above the threshold level, with a normal serum parathyroid hormone concentration and normal bone histology. • Vitamin D insufficient
Serum 25(OH)D levels below the threshold, resulting in a tendency to increase the serum parathyroid hormone concentration and enhance bone turnover, but sufficiently high to maintain normal bone mineralization. • Vitamin D deficient
Serum 25(OH)D levels far below the threshold and sufficiently low to increase the serum parathyroid hormone level, enhance bone turnover and result in incipient or overt osteomalacia in adults and rickets and osteomalacia in children. Such classification has led to establishing the diagnosis of inadequate vitamin D status in a substantial proportion of the populations in diverse countries worldwide.

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However, although the justification for classification is well established, diagnosis of vitamin D status in an individual according to this schema is fraught with difficulty. In this regard, several factors confound the ability to set the decision limits for classification of patients at uniform and widely accepted levels. Among these are variables such as: (1) dietary calcium, which may independently influence the set-point for a threshold value in an individual patient, as described above; (2) use of uniquely different patient populations to establish threshold values; and (3) the interlaboratory variability of 25(OH)D measurements performed by similar or different techniques. Thus, the 25(OH)D threshold below which parathyroid hormone increase has been variably reported as 10–44 ng/ml [40, 47–49, 58], while that for optimal calcium absorption ranges from 26–36 ng/ml. Not surprisingly, therefore, Glendenning et al. [59] confirmed the significant clinical conundrum that precludes simple classification of patients according to their vitamin D status. They illustrated that, considering somewhat arbitrary medical decision limits of 12 and 20 ng/ml for vitamin D deficiency and insufficiency, respectively, measurement of 25(OH)D by two different immunoassays in duplicate samples resulted in diagnostic discordance in 42 of 130 (32%) patients. Thus, the variability in assay results not only affects efforts to establish threshold values, but the applicability of the resulting decision limits to patients when assay systems used to measure 25(OH)D in the patients differ from those employed in an original study, in which decision limits were established. Ironically, although the most commonly used decision limit to establish vitamin D insufficiency in current use is 32 ng/ml, according to a study by Heaney et al. [21], the assay used by the investigators was a commercially available measurement no longer available, thereby creating an assay dependent diagnostic discordance.

B. What is the Basis for 25-Hydroxyvitamin D Assay Variability The considerations noted above have clearly outlined the importance of developing standardized assays for 25(OH)D, which overcome the variability currently complicating this measurement. Such efforts will require appreciating the historical development of the assays and understanding the reasons for the disparate results. 1. COMPETITIVE BINDING PROTEIN ASSAYS

Until 1977 various competitive binding protein assays for 25(OH)D dominated the literature. For the most part these assays utilized the vitamin D-binding protein from rat serum as a specific binding agent, required some type

of organic extraction and sample purification by column chromatography, and employed [3H]-25(OH)D as a tracer to estimate endogenous loses of the metabolite during the extraction and purification procedures [8, 16, 17]. Not surprisingly, variations in extraction technique, incubation conditions, metabolite recovery and purification procedures are legion in these assays. In fact, some of the 25(OH)D assays use improper extraction techniques, resulting in matrix (lipid) interference with the assay and false elevations of the results [60]. Moreover, use of the vitamin Dbinding protein limited the specificity of the assays, since 24,25(OH)2D and 25,26(OH)2D compete with 25(OH)D for binding and are present in sufficient amounts in variable patient populations to again falsely elevate the assay values [39]. Thus, the “normal” ranges for such assays vary from 18–36 ng/ml to 20–100 ng/ml. Regardless, the requirement for sample extraction with an organic solvent, drying under N2, purification by column chromatography and recovery estimates to account for endogenous losses of 25(OH)D, rendered these assays relatively cumbersome. Hence, although they were appropriate for research laboratories, they did not qualify for use in high-throughput clinical laboratories. 2. RADIOIMMUNOASSAYS AND CHEMILUMINESCENT ASSAYS

Not surprisingly, as the clinical demand for serum 25(OH)D analysis increased, attempts to offer simpler, rapid, yet valid assays mounted. To this end, elimination of liquid–liquid organic extraction and chromatographic prepurification and evaporation of organic solvents was sought. With the introduction of the first “valid” radioimmunoassay (RIA) for assessing circulating 25(OH)D in 1985, this goal was partially met [61]. This RIA required only an acetonitrile extraction step, eliminating the need for sample prepurification prior to assay and a requirement for organic solvent evaporation, albeit [3H]-25(OH)D was used as the tracer. In 1993, further advances permitted development and use of [125I]-25(OH)D as tracer, increasing assay sensitivity and reducing the need for sample purification [62]. Thus, use of RIAs with [125I]-25(OH)D as tracer became a common choice of clinicians worldwide for measurement of 25(OH)D. Indeed, such an assay became the first test for vitamin D approved by the Food and Drug Administration for clinical diagnosis in the United States. The value of the RIA established by Hollis [62] and subsequently offered by DiaSorin, was amplified because the antibody used was raised against the synthetic vitamin D analog 23,24,25,26,27-pentanor vitamin D-C(22)-carboxylic acid. Coupling this compound to bovine serum albumin permitted generation of antibodies that theoretically cross-react equally with 25(OH)D3 and 25(OH)D2, since the analog retained the intact structure

Chapter 31 New Horizons for Assessment of Vitamin D Status in Man

of vitamin D only up to carbon 22, a common structure area of the 25(OH)D metabolites. Unfortunately, not all antibodies subsequently developed for commercially available assays were similarly developed and thus do not interact equally with both 25(OH)D3 and 25(OH)D2. More recently, DiaSorin Corporation (Stillwater, MN) and Nichols Institute Diagnostics (San Clemente, CA) have introduced methods for the direct (no extraction) quantitative determination of 25(OH)D in serum or plasma utilizing a chemiluminescent assay. The assay offered by DiaSorin (performed on the LIAISON® platform) utilizes a specific antibody identical to that used in their RIA and similar to that originally developed by Hollis [62], which is coated onto magnetic particles. Tracer vitamin D is linked to an isoluminol derivative and, during incubation of the sample, the 25(OH)D is dissociated from its binding protein, competing with the labeled vitamin D for binding sites on the antibody. After incubation, unbound material is removed with a wash, starter reagents added and a flash chemiluminescent reaction created, the light signal from which is measured by a photomultiplier as relative light units. By contrast, the assay developed by Nichols Institute Diagnostics (performed on the ADVANTAGE® platform), although executed quite similarly to the DiaSorin technique, utilizes human serum vitamin D-binding protein as a competitive binder instead of an antibody. Despite the widespread availability of RIA and chemiluminescent 25(OH)D assays, which were created in response to the demand for rapid measurements of clinical samples, the reliability of these tests is a matter of great debate. In this regard, commercial binding/immunochemical assays in general have problems with accuracy and often generate false-positive and false-negative results [63]. The difficult task of immunoassay standardization and external quality assessment compounds the accuracy issue [64]. The major problem in measuring 25(OH)D by these methods is attributable to the molecule itself, since this metabolite is one of the most hydrophobic compounds measured by RIA or competitive protein binding assay (CPBA). When combined with the fact that it exists in two forms, 25(OH)D2 and 25(OH)D3, substantial analytical problems are created. The lipophilic nature of 25(OH)D renders it especially vulnerable to matrix effects in any RIA or CPBA. Such matrix effects result from the presence of a compound in the sample assay tubes, which is not present in standard assay tubes. These substances are usually lipid, but in the chemiluminescent assays, performed without sample prepurification, anything contained in a serum or plasma sample may serve as a substance creating a matrix effect. Regardless, such matrix factors simply change the ability of the binding agent, antibody, or binding protein to associate with 25(OH)D in a sample. When this occurs, it markedly

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diminishes the validity of the assay. Although experience indicates that CPBA and/or chemiluminescent assays are more susceptible to such matrix effects, RIAs suffer similar interference [65]. Therefore, it is not surprising that significant concern has emerged regarding the currently available chemiluminescent assays and RIAs. Indeed, recent studies have clearly established that the Nichols Advantage assay performs inadequately on clinical samples, correlating poorly with both RIA and HPLC determinations (Fig. 2) [5, 66]. This is not unexpected since the vitamin D-binding protein is susceptible to the matrix effects noted above [67]. Perhaps more importantly, however, the validity of the Nichols Advantage assay is further confounded by limited ability to detect 25(OH)D2, thereby failing to function adequately in monitoring therapy with ergocalciferol (a high-dose vitamin D preparation commonly prescribed in the United States) [68, 69]. Such shortcomings have resulted in withdrawal of this assay from use in many commercial laboratories. There is less certainty regarding the Liaison chemiluminiscent assay. Although Hollis [66] and Ersfeld et al. [70] indicate that the automated assay is in excellent agreement with the widely used DiaSorin RIA in both normal and chronic renal failure patients, several investigators [69, 71] found poor correlation of this assay with RIA and HPLC determinations (Fig. 2). Hence, question remains regarding the clinical utility of the Liaison assay. Unfortunately, recent studies have also raised questions regarding the accuracy and precision of measurements obtained from any single RIA. In this regard, several investigators have compared serum 25(OH)D levels obtained by RIA to those measured by HPLC, a gold standard for the assay frequently used in the research environment. Although Hollis [66] and Turpeinen (71) reported excellent correlation of the values obtained by RIA to those by HPLC, Lips et al. [19] and Gemar et al. [69] have found significant discordance between the values obtained by these assay techniques (Fig. 3). Moreover, Binkley and colleagues [5] have discovered that the reproducibility of the RIA measurement (DiaSorin) is highly dependent upon the laboratory in which the assay is performed (Fig. 4) and varies significantly upon repetitive measures in the same laboratory (Fig. 5). In addition, Vieth et al. [72] have raised significant questions regarding RIA assay validity in samples with low 25(OH)D concentration. Moreover, Hollis found significant differences in the performance of the available RIAs. He compared quantification of circulating 25(OH)D, using two commercially available Food and Drug Administration-approved 125I-based RIA kits (DiaSorin, Stillwater, MN and IDS Ltd, Tyne and Wear,

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Figure 2

(A) Correlation between serum 25(OH)D levels obtained in duplicate samples by the Nichols Advantage Assay and an HPLC assay performed at the University of Wisconsin. The data are plotted using a Lowess curve and 95% confidence limits. In general the correlation is better when values are <30 ng/ml. However, as the values increase the 95% confidence limits broaden considerably. In any case the positive bias of the Nichols Advantage Assay is 18.7 ng/ml. Moreover, the correlation is not linear throughout the range of values, challenging the assumption that establishing a normal reference range for the Nichols Advantage Assay overcomes the dilemma of assessing vitamin D status in individuals. (B) Correlation between serum 25(OH)D levels obtained in duplicate samples by the DiaSorin Liaison Assay and an HPLC assay performed at the University of Wisconsin. The data are plotted using a Lowess curve and 95% confidence limits. The positive bias of the DiaSorin Liaison Assay is 5.9 ng/ml. However, the correlation is not linear throughout the range of values, challenging the assumption that establishing a normal reference range for the DiaSorin Liaison Assay overcomes the dilemma of assessing vitamin D status in individuals.

United Kingdom) [73]. Both methods quantitatively recovered 25(OH)D3 added to serum, but only the DiaSorin kit quantitatively recovered 25(OH)D2, because the primary antibody from the IDS kit had unequal reactivity with pure 25(OH)D2 and 25(OH)D3. These data indicate that some immunoassays may underestimate 25(OH)D when 25(OH)D2 constitutes an appreciable part of the total.

Figure 3

3. HPLC ASSAYS For 25-HYDROXYVITAMIN D

Validation of the commercially available rapid 25(OH)D assays has required a gold standard for comparison. For the most part the HPLC method for detection of circulating 25(OH)D has served this role. First introduced in 1977 from the laboratory of Dr. Hector DeLuca, this assay allowed assessment of 25(OH)D2 and 25(OH)D3 by

Correlation between serum 25(OH)D levels obtained in duplicate samples by the DiaSorin RIA and an HPLC assay at the University of Wisconsin. The RIA was performed in a commercial laboratory (A) and a research laboratory (B) with extensive experience in the measurement, using two different sets of samples. The data are plotted using Lowess curves and 95% confidence limits. The relationships are similar in both cases, albeit the correlation for samples measured in the research laboratory (B) is better. Nevertheless, in both cases, there is significant discordance between the 25(OH)D levels obtained by RIA and HPLC. Indeed, a positive bias for the RIA measurements exists for both sets of data. Moreover, the relationships are curvilinear, raising significant questions regarding whether establishing a normal reference range for the RIA will permit accurate identification of vitamin D status in individuals.

Chapter 31 New Horizons for Assessment of Vitamin D Status in Man

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Figure 4

Correlation between 25(OH)D measurements obtained in duplicate samples by the DiaSorin RIA and an HPLC performed at the University of Wisconsin. The samples measured by RIA in two different laboratories (A and B) were identical. The correlation between the RIA and HPLC data was highly variable, according to the laboratory in which the measurements were made. As a consequence the measurements made by RIA in the two different laboratories had poor correlation with each other (C). These observations document that the reproducibility of the DiaSorin RIA measurements is highly dependent upon the laboratory in which the assay is performed.

Figure 5 Correlation between RIA measurements of 25(OH)D made in the same laboratory on duplicate samples. The variability in the performance of the DiaSorin RIA is again demonstrated despite repetitive measurements in the same laboratory.

UV detection [74]. Although this assay is very accurate if validated and performed by experienced personnel, inaccurate HPLC methods exist and can provide misleading results. Underlying such inaccuracy is co-migration of UV-detectable products with the vitamin D metabolites, particularly 25(OH)D2 [68]. Regardless, over subsequent years additional quantitative HPLC assays have been developed based on UV detection and normal-phase separation [75], combined use of normal- and reversed-phase separations [76], or reversed-phase separation alone [77]. Recently, a reversed-phase HPLC method for 25(OH)D3 in human plasma has been developed with normal-phase prepurification of the sample [78] or liquid extraction only [79]. The initial attempts to develop such assays had disadvantages, including relatively large sample volume requirements, slow sample throughput, use of radiolabeled 25(OH)D to trace recovery of the endogenous metabolite and employment of imprecise normal-phase chromatography, limitations which confined their use to research laboratories. However, the recently developed HPLC assays involve relatively simple extraction procedures, obviate the need for radiolabeled 25(OH)D, and use reproducible

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reversed-phase chromatography. Hence relatively simple and sensitive HPLC methods are now available for 25(OH)D measurements in commercial laboratories [71]. 4. LIQUID CHROMATOGRAPHY/TANDEM MASS SPECTROMETRY ASSAY OF 25-HYDROXYVITAMIN D

Within the last 4 years, the evident problems inherent to the various assay methods for 25(OH)D have resulted in efforts to create a reference method for the measurement of 25(OH)D3 and 25(OH)D2. Although highly accurate gas chromatography–mass spectrometry methods were developed years ago [80–83], they are extremely complex and did not gain widespread use for quality-control programs or routine assay validation. Since liquid chromatographytandem mass spectrometry (LC/MS/MS) requires less time-consuming sample purification and far shorter run times and has been recently accepted in clinical chemistry as a reference system [84, 85], convenient and specific isotope-dilution LC/MS/MS methods for quantification of 25(OH)D have been developed. The principle of isotope dilution with stable-isotope-labeled internal standard compounds and mass spectrometric detection is generally accepted as yielding the highest attainable analytical accuracy. Moreover, any matrix effect of an assay is compensated, in part, by isotope dilution standardization. Vogeser et al. [86] recently validated such an assay for 25(OH)D3. The technique includes semiautomated sample preparation and the practicable LC/MS/MS methodology, allowing for use in routine laboratory settings. A multicenter validation of the method is currently planned. Not surprisingly, however, recent studies have indicated that assay of 25(OH)D by LC/MS/MS is not without flaws. It appears that the measurement of 25(OH)D3 is confounded by the concurrent assay of 3-epi-25(OH)D3 [87], which is increased to interfering levels in youths with a variety of metabolic bone diseases [88]. In any case, this method can also be adopted for measurement of 25(OH)D2, when circulating levels of this metabolite exceed approximately 5.0 ng/ml, values likely encountered in patients receiving ergocalciferol supplements. Alternatively, simultaneous determination of 25(OH)D3 and 25(OH)D2 is possible with greater sensitivity using LC/MS/MS and Cookson-type derivatization, a procedure that is complex and labor-intensive [89].

C. Contemporary Approach to Measurement of 25-Hydroxyvitamin D It is clear from the above that vitamin D status is the complex result of sunshine exposure, latitude, sunscreen use, clothing, skin pigmentation, dietary intake, and use of vitamin D supplements. The consequences of vitamin D

deficiency, however, depend on calcium intake and other variables, which independently influence parathyroid function, vitamin D metabolism, and other variables. Thus, establishing the diagnosis of vitamin D insufficiency or deficiency depends upon: (1) defining the threshold levels of 25(OH)D, below which the variable complications of inadequate vitamin D manifest; and (2) ascertaining the serum 25(OH)D levels in at-risk patients. Unfortunately, substantial interlaboratory variation has confounded the 25(OH)D measurement, secondary to assay flaws, which are idiosyncratic to the method used, and to the technical expertise available in individual laboratories. Although there is no ideal solution to these problems at the present time, there are practical approaches to improving the assessment of the vitamin D status in patients. In this regard, physicians and laboratory directors must employ careful judgment in choosing testing methods for clinical applications. In our opinion, such judgment should result in coordinated efforts of the clinical and laboratory communities to implement existing methodologies, which measure 25(OH)D3 and 25(OH)D2 individually and with minimal interference. With the development of such assays, we envision that physicians and the laboratory community worldwide will strive for universal standardization of methods through the availability of validated primary 25(OH)D3- and 25(OH)D2- containing serum standards. Recognizing the need for such standardization in the United States, the National Institute of Standards and Technology (NIST), working with the Office of Dietary Supplements at the National Institutes of Health, has initiated efforts to prepare Standard Reference Material (SRM) for 25(OH)D in serum. At present it is anticipated that serum samples containing low (16 ng/ml) and normal (80 ng/ml) levels of 25(OH)D3 and 25(OH)D2, respectively, will be available. Coupled with a program to test the proficiency of available assays with clinical samples containing both metabolites, the availability of the SRM will markedly improve the ability of physicians to accurately assess the vitamin D status in patients and to compare results between populations and clinical studies. In the following sections of this chapter, we detail pragmatic ongoing pursuits, designed to improve the availability of optimal assays for 25(OH)D, noting caveats to each. We have focused on the newly emerging commercially available HPLC and LC/MS/MS assays, which provide selective measurements of 25(OH)D3 and 25(OH)D2 and the widely available DiaSorin RIA. 1. HPLC ASSAY OF 25(OH)D3 AND 25(OH)D2 IN THE CLINICAL LABORATORY

Although HPLC techniques for measurement of 25(OH)D have existed since 1977 [74], they have rarely

Chapter 31 New Horizons for Assessment of Vitamin D Status in Man

been offered in the clinical laboratory because of relatively large sample volume requirements, slow sample throughput, use of radiolabeled 25(OH)D to trace recovery of the endogenous metabolite and employment of normal phase chromatography with variable retention times, due to unanticipated changes in sorbent characteristics. To overcome such limitations, we have recently developed an assay in which 25(OH)D3 and 25(OH)D2 are selectively measured by semiautomatic solid-phase extraction and isocratic reverse-phase HPLC [69, 90]. This assay requires as little as 0.5 ml of serum, includes laurophenone as a nonradioactive tracer to assess recovery of the vitamin D metabolites and permits reasonably rapid processing of samples with a 17-minute elution time from reverse-phase HPLC [90]. Indeed, the time to the first result is little different than the 3.5 hours required for RIA of 25(OH)D. The assay involves four steps: (1) precipitation of protein and addition of laurophenone as an internal standard to trace recovery; (2) extraction of vitamin D metabolites and standard from the sample supernatant with a solidphase sorbent, using the automated Gilson ASPEC XL4 (Gilson, Inc., Middleton, WI); (3) concentration of the extract; and (4) HPLC analysis. a. Protein Precipitation • Into a test tube containing 2.0 ml of precipitation reagent (laurophenone 400 µg/L in acetonitrile), add 1 ml of serum. Do not mix. Allow the tube to set at room temperature for 5 minutes. • Vortex-mix the tube and allow to set for an additional 5 minutes. • Centrifuge the tube for 10 minutes and decant clear supernatant into a glass test tube. b. Extract Compounds with Solid-Phase Sorbent The automated Gilson ASPEC XL4 will: • Prime the Strata-X sorbent cartridge* with 2 ml acetonitrile followed by 1 ml acetonitrile/water, 35/65, v/v.
Phenomenex Strata-X™ surface-modified styrenedivinylbenzene, 60 mg sorbent in 1 ml-size cartridge. • Add 1.0 ml water to the sample supernatant. • Load 3.5 ml of sample onto the Strata-X sorbent. • Wash the sorbent with 2 ml acetonitrile/water, 35/65, v/v. • Elute the compounds with 2 ml acetonitrile. c. Concentrate Extract • Evaporate acetonitrile in extract to dryness under nitrogen (10 psi) using a Zymark Turbo/Vac at 35º for 25 minutes. • Reconstitute the dry sample extract using a two-step process:
Add 150 µl of ethyl acetate/acetonitrile, 5/95, v/v, to the dry extract and vortex-mix for 10 seconds






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Add 110 µl of water and vortex-mix for 10 seconds Centrifuge tube to settle precipate Load HPLC autosampler with clear liquid and inject 80 µl onto the HPLC column.

d. HPLC Analysis The HPLC parameters are: • Thermo Integrated HPLC instrument with UV3000 detector, P 4000 pump, AS 2000 autosampler, and SCM 1000 solvent system. • Methanol/Water, 670/330, v/v, mobile phase. • Columns:
Analytical – Zorbax™ StableBond Cyanoprpyl (SB-CN), 4.6 × 250 mm, 5-micron particle
Guard – Zorbax SB-CN, 4.6 × 12.5 mm, 5-µm particle
Mobile Phase Conditioning – 4.6 × 250 mm column manually packed with silica, 80-µm particle. • Oven temperature at 50⬚C. • Flow rate at 1.2 ml/minute. • Detection at 275 nm. • Run time at 17 minutes. Using this technique, extracted drug-free serum or plasma results in a chromatogram that is free of detectable peaks, which co-elute with the internal standard, laurophenone, and 25(OH)D2. Moreover, hemoglobin, bilirubin, and lipids do not cause interference or adversely affect recovery of the internal standard, 25(OH)D3 or 25(OH)D2. Figure 6 illustrates representative chromatograms. Extracted serum to which calibrator has been added results in a chromatogram that documents the presence and clear separation of eluted peaks of the internal standard, 25(OH)D3 and 25(OH)D2. Also documented is appropriate detection of an optimal concentration of 25(OH)D3, as well as a high concentration of 25(OH)D2, due to treatment with 25(OH)D2 and an accordingly increased concentration of this metabolite. Between-run precision coefficients of variation (CV) for the assay ranged from 2.6–4.9% for 25(OH)D3 and 3.2–12.6% for 25(OH)D2 (Table II). Upon additions of 25(OH)D3 and 25(OH)D2 to serum samples, we observed 95–102% recovery of the two analytes. Further, the assay was linear from 5 ng/ml to at least 100 ng/ml with a minimal detection limit of 3–4 ng/ml (Table III). Moreover, the results for patient samples tested by both HPLC and LC/MS/MS were nearly identical (Fig. 7). These observations clearly demonstrate that the HPLC assay for the measurement of 25(OH)D3 and 25(OH)D2 provides quality data and is suited for use in a clinical laboratory. Indeed, at the present time, we utilize this assay in the Clinical Laboratories of the University of Wisconsin

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Figure 6

Representative chromatograms obtained during HPLC analysis of 25(OH)D3 and 25(OH)D2 in serum samples. (A) Extracted serum to which calibrator has been added results in a chromatogram that documents the presence and clear separation of eluted peaks of the internal standard (laurophenone), 25(OH)D3 (68 ng/ml), and 25(OH)D2 (63 ng/ml). (B) Extracted serum sample containing an increased concentration of 25(OH)D2 (89.6 ng/ml), due to treatment with pharmacological amounts of this metabolite. (C) Extracted serum sample containing an optimal concentration of 25(OH)D3 (34.9 ng/ml), as well as a low concentration of 25(OH)D2 (<5 ng/ml).

Hospital and Clinics as the exclusive technique to measure 25(OH)D metabolites. Additional advantages of the assay are the ability of multiple analysts to successfully perform the HPLC method and the reasonable cost of establishing the method. Indeed, for hospital and research laboratories, this HPLC methodology seems ideal as an assay technique that will meet the goals for the ongoing optimization of 25(OH)D assays. Of course this HPLC assay is subject to interference by co-chromatography of interfering substances or vitamin D metabolites present in adequate concentration to confound measurement of 25(OH)D3 or 25(OH)D2. Thus far, we have identified interference only from substances leached from serum-separating gels used in some blood collection tubes. Therefore, it is essential that SST tubes are not used for blood collection when employing this HPLC assay. As studies progress, we anticipate that the 3-epi-25(OH)D3 [87] identified in high concentrations in children may interfere with measurement of 25(OH)D3. 2. LIQUID CHROMATOGRAPHY-TANDEM MASS SPECTROSCOPY ASSAY OF 25(OH)D3 AND 25(OH)D2 IN THE CLINICAL LABORATORY

As noted previously, convenient and specific isotopedilution LC/MS/MS methods for quantification of Table II.

Between Run Precision for the HPLC Assay

Control (n = 20) 1 2 3 4 5

25(OH)D3 25(OH)D2 25(OH)D3 25(OH)D2 25(OH)D3 25(OH)D2 25(OH)D3 25(OH)D2 25(OH)D3 25(OH)D2

Mean (ng/ml)

SD (ng/ml)

CV (%)

14.26 4.38 45.03 22.00 82.22 59.81 11.54 88.87 24.56 11.96

0.56 0.55 1.16 1.09 2.72 1.90 0.98 3.01 1.23 0.53

3.9 12.6 2.6 4.9 3.3 3.2 8.5 3.4 5.0 4.5

25(OH)D have been developed. Although the initial intent for development of such assays seemingly was to provide a reference method for standardization of 25(OH)D measurements, several commercial laboratories, including the Mayo Medical Laboratories [91] and the Nichols Laboratories, currently offer assay of the 25(OH)D metabolites by this technique. The advantages of this method are use of a relatively small volume of serum, a rapid processing time (60 minutes to the first result) and, like the HPLC systems, absence of interference by matrix effects. Moreover, accuracy and precision are respectable. Nevertheless, the cost of establishing such an assay system is prohibitive for many hospital and research laboratories. However, the purchase cost for individual samples from the commercial laboratories is quite reasonable. There is little published material on the methodological details used for LC/MS/MS by the commercial laboratories. However, an abstract detailing in brief the techniques employed at the Mayo Medical Laboratories recently appeared [91]. Deuterated stable isotope (d3-25(OH)D3) is added to a 0.2 ml plasma sample as internal standard. The 25(OH)D3, 25(OH)D2, and the internal standard are extracted using acetonitrile precipitation. Extracts are further purified online and analyzed by LC/MS/MS using multiple Table III. Linearity and Detection Limits of HPLC Assay 25(OH)D3 Concentration range n r Regression equation x = expected concentration y = analytical concentration Sy/x = Lower limit of detection Lower limit of quantitation

25(OH)D2

5–100 (ng/ml) 21 0.9972

5–100 (ng/ml) 21 0.9985

y = 1.00x + 0.873 (ng/ml) 2.61 (ng/ml) 3–4 (ng/ml) 5 (ng/ml)

y = 1.00x – 0.034 (ng/ml) 1.92 (ng/ml) 3–4 (ng/ml) 5 (ng/ml)

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Chapter 31 New Horizons for Assessment of Vitamin D Status in Man

Of some concern regarding this assay is the possibility that the measurements may be compromised to some extent by ion suppression. This is particularly plausible for measurements of 25(OH)D2, since only d3-25(OH)D3 is used as the internal standard, despite the availability of d3-25(OH)D2. Moreover, as noted above, measurement of 25(OH)D3 may be confounded by the concurrent assay of 3-epi-25(OH)D3 [88]. Nevertheless, the availability of commercially available LC/MS/MS measurements of 25(OH)D will undoubtedly help immeasurably in creating a favorable environment for standardizing the assay for 25(OH)D metabolites. 3. RADIOIMMUNOASSAY OF 25(OH)D

Figure 7 Correlation between 25(OH)D3 and 25(OH)D2 measurements made in duplicated samples by an HPLC assay performed at the University of Wisconsin and a LC/MS/MS assay. (A) The measurement of 25(OH)D3 by these techniques is virtually identical. (B) The measurement of 25(OH)D2 by these techniques is likewise virtually identical.

reaction monitoring. Interassay coefficients of variation are less than 15% at various 25(OH)D3 and 25(OH)D2 levels and precision is set at 2 ng/ml and 4.2 ng/ml for 25(OH)D3 and 25(OH)D2, respectively, based on interassay coefficients of variation of 12.9% and 14.0%. Such betweenrun precision compares favorably to that achieved with the HPLC assay described above. As noted above, Figure 7 illustrates the excellent correlation between 25(OH)D3 and 25(OH)D2 values obtained from analysis of multiple-patient samples by the LC/MS/MS and HPLC techniques. By contrast, the correlation between LC/MS/MS and RIA is less precise with an r2 of 0.86 and occurs with a positive bias of the RIA, creating a line significantly removed from the line of identity [91]. Table IV.

Clearly, among the well-established commercial measurements for 25(OH)D, the DiaSorin RIA offers the best commonly available alternative worldwide. The antibody developed for this measurement recognizes both 25(OH)D3 and 25(OH)D2, thereby allowing valid assessment of vitamin D status in patients receiving ergocalciferol. Moreover, several studies indicate that a measurement <30 ng/ml generally predicts a 25(OH)D concentration that is at or below this level, albeit the accuracy of the measurement in this range is lacking, as previously noted by Vieth et al. [72]. Therefore, reliable recognition of abnormally low vitamin D status is possible. In contrast, as the values exceed approximately 30 ng/ml, the assay displays variable accuracy and, in many cases, the relationship to the 25(OH)D concentration obtained by HPLC or LC/MS/MS exhibits a significantly different slope than that throughout the lower range of values (Fig. 3). Such variable correlation prevents establishing a valid reference range for the measurement throughout the span of values commonly encountered. An additional caveat confounding use of this assay is the technical expertise necessary to perform the assay, which contributes to an unpredictable variability in measurements from different laboratories (Figs 3 and 4). These observations have been reinforced by the results obtained from 36 laboratories, using the DiaSorin RIA, on samples distributed by DEQAS (Table IV) [92]. Such variability would clearly create a significant diagnostic discordance were individual samples measured in

DiaSorin RIA Mean and Range of Determinations on DEQAS Samples1 Sample number

Method DiaSorin RIA (n = 36) Method mean (ng/ml) Range (ng/ml) 1Samples

1

2

3

4

5

12.6 6.9–17.0

23.6 16.7–31.7

33.7 26.2–46.8

42.2 32.0–83.6

27.0 18.8–42.4

1–3 contain predominantly 25(OH)D3 and samples 4 and 5 contain 80% and 66% 25(OH)D2, respectively.

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different laboratories. Indeed, Vieth et al. have reported that the DiaSorin RIA exhibits a mean between-laboratory CV of 20–30%. Nevertheless, physicians in localities where the DiaSorin RIA is available may use this assay to confirm the presence of abnormal vitamin D status in patients clinically at risk for insufficiency and deficiency. They can refer to the studies of Heaney et al. [21, 56] and Chapuy et al. [40] to obtain the appropriate decision limits for diagnostic purposes, since these investigators used comparable immunoassays. In contrast, physicians should exert caution when using this assay to determine the possibility of vitamin D toxicity or to track the effects of therapy with vitamin D, since measurements at higher levels randomly display substantial discordance.

III. MEASUREMENT OF 1,25-DIHYDROXYVITAMIN D Measurement of 1,25(OH)2D is diagnostic for several clinical conditions, including vitamin D-dependent rickets, types 1 and 2, as well as those associated with hypercalcemia secondary to overproduction of this metabolite, such as sarcoidosis, tuberculosis, Hodgkin’s disease, fungal infections, Wegener’s granulomatosis, and lymphoma. Assessment of circulating 1,25(OH)2D levels in other circumstances is generally for confirmatory testing, such as in chronic renal failure, X-linked hypophosphatemia, hypoparathyroidism, and hyperparathyroidism.

A. Current Assays D. The Contemporary Normal Range for Serum 25(OH)D With movement of the medical and scientific communities towards standardizing 25(OH)D measurements and the adaptation of new assay methods, due consideration regarding establishing normal ranges for this measurement is required. Although it should not be difficult to determine reference ranges for the new assays in ostensibly normal patient populations, determining the decision limits for various vitamin D states may prove more difficult. Such decision limits have been created in various studies wherein calcium absorption, parathyroid hormone, and other factors have been used as endpoints. However, these studies have employed a wide range of serum 25(OH)D assays to establish the thresholds or decision limits. Clearly, this extensive literature is of extreme value and the medical community cannot discard this wealth of information. However, based on previous experience, we are likely to discover that there are substantial differences in the performance characteristics of the contemporary assays with those used in previous studies. Although such differences may be attributed, in part, to latitude and population characteristics [43, 45], interlaboratory variation is likely the primary underlying factor. Indeed, Lips et al. [19] clearly documented that 25(OH)D values from different laboratories are not comparable unless a careful cross-calibration has been performed. Thus, we should attempt to obtain such cross-calibration between the assays employed in studies wherein decision limits were defined and the newly emerging assays, in order that we can adapt the decision limits to the evolving contemporary assays. If we fail to do this, all assays will continue to discriminate well between low, average, and high values, but definition of vitamin D deficiency and insufficiency in absolute terms will remain elusive.

The measurement of 1,25(OH)2D has presented a difficult challenge because of a low circulating concentration, instability, a lipophilic structure, and the relative abundance of 25(OH)D, the precursor for the synthesis of the active vitamin D metabolite. With the introduction in 1984 of a radioreceptor assay, utilizing solid-phase extraction of 1,25(OH)2D from serum, along with silica cartridge purification and a vitamin D receptor isolated from calf thymus, measurement of circulating levels of this metabolite became widely available [93]. This assay was further simplified in 1986 by decreasing the required purification steps [94]. Nevertheless, the difficulty in isolating vitamin D receptor from calf thymus and the specificity of the receptor for the ligand, which allowed use of only [3H]-25(OH)D as the tracer for the assay (and not [125I]-based tracer), remained distinct limits of this assay. In 1996, however, Hollis et al. [95] reported development of an RIA for 1,25(OH)2D, which utilizes [125I]-25(OH)D tracer, as well as standards in an equivalent serum matrix, eliminating the need for individual sample recoveries, increasing the sensitivity of the assay and decreasing the requisite sample volume to 0.5 ml of serum. The antibody used for this measurement has 100% cross-reactivity with 1,25(OH)2D3 and 1,25(OH)2D2 and purification of the serum sample does not require HPLC and, in fact, requires only a single cartridge format. Moreover, the antibody does not detect any interfering 1α-hydroxylated vitamin D metabolites. These advances created an assay that is readily adaptable to the commercial clinical laboratory. Indeed, both of these assays are widely available commercially and provide comparable accurate and precise measurements of circulating 1,25(OH)2D. Hence, when necessary, quantitation of this metabolite is reasonably easily obtained.

Chapter 31 New Horizons for Assessment of Vitamin D Status in Man

References 1. Rhodes, C. J., Claridge, P. A., Traffold, D. J. H., and Makin, K. L. J. (1983). An evaluation of the use of Sep-Pak Qg cartridges for the extraction of vitamin D3 and some of its metabolites from plasma and urine. J. Steroid Biochem. 19, 1349–1354. 2. Heubi, J. E., Hollis, B. W., and Tsang, R. C. (1990). Bone disease in chronic childhood cholestasis. II. Better absorption of 25(OH)D than vitamin D in extrahepatic biliary atresia. Pediatr. Res. 27, 26–31. 3. Standing Committee on the Scientific Evaluation of Dietary Reference Intakes, Food and Nutrition Board, Institute of Medicine: Dietary reference intakes for Calcium, Phosphorus, Magnesium, Vitamin D, and Fluoride”. (1997). National Academy Press, Washington, DC. 4. Nongen, M., Ginkel v F. C., Vijgh vd W. J. F., Kuipers, S., Netelenbos, J. C., and Lips, P. (1984). An international comparison of vitamin D metabolite measurements. Clin. Chem. 30, 399–403. 5. Binkley, N., Krueger, D., Cowgill, C., Plum, L., Lake, E., Hansen, K., DeLuca, H., and Drezner, M. (2004). Assay variation confounds the diagnosis of hypovitaminosis D: a call for standardization. J. Clin. Endocrinol. Metab. 89, 3152–3157. 6. Brazerol, W. F., McPhee, A. J., Mimouni, F., Specker, B. L., and Tsang, R. C. (1988). Serial ultraviolet B exposure and serum 25 hydroxyvitamin D response in young adult American blacks and whites: no racial differences. J. Am. College Nut. 7, 111–118. 7. Salamone, L. M., Dallal, G. E., Zantos, D., Makrauer, F., and DawsonHughes, B. (1993). Contributions of vitamin D intake and seasonal sunlight exposure to plasma 25-hydroxyvitamin D concentration in elderly women. Am. J. Clin. Nutr. 58, 80–86. 8. Haddad, J., and Chyu, K. (1971). Competitive protein-binding radioassay for 25-hydroxycholecalciferol. J. Clin. Endocrinol. Metab. 33, 992–995. 9. Goswami, R., Gupta, N., Goswami, D., Marwaha, R. K., Tandon, N., and Kochupillai, N. (2000). Prevalence and significance of low 25-hydroxyvitamin D concentrations in healthy subjects in Delhi. Am. J. Clin. Nutr. 72, 472–475. 10. Outila, T. A., Karkkainen, M. U. M., Seppanen, R. H., and LambergAllardt, C. J. E. (2000). Dietary intake of vitamin D in premenopausal, healthy vegans was insufficient to maintain concentrations of serum 25-hydroxyvitamin D and intact parathyroid hormone within normal ranges during the winter in Finland. J. Am. Dietetic Assoc. 100, 434–440. 11. Guzel, R., Kozanoglu, E., Guler-Uysal, F., Soyupak, S., and Sarpel, T. (2001). Vitamin D status and bone mineral density of veiled and unveiled Turkish women. J. Women’s Health & Gender-Based Med. 10, 765–770. 12. Melin, A., Wilske, J., Ringertz, H., and Saaf, M. (2001). Seasonal variations in serum levels of 25-hydroxyvitamin D and parathyroid hormone but no detectable change in femoral neck bone density in an older population with regular outdoor exposure. J. Am. Geriatric Soc. 49, 1190–1196. 13. Chapuy, M. C., Pamphile, R., Paris, E., Kempf, C., Schlichting, M., Arnaud, S., Garnero, P., and Meunier, P. J. (2002). Combined calcium and vitamin D3 supplementation in elderly women: Confirmation of reversal of secondary hyperparathyroidism and hip fracture risk: The Decalyos II study. Osteoporosis Int. 13, 257–264. 14. Harris, S. S., Soteriades, E., Coolidge, J. A., Mudgal, S., and DawsonHughes, B. (2000). Vitamin D insufficiency and hyperparathyroidism in a low income, multiracial, elderly population. J. Clin. Endocrinol. Metab. 84, 4125–4130. 15. Holick, M. F., Siris, E. S., Binkley, N., Beard, M. K., Khan, A., Katzer, J. T., Petruschke, R. A., Chen, E., and de Papp, A. E. (2005). Prevalence of Vitamin D inadequacy among postmenopausal North American women receiving osteoporosis therapy. J. Clin. Endocrinol. Metab. 90, 3215–3224. 16. Belsey, R., DeLuca, H., and Potts, J. (1971). Competitive binding assay for vitamin D and 25-OH vitamin D. J. Clin. Endocrinol. Metab. 33, 554–557.

525

17. Belsey, R., DeLuca, H., and Potts, J. (1974). A rapid assay for 25-OHvitamin D3 without preparative chromatography. J. Clin. Endocrinol. Metab. 38, 1046–1051. 18. Carter, G. D., Carter, C. R., Gunter, E., Jones, J., Jones, G., Makin, H. L., and Sufi, S. (2004). Measurement of vitamin D metabolites: An international perspective on methodology and clinical interpretation. J. Steroid Biochem. Molec. Biol. 89, 267–271. 19. Lips, P., Chapuy, M., Dawson-Hughes, B., Pols, H., and Holick, M. (1999). An international comparison of serum 25-hydroxyvitamin D measurements. Osteoporosis Int. 9, 394–397. 20. Feleke, Y., Abdulkadir, J., Mshana, R., Mekbib, T., Brunvand, L., Berg, J., and Falch, J. (1994). Low levels of serum cacidiol in an African population compared to a North European population. Eur. J. Endocrinol. 141, 358–360. 21. Heaney, R., Dowell, S., Hale, C., and Brendich, A. (2003). Calcium absorption varies within the reference range for serum 25-hydroxyvitamin D. J. Am. Coll. Nutr. 22, 142–146. 22. Lester, E., Skinner, R. K., and Wills, M. R. (1977). Seasonal variation in serum 25-hydroxyvitamin D in the elderly in Britian. Lancet 1, 979–998. 23. Dawson-Hughes, B., Dallal, G. E., Krall, E. A., Harris, S., Sokoll, L. J., and Falconer, G. (1991). Effect of vitamin D supplementation on wintertime and overall bone loss in healthy postmenopausal women. Ann. Intern. Med. 115, 505–512. 24. Krall, E. A., Sayhoun, N., Tannenbaum, S., Dallal, G. E., and DawsonHughes, B. (1989). Effect of vitamin D intake on seasonal variations in parathyroid hormone secretion in postmenopausal women. N. Engl. J. Med. 321, 1777–1783. 25. Webb, A. R., Pilbeam, C., Hanafin, N., and Holick, M. F. (1990). An evaluation of the relative contributions of exposure to sunlight and of diet to the circulating concentrations of 25-hydroxyvitamin D in an elderly nursing home population in Boston. Am. J. Clin. Nutr. 51, 1075–1081. 26. Lund, B., and Sorensen, O. H. (1979). Measurement of 25-hydroxyvitamin D in serum and its relation to sunshine, age and vitamin D intake in the Danish population. Scandanavian J. Clin. Invest. 39, 23–30. 27. Poskitt, E. M. E., Cole, T. J., and Lawson, D. E. M. (1979). Diet, sunlight and 25-hydroxyvitamin D in healthy children and adults. B.M.J. 1, 221–223. 28. Rejnmark, L., Jorgensen, M. E., Pedersen, M. B., Hansen, J. C., Heickendorff, L., Lauridsen, A. L., Mulvad, G., Siggaard, C., Skjoldborg, H., Sorensen, T. B., Pedersen, E. B., and Mosekilde, L. (2004). Vitamin D insufficiency in Greenlanders on a westernized fare: Ethnic differences in calcitropic hormones between Greenlanders and Danes. Calcif. Tissue Int. 74, 255–263. 29. Holick, M. F. (2003). Vitamin D: A millennium perspective. J. Cell. Biochem. 88, 296–307. 30. Holick, M. F. (1985). The photobiology of vitamin D and its consequences for humans. Ann. N. Y. Acad. Sci. 453, 1–13. 31. Holick, M. F., MacLaughlin, J. A., and Clark, M. B. (1982). Photosynthesis of previtamin D3 in human skin. Science 216, 1001–1003. 32. Holick, M. F. (1995). Environmental factors that influence the cutaneous production of vitamin D. Am. J. Clin. Nutr. 61, 638S–645S. 33. Jablonski, N. G. (2004). The evolution of human skin and skin color. Ann. Rev. Anthropol. 33, 585–623. 34. Rajakumar, K. (2003). Vitamin D, cod-liver oil, sunlight and rickets: A historical perspective. Pediatrics 112, e132–e135. 35. Hollis, B. W. (2005). Circulating 25-hydroxyvitamin D levels indicative of vitamin D sufficiency: Implications for establishing a new effective dietary intake recommendation for vitamin D. J. Nutrition 135, 317–322. 36. Better, O. S., Shabtai, M., Kedar, S., Melamud, A., Berenheim, J., and Chaimovitz, C. (1980). Increased incidence of nephrolithiasis (N) in lifeguards (LG) in Israel. Adv. Exp. Med. Biol. 128, 467–472.

526 37. Tangpricha, V., Turner, A., Spina, C., Decastro, S., Chen, T. C., and Holick, M. F. (2004). Tanning is associated with optimal vitamin D status (serum 25-hydroxyvitamin D concentration) and higher bone mineral density. Am. J. Clin. Nutr. 80, 1645–1649. 38. Haddock, L., Corcino, J., and Vazques, M. D. (1982). 25(OH)D serum levels in the normal Puerto Rican population and in subjects with tropical sprue and parathyroid disease. Puerto Rico Health Sci. J. 1, 85–91. 39. Schimidt-Gayk, H. (1999). Measurement of calcium, phosphate, parathyroid hormone and vitamin D. In “Dynamics of Bone and Cartilage Metabolism” (M. Seibel, S. Robins and J. Bilezikian, eds.), pp. 375–399. Academic Press, San Diego, CA. 40. Chupuy, M., Preziosi, P., Maamer, M., Arnaud, S., Galan, P., Hercberg, S., and Meunier, P. (1997). Prevalence of vitamin D insufficiency in an adult normal population. Osteoporosis Int. 7, 439–443. 41. Netelenbos, J., Jongen, M., van der Vijgh, W., Lips, P., and van Ginkel, F. (1985). Vitamin D status in urinary calcium stone formation. Arch. Intern. Med. 145, 681–685. 42. Wielen, v. R., Lowik, M., Berg, v. H., de Groot, L. C., Haller, J., Moreiras, O., and Staveren, v. W. (1995). Serum vitamin D concentrations among elderly people in Europe. Lancet 346, 207–210. 43. van der Wielen, R. P., Lowik, M. R., van den Berg, H., de Groot, L. C., Haller, J., Moreiras, O., and van Staveren, W. A. (1995). Serum vitamin D concentrations among elderly people in Europe. Lancet 346, 207–210. 44. Zhao, X. H. (1992). Nutritional situation of Beijing residents. Southeast Asian J. Trop. Med. Pub. Health 23(Suppl 3), 65–68. 45. McKenna, M. (1992). Differences in vitamin D status between countries in young adults and the elderly. Am. J. Med. 93, 69–77. 46. Lips, P., Duong, T., Oleksik, A., Black, D., Cummings, S., Cox, D., and Nickelsen, T. (2001). A global study of vitamin D status and parathyroid function in postmenopausal women with osteoporosis: Baeline data from the Multiple Outcomes of Raloxifene Evaluation Clinical Trial. J. Clin. Endocrinol. Metab. 86, 1212–1221. 47. Thomas, M. K., Lloyd-Jones, D. M., Thadhani, R. I., Shaw, A. C., Deraska, D. J., Kitch, B. T., Vamvakas, E. C., Dick, I. M., Prince, R. L., and Finkelstein, J. S. (1998). Hypovitaminosis D in medical inpatients. N. Engl. J. Med. 338, 777. 48. Dawson-Hughes, B., Harris, S. S., and Dallal, G. E. (1997). Plasma calcidiol, season, and serum parathyroid hormone concentrations in healthy elderly men and women. Am. J. Clin. Nutr. 65, 67–71. 49. Lips, P., Wiersinga, A., van Ginkel, F. C., Jongen, M. J. M., Netelenbos, J. C., Hackeng, W. H. L., Delmas, P. D., and van der Vijgh, W. J. F. (1988). The effect of vitamin D supplementation on vitamin D status and parathyroid function in elderly subjects. J. Clin. Endocrinol. Metab. 67, 644–650. 50. Ooms, M. E., Lips, P., Roos, J. C., van der Vijgh, W. J., Popp-Snijders, C., Bezemer, P. D., and Bouter, L. M. (1995). Vitamin D status and sex hormone binding globulin: determinants of bone turnover and bone mineral density in elderly women. J. Bone Miner. Res. 10, 1177–1184. 51. Lips, P., Pluijm, S. M. F., Popp-Snijders, C., and Smit, J. H. (2001). Vitamin D status, sex hormone binding globulin, IGF-1 and markers of bone turnover as determinants of bone mass and fractures in the Longitudinal Aging Study Amsterdam. J. Bone Miner. Res. 16(Suppl 1), S166. 52. Trivedi, D. P., Doll, R., and Khaw, K. T. (2003). Effect of four monthly oral vitamin D3 (cholecalciferol) supplementation on fractures and mortality in men and women living in the community: randmized double blind controlled trial. BMJ 326, 469–474. 53. Scotti, A., Bianchini, C., Gianalfredo, A., and Marzo, A. (2001). Absorption of calcium administered alone or in fixed combination with vitamin D to healthy volunteers. Arzneim-Forsch/Drug Res. 51, 493–500. 54. Bischoff-Ferrari, H. A., Dietrich, T., Orav, E. J., and Dawson-Hughes, B. (2004). Positive association between 25-hydroxy vitamin D levels and bone mineral density: a population-based study of younger and older adults. Am. J. Med. 116, 634–639.

GARY L. LENSMEYER ET AL. 55. Lips, P. (2005). How to define normal values for serum concentrations of 25-hydroxyvitamin D? An overview. In (D. Feldman, J. W. Pike, and F. H. Glorieux, eds.), “Vitamin D” pp. 1019–1027. Elsevier Academic Press, Amsterdam. 56. Heaney, R. P. (2004). Functional indices of vitamin D status and ramifications of vitamin D deficiency. Am. J. Clin. Nutr. 80(6 Suppl), 1706S–1709S. 57. Dawson-Hughes, B., Heaney, R. P., Holick, M. F., Lips, P., Meunier, P. J., and Vieth, R. (2005). Estimates of optimal vitamin D status. Osteoporosis Int. 16, 713–716. 58. Peacock, M. (1998). Effects of calcium and vitamin D insufficiency on the skeleton. Osteoporosis Int. 8, 545–551. 59. Glendenning, P., Noble, J. M., Taranto, M., Musk, A. A., McGuniess, M., Goldswain, P. R., Fraser, W. D., and Vasikaran, S. D. (2003). Issues of methodology, standardization and metabolite recognition for 25hydroxyvitamin D when comparing the DiaSorin radioimmunoassay and the Nichols Advantage automated chemiluminescence proteinbinding assay in hip fracture cases. Ann. Clin. Biochem. 40, 546–551. 60. Dean, B., Kolavcic, M., Wark, J., and Harrison, L. (1988). Chromatography of serum on Sep-pak C18 corrects falsely elevated vitamin D metabolite levels measured by protein binding assay. Clin. Chim. Acta. 176, 169–178. 61. Hollis, B. W., and Napoli, J. (1985). Improved radioimmunoassay for vitamin D and its use in assessing vitamin D status. Clin. Chem. 31, 1815–1819. 62. Hollis, B. W., Kamerud, J., Selvaag, S., Lorenz, J., and Napoli, J. (1993). Determination of vitamin D status by radioimmunoassay with an 125I-labeled tracer. Clin. Chem. 39, 529–533. 63. Marks, V. (2002). False-positive immunoassays results: A multicenter survey of erroneous immunoassay results from assays of 74 analytes in 10 donors from 66 laboratories in seven countries. Clin. Chem. 48, 2008–2016. 64. Stenman, U.-H. (2001). Immunoassay standardization: Is it possible, Who is responsible, Who is capable? Clin. Chem. 47, 815–820. 65. Bouillon, R., Van Herck, E., Jans, I., Tan, B. K., Van Baelen, H., and DeMoor, P. (1984). Two direct (nonchromatographic) assays for 25-hydroxyvitamin D. Clin. Chem. 30, 1731–1736. 66. Hollis, B. W. (2005). Detection of vitamin D and its major metabolites. In (D. Feldman, J. Pike and F. Glorieux, eds.), “Vitamin D” pp. 931–950. Elsevier Academic Press, Amsterdam. 67. Dorantes, L., Arnaud, S., and Arnaud, C. (1978). Importance of the isolation of 25-hydroxyvitamin D before assay. J. Lab. Clin. Med. 91, 791–796. 68. Hollis, B. W. (2004). The determination of circulating 25-hydroxyvitamin D: No easy task. J. Clin. Endocrinol. Metab. 89, 3149–3151. 69. Gemar, D., Binkley, N., Krueger, D., Engelke, J., and Drezner, M. (2004). Assessment of vitamin D status: an imposing dilemma. J. Bone Miner. Res. 19, Suppl 1, S343. 70. Ersfeld, D., Rao, D., Body, J., Sackrison, J., Jr., Miller, A., Parikh, N., Eskridge, T., Polinske, A., Olson, G., and MacFarlane, G. (2004). Analytical and clinical validation of the 25 OH vitamin D assay for the LIAISON automated analyzer. Clin. Biochem. 37, 867–874. 71. Turpeinen, U., Hohenthal, U., and Stenman, U.-H. (2003). Determination of 25-hydroxyvitamin D in serum by HPLC and immunoassay. Clin. Chem. 49, 1521–1524. 72. Vieth, R., Chan, A., and Pollard, A. (1995). 125I-RIA kit cannot distinguish vitamin D deficiency as well as a more specific assay for 25-hydroxyvitamin D. Clin. Biochem. 28, 175–179. 73. Hollis, B. W. (2000). Comparison of commercially available 125I-based RIA methods for the determination of circulating 25(OH)D. Clin. Chem. 46, 1657–1661. 74. Eisman, J. A., Shepard, R. M., and DeLuca, H. F. (1977). Determination of 25-hydroxyvitamin D2 and 25-hydroxyvitamin D3 in human plasma using high-pressure liquid chromatography. Anal. Biochem. 90, 298–305. 75. Gilbertson, T. J., and Stryd, R. P. (1977). High-performance liquid chromatographic assay for 25-hydroxyvitamin D3 in serum. Clin. Chem. 23, 1700–1704.

Chapter 31 New Horizons for Assessment of Vitamin D Status in Man 76. Jones, G. (1978). Assay of vitamins D2 and D3, and 25-hydroxyvitamin D2 and D3 in human plasma by high-performace liquid chromatography. Clin. Chem. 24, 287–298. 77. Aksnes, L. (1994). Simultaneous determination of retinol, α-tocopherol, and 25-hydroxyvitamin D inhuman serum by high-performance liquid chromatography. J. Pediatr. Gastrenterol. Nutr. 18, 339–343. 78. Shimada, K., Mitamura, K., Kitama, N., and Kawasaki, M. (1997). Determination of 25-hydroxyvitamin D3 in human plasma by reversedphase high-performance liquid chromatography with ultraviolet detection. J. Chromatogr. 689, 409–414. 79. Alvarez, J. C., and De Mazancourt, P. (2001). Rapid and sensitive high-performance liquid chromatographic method for simultaneous determination of retinol, α-tocopherol, 25-hydroxyvitamin D3 and 25-hydroxyvitamin D2 in human plasma with photodiode-array ultraviolet detection. J. Chromatogr. 755, 129–135. 80. Campbell, J. A., Squires, D. M., and Babcock, J. C. (1969). Synthesis of 25-hydroxycholecalciferol, the biologically effective metabolite of vitamin D3. Steroids 13, 567–577. 81. De Leenheer, A. P., and Cruyl, A. A. (1978). Vitamin D3 in plasma: quantitation by mass fragmentography. Anal. Biochem. 91, 293–303. 82. Coldwell, R. D., Trafford, D. J., Varley, M. J., Kirk, D. N., and Makin, H. L. (1989). Measurement of 25-hydroxyvitamin D2, 25-hydroxyvitamin D3, 24,25-dihydroxyvitamin D2 and 25, 26-dihydroxyvitamin D2 in a single plasma sample by mass fragmentography. Clin. Chim. Acta. 28, 157–168. 83. Björkhem, I., and Holmberg, I. (1980). Mass fragmentographic assay of 25-hydroxyvitamin-D3. Methods Enzymol 67, 385–393. 84. Kobold, U., Jeppsson, J. O., Dulffer, T., Finke, A., Hoelzel, W., and Miedema, K. (1997). Candidate reference methods for hemoglobin A1c. Clin. Chem. 43, 1944–1951. 85. Jeppsson, J. O., Kobold, U., Barr, J., Finke, A., Hoelzel, W., Hoshino, T., Miedema, K., Mosca, A., Mauri, P., Paroni, R., Thienpont, L., Umemoto, M., and Weykamp, C. (2002). Approved IFCC reference method for the measurement of HbA1c in human blood. Clin. Chem. Lab. Med. 40, 78–89. 86. Vogeser, M., Kyriatsoulis, A., Huber, E., and Kobold, U. (2004). Candidate reference method for the quantification of circulating

87.

88. 89.

90.

91.

92.

93.

94.

95.

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25-hydroxyvitamin d3 by liquid chromatography-tandem mass spectrometry. Clin. Chem. 50, 1415–1417. Kamao, M., Hatakeyama, S., Sakaki, T., Sawada, N., Inouye, K., Kubodera, N., Reddy, G. S., and Okano, T. (2005). Measurement and characterization of C-3 epimerization activity toward vitamin D3. Arch. Biochem. Biophys. 436, 196–205. Hollis, B. W. (2005). Personal Communication. Higashi, T., Awad, D., and Shimada, K. (2001). Simultanous determination of 25-hydroxyvitamin D2 and 25-hydroxyvitamin D3 in human plasma by liquid chromatography-tandem mass spectrometry employing derivatization with a Cookson-Type reagent. Biol. Pharm. Bull. 24, 738–743. Lensmeyer, G. L., Wiebe, D. A., Binkley, N., Drezner, M. K., Singh, R., and Darcy, T. P. (2005). Clinically reliable assay for routine monitoring of 25-hydroxyvitamin D3 and 25-hydroxyvitamin D2 in serum using a semi-automated extraction and HPLC. Clin. Chem. 51, Suppl, A. Fenske, J. S., Peieper, K. A., Belisle, K. J., Eastvold, M., and Singh, R. J. (2005). LC-MS/MS analysis of 25 OH Vitamin D2 and D3 compared to the DiaSorin LIAISION® and RIA methods. Clin. Chem. 51, Suppl, A114. Carter, G. D., Carter, R., Jones, J., and Berry, J. (2004). How accurate are assays for 25-hydroxyvitamin D? Data from the international vitamin D external quality assessment scheme. Clin. Chem. 50, 2195–2197. Reinhardt, T. A., Horst, R. L., Orf, J. W., and Hollis, B. W. (1984). A microassay for 1,25-dihydroxyvitamin D not requiring high performance liquid chromatography: Application to clinical studies. J. Clin. Endocrinol. Metab. 58, 91–98. Hollis, B. W. (1986). Assay of circulating 1,25-dihydroxyvitamin D involving a novel single-cartridge extraction and purification procedure. Clin. Chem. 32, 2060–2063. Hollis, B. W., Kamerud, J. Q., Kurkowski, A., Beaulieu, J., and Napoli, J. L. (1996). Quantification of circulating 1,25-dihydroxyvitamin D by radioimmunoassay with an 125I-labeled tracer. Clin. Chem. 42, 586–592.

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Chapter 32

Measurement of Biochemical Markers of Bone Formation Kim E. Naylor, Richard Eastell

I. II. III. IV.

From the Academic Unit of Bone Metabolism, University of Sheffield, Sheffield, UK

V. Bone Alkaline Phosphatase VI. Osteocalcin VII. Discussion References

Abstract Introduction Propeptides of Type I Procollagen Total Alkaline Phosphatase

I. ABSTRACT

which are usually measured in urine. There are many commercially available and in-house assays for the measurement of biochemical markers of bone formation. Established assays have now been adapted for measurement on automated systems potentially improving precision, accuracy, and sample processing time.

There are several biochemical markers of bone formation. These markers do not always behave in identical ways as each is reflecting a different stage of osteoblast differentiation, each has different pathways for clearance, and each has different degrees of specificity for bone. It is important to measure the bone formation marker that is best able to provide the information that is required for a specific investigation. The measurement of biochemical markers of bone formation provides a noninvasive tool to assess changes in bone metabolism. They are most useful when bone formation is of specific interest, such as monitoring anabolic treatments for osteoporosis. Measurements can be done repeatedly to investigate the time course of change over a relatively short time period in comparison to other methods. There are several sources of preanalytical and analytical variability that should be minimized to obtain optimum results. Bone formation markers are measured in serum and tend to be less variable than bone resorption markers, Dynamics of Bone and Cartilage Metabolism

II. INTRODUCTION Biochemical markers of bone formation are all proteins secreted by the osteoblasts (Table I). They are not all specific to the osteoblast and this is one of the reasons why bone formation markers do not always behave in the same way (Table II). Each bone formation marker reflects a different stage of osteoblast differentiation [1–3]. The proliferation period is associated with expression of collagen type I genes, suggesting that the propeptides of type I procollagen (C terminal PICP and N terminal PINP) would be markers of this phase of bone formation. Bone alkaline phosphatase (bone ALP) is expressed at the matrix 529

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Table I.

Biochemical Markers of Bone Formation

Propeptides of Type I procollagen Total alkaline phosphatase Bone alkaline phosphatase Osteocalcin

C terminal PICP N terminal PINP Total ALP Bone ALP OC

maturation phase and osteocalcin (OC) is produced at the mineralization phase of bone formation [4]. Biochemical markers of bone formation are cleared by different mechanisms, e.g. liver, kidney, or both, and have different halflives ranging from minutes (PINP) to days (bone ALP). The use of bone formation markers is most informative when we specifically want to study bone formation; for example, studying acute decreases in bone formation, such as the effects of glucocorticoids, or the increase in bone formation such as the effect of anabolic treatments used for osteoporosis or the evaluation of bone metastases (osteosclerotic). Bone formation markers may also be useful to study states of high bone remodeling or changes in bone remodeling that result from antiresorptive treatment. The reason that they may be useful in these situations is that bone formation markers may be more bone specific or less variable than bone resorption markers as they are measured in serum, unlike the bone resorption markers which are typically measured in urine and, hence, have the added variability from a creatinine correction to the marker measurement. It may be interesting to measure bone formation markers along with bone resorption markers as it gives an index of bone balance [5]. Bone density measurements are a valuable tool for the diagnosis of metabolic bone disease such as osteoporosis. A drawback with this method is that changes are relatively slow and small in comparison to the variability of the measurement. Repeated measurements have to be made over time intervals of at least 12 months to detect a true change in bone density which is greater than the error of the measurement itself. Bone density measurements are therefore not the best technique to employ if monitoring change in bone metabolism such as in response to treatment (Table III). Biochemical markers of bone turnover offer an alternative method of assessing bone metabolism, Table II.

Response of Bone Formation Markers

Reasons why bone formation markers do not all respond in the same way Reflect different phases of differentiation of the osteoblast May not be specific to bone Clearance mechanisms differ Change in marker may not represent osteoblast activity, but a specific effect on gene transcription (or translation)

Table III. How to Monitor Treatment? Bone mineral density

Bone turnover markers

This test is used for the diagnosis of osteoporosis Precise Slow to change Change is small Unlikely to achieve a value with treatment in the normal range Not a good surrogate What would happen if treatment were not given?

This test is not used for the diagnosis of osteoporosis Imprecise Quick to change Change is large Likely to achieve a value with treatment in the normal range Good surrogate No change if no treatment

which is relatively inexpensive, noninvasive, requires no special patient preparation, and can be repeated over short time intervals providing quick results. For optimum measurement results, it is important to identify and minimize sources of preanalytical variability (Tables IV and V). Bone markers are quick to change in response to therapy and the change is usually large, achieving a value within the normal range in response to treatment. The introduction of automated assays has greatly enhanced the accessibility and sample throughput of bone marker assays.

III. PROPEPTIDES OF TYPE I PROCOLLAGEN A. General Description Type I collagen accounts for approximately 90% of the bone matrix, the remainder being noncollagenous proteins. The biosynthesis of type I collagen is a complex process involving both intracellular and extracellular modifications. During type I collagen biosynthesis the carboxy (PICP) and amino (PINP) terminal propeptides are released and can be measured in serum as markers of collagen formation (see Chapter 24).

B. Specificity and Clearance The propeptides of type I procollagen have a short halflife, PICP 6–9 minutes and PINP 1 minute as determined in the rat model. Both PICP and PINP are cleared by the liver endothelial cells but by different mechanisms; PICP by the mannose receptors and PINP by scavenger receptors [6, 7]. The clearance of PICP is sensitive to hormonal changes, which should be considered when measured in patients during puberty and the menopause or those taking hormone replacement therapy or aromatase inhibitors. Both PINP and PICP show a dose-dependent decrease in

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Chapter 32 Measurement of Biochemical Markers of Bone Formation Table IV. Source Age Gender Pregnancy and lactation Fracture Disease Drugs Immobility

Uncontrollable Sources of Preanalytical Variability

Action Use appropriate reference range Use appropriate reference range Appropriate interpretation of result Record details of previous fracture Medical history should be recorded Record details of all medication Record history of immobility

Note Adolescents should be based on pubertal status Men and women (pre- and post-menopausal reference ranges) Weeks of gestation or postpartum Date, site, and type of fracture Important: renal impairment, liver disease, diabetes, thyroid disease Important: corticosteroids, anticonvulsants, heparin, GnRH agonists Date and duration

Adapted from reference [52].

response to treatment with a novel selective estrogen receptor modulator (SERM) [8]. As the procollagen propeptides are cleared by the liver, they are elevated in patients with liver disease and in some medical conditions it is therefore not appropriate to measure them. For example, in a study of patients with primary bilary cirrhosis the collagen-related markers of bone turnover reflect severity of liver fibrosis and do not reflect bone turnover [9]. As type I collagen is the most abundant type of collagen in the body it is possible that some of the procollagen propeptides present in serum could be from nonbone sources [10]. Bone is, however, a metabolically active tissue with higher turnover than soft tissues and type I procollagen peptides should therefore mainly reflect type I bone collagen formation. In a study of untreated postmenopausal osteoporotic women, PINP had significantly higher diagnostic accuracy to distinguish between osteoporotic women and controls, compared to PICP [11].

C. Assays Several assays are available for the measurement of propeptides of type I procollagen including RIA and ELISA methods. The PINP assay has been adapted for measurement by autoanalyzer (Roche Diagnostics, Germany). Assays are available to measure both the highand low-molecular-weight forms of PINP [12–14]. It is

proposed that the high-molecular-weight form of PINP represents a trimeric form which is unstable at body temperature and thermal transition produces a monomeric α1 chain form of PINP [14]. The ratio of the two molecular weight forms could be different in different metabolic bone diseases or could change in response to therapy.

D. Variability 1. CONTROLLABLE

PICP has a circadian rhythm with a nocturnal peak and an afternoon nadir. There does not appear to be a seasonal variation in levels of PICP. Woitge et al. reported that, like other biochemical markers of bone formation, PICP was higher in the winter compared to summer levels in women, but this was not statistically significant [15]. Blumsohn et al. [16] reported a small increase in bone formation markers during the summer months but concluded that this was unlikely to be of clinical significance. Serum PICP is lower in endurance-trained athletes compared to sedentary controls, but other bone formation markers were not significantly different between the two groups [17, 18]. 2. UNCONTROLLABLE

Bone turnover is high during growth in children, particularly in adolescence. PICP and, to a greater extent, PINP are higher in children compared to adults with an increase

Table V. Controllable Sources of Preanalytical Variability Source Circadian Fasting Diet Seasonal Exercise

Adapted from reference [52].

Action Collect serial samples at same time of day Collect samples after overnight fast Record details of calcium supplementation Consider sample collection time of year for large longitudinal studies Record details of exercise

Note Define collection time window in protocol Calcium may suppress bone turnover Not important for individuals Refrain from intensive exercise 24 hours before sample collection

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during puberty [19]. Bone formation markers are elevated following fracture. In a study of Colles’ fracture PICP was significantly elevated up to 5 weeks following fracture, returning to baseline at 9 months post fracture [20]. In a study of patients with ankle fracture bone formation markers were increased significantly between 1 and 4 weeks following fracture and PINP remained elevated at 1 year post fracture [21]. This was also found to be the case for patients with distal forearm fracture [22].

E. Evaluating Change in Bone Formation 1. GLUCOCORTICOIDS

Glucocorticoids are known to inhibit bone formation. In vitro work has shown that supraphysiological doses of glucocorticoids directly inhibit osteoblast function through their effect on type I collagen synthesis [23]. Cooper et al. reported bone formation markers to be decreased in men taking glucocorticoids but bone resorption markers were unchanged (Fig. 1) [24]. 2. OSTEOGENESIS IMPERFECTA

As the propeptides of type I procollagen are markers of bone collagen formation, they may be useful in diseases of impaired collagen formation such as osteogenesis imperfecta, in which condition they are often decreased [25].

have abnormal baseline PINP, with a significant decrease following treatment [26]. Others have reported bone formation markers to be sensitive in monitoring the response to therapy in Paget’s disease of bone with a decrease of the same order of magnitude as seen for bone resorption markers (Fig. 2) [27]. 2. ESTROGEN

In menopausal women treated with estrogen therapy there is a more dynamic response in PINP compared to PICP [28]. Peris et al. reported bone formation markers to be elevated in women following surgical menopause, PINP having the largest magnitude of change with an increase of 101% compared to an increase of 44% for bone ALP and 37% for osteocalcin [29]. During pregnancy, PICP and PINP are significantly increased during the third trimester by 35% and 66%, respectively, compared to prepregnancy baseline values [30]. 3. RENAL DISEASE

Serum PINP is elevated compared to healthy controls in patients with end-stage renal disease and renal transplant patients [31]. A recent study of patients with chronic renal failure found PINP to be significantly related to decrease in BMD of the radius [32].

F. Evaluating Change in Bone Remodeling 1. ANTIRESORPTIVE THERAPY

Bone formation markers are sensitive to changes in bone turnover that result from antiresorptive therapy. In a study of the response to bisphosphonate therapy in Paget’s disease of bone, more than 95% of patients were found to

Figure 1

The effect of 5 mg prednisolone twice daily on biochemical bone markers in healthy males. Data are percentage change from baseline and values are mean ± SE, n = 20 [24]. Reproduced with permission from Cooper et al., J. Clin Endocrinol Metab 2003; 88:3874–7[24].

Figure 2 Mean percentage (± SEM) of variation of biochemical markers of bone turnover in relation to baseline values in patients with Paget’s disease at 1 and 6 months from the end of treatment with tiludronate (#significant difference compared with other markers of the group). Reproduced with permission from reference [27].

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The propeptides of type I procollagen may prove to be useful to assess bone involvement in cancer patients. In a study of prostate cancer patients, those with bone metastases had elevated PINP compared to those with no metastases [33]. Jung et al. have also reported PINP, along with other bone turnover markers, to be significantly increased in prostate cancer patients that have bone metastases compared to patients with no bone involvement [34]. In a study of patients with ovarian cancer, a low PICP to PINP ratio was associated with fast progression of disease and poor clinical outcome [35].

IV. TOTAL ALKALINE PHOSPHATASE A. General Description The alkaline phosphatase enzymes hydrolyze organic phosphates at a high pH. In adults the majority of circulating alkaline phosphatase is derived from bone and liver. In metabolic bone disease with increased osteoblast activity the bone fraction is elevated.

B. Specificity and Clearance Total ALP lacks sensitivity and specificity as a bone formation marker due to the contribution of alkaline phosphates from nonskeletal sources. Total ALP is not as sensitive as specific bone formation markers in diseases with subtle changes in bone turnover such as osteoporosis [11]. However, for bone disease with dramatic changes in bone metabolism such as Paget’s disease of bone, total ALP provides adequate information on changes in bone turnover. Total ALP is cleared by the liver and is elevated in patients with liver disease with involvement of the biliary tracts.

C. Assays The measurement of serum total alkaline phosphatase (total ALP) has been validated against bone mineralization measured by calcium kinetic methods [36]. It is usually measured by an automated colorimetric method, which is relatively inexpensive with good reproducibility.

D. Variability 1. CONTROLLABLE

The circadian rhythm of total ALP is small and has no significant effect. Changes during the menstrual cycle are

not significant to the measurement of total ALP. There does not appear to be a clinically significant seasonal effect on total ALP. 2. UNCONTROLLABLE

Due to the presence of nonbone isoforms of alkaline phosphatase, total ALP is elevated in certain medical conditions including liver disease, pregnancy, and malignancy. Total ALP is low in some diseases such as hypophosphatasia, hypothyroidism, celiac disease, and in patients treated with glucocorticoid therapy. In some cases the cause of elevated total ALP is not apparent and electrophoresis methods to separate different isoforms may be useful. Low total ALP can be caused by arrested bone growth as occurs, for example, in achondroplasia. Hypophosphatasia is also a cause of low total ALP and this can be genetic or due to low magnesium levels associated with celiac disease.

E. Evaluating Change in Bone Remodeling Total ALP is elevated in bone disease where there is a dramatic change in bone turnover such as Paget’s disease of bone, osteomalacia, and primary hyperparathyroidism, but is less reliable to identify the subtle changes in bone turnover that occur in osteoporosis [11, 37]. Total ALP decreases in response to antiresorptive treatment in Paget’s disease of bone [26, 27].

V. BONE ALKALINE PHOSPHATASE A. General Description Circulating total ALP consists of several isoenzymes of alkaline phosphatase, which differ by their post-translational modification, providing a basis for their analytical separation (see also Chapter 9). There are four genes that code for alkaline phosphatase. The genes that code for germ cell, placental, and intestinal isoforms are located on chromosome 2. The tissue nonspecific alkaline phosphatase gene codes for liver, kidney, and bone alkaline phosphatase and is located on chromosome 1 [38].

B. Specificity and Clearance Bone alkaline phosphatase has a half-life of around 40 hours and like other glycoproteins is cleared by the liver. Immunoassays for bone alkaline phosphatase (bone ALP) show greater bone specificity than total alkaline phosphatase. In certain conditions there may be a possibility of cross-reactivity from other isoforms of alkaline

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phosphatase such as liver and placental isoenzyme, for example, in patients with liver disease, or during pregnancy, respectively.

C. Assays Several techniques have been adopted to quantify liver and bone isoenzymes including heat inactivation, chemical inhibition, electrophoresis, wheat germ lectin precipitation, and immunoassays [39]. The heat inactivation method utilizes the half-inactivation time of alkaline phosphatase isoenzymes at 56⬚C [40]. The electrophoresis method separates on the basis that the liver isoform is more negatively charged than the bone isoform. Electrophoresis allows the accurate quantitation of the various isoforms of alkaline phosphatase with selective inhibition (e.g. with neuraminidase or wheat germ lectin) improving separation [41]. The wheat germ lectin precipitation method separates the bone from the liver isoenzyme as wheat germ lectin preferentially binds to the N-acetylglucosamine residues on the bone isoenzyme. The liver isoenzyme remaining in the supernatant is subtracted from total enzyme activity to give the bone alkaline phosphatase result. A drawback with this technique is the variable affinity of wheat germ lectin for the bone isoenzyme and it is, therefore, important to standardize the assay. Immunoassays provide a rapid and convenient alternative to other measurement methods. They have been developed to utilize monoclonal antibodies directed at both the liver [42] and the bone isoforms [43–46]. Automated immunoassays for bone ALP have now been developed.

D. Variability 1. CONTROLLABLE

Alkaline phosphatase has a low day-to-day variability, with a half-life of around 40 hours [47]. Bone ALP exhibits a circadian rhythm, although there are conflicting results as to the pattern with some reports of two peaks [48] and others reporting a single nocturnal peak [49]. Like other bone formation markers, bone ALP is elevated during the luteal phase of the menstrual cycle [48]. However, this increase is in the order of 10% and not regarded as clinically significant. Studies have conflicting results with regard to seasonal variability. Some report alkaline phosphatases to be lower in the summer compared to winter [50] whereas others report a decrease in the winter [51]. These different findings may be due to the study design as some are cross sectional and others longitudinal, but also may be due to the geography of the study site [52].

2. UNCONTROLLABLE

Bone ALP is affected by age, gender, and hormonal status [41]. In children, bone ALP correlates with height velocity and weight gain until puberty when there is a dramatic increase [19, 53]. During pregnancy bone ALP is significantly elevated during the third trimester [30, 54, 55] in comparison to nonpregnant, nonlactating levels. It is important that assays used to measure bone alkaline phosphatase during pregnancy do not cross-react with the placental isoform. Fracture causes an increase in bone formation markers, which is sustained for months following the injury. Following fracture there was a significant increase in total and bone alkaline phosphatase [56, 57], which was maintained during the 20-week study period. Bone ALP was also elevated in patients with fractures of the distal forearm and remained elevated at 1 year after fracture [22].

E. Evaluating Change in Bone Formation Bone ALP measured by immunoassay is more sensitive than total ALP for clinical investigation of patients with metabolic bone disease [27, 43–45]. However, despite this improved sensitivity it has been argued that total ALP provides sufficient diagnostic information for most clinical situations, particularly in more active bone disease [27, 58].

F. Evaluating Change in Bone Remodeling 1. ANTIRESORPTIVE THERAPY

Bone ALP decreases in response to alendronate treatment in postmenopausal osteoporosis [59, 60]. After treatment for between 8–12 weeks bone ALP had decreased to within the premenopausal reference range [60]. In patients with Paget’s disease of bone both total ALP and bone ALP decrease in response to bisphosphonate therapy [26, 27]. 2. ESTROGEN

Serum estradiol is inversely related to bone ALP in a population based study of postmenopausal women [61]. Bone ALP has been used to evaluate the third-generation aromatase inhibitors such as anastrazole used in the treatment of breast cancer. These reduce circulating estrogen which may affect bone turnover, resulting in increased fracture risk [62]. 3. MYELOMA

Bone markers may have a role in the assessment of patients with myeloma-induced bone disease. Specific markers may be used as noninvasive measures of

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Chapter 32 Measurement of Biochemical Markers of Bone Formation

bone involvement. One study suggests that serum bone ALP reflects suppressed bone formation rate and may be useful to differentiate between benign and myelomainduced osteoporosis [63]. However, other data showed bone formation markers to be less informative whereas bone resorption markers were predictive of progressive bone events [64].

VI. OSTEOCALCIN A. General Description 1. INTACT AND LARGE N-TERMINAL MID-FRAGMENT OSTEOCALCIN

Osteocalcin or bone Gla protein is an established marker of bone formation. It is a vitamin K-dependent protein produced by osteoblasts [65]. Osteocalcin accounts for up to 20% of the noncollagenous protein in bone, with an affinity for bone mineral; however, its function is unknown. Circulating osteocalcin is heterogeneous: in addition to the intact molecule there are several osteocalcin fragment molecules, which are the result of proteolyic hydrolysis of peptide bonds involving arginine residues [66, 67]. Garnero et al. [68] characterized several immunoreactive forms of human osteocalcin using monoclonal antibodies (Fig. 3). The large N-terminal mid-fragment (1-43) accounted for a high proportion of circulating total osteocalcin in healthy subjects and those with osteoporosis. As osteocalcin is incorporated into the bone matrix, it is feasible that some osteocalcin may be released into the circulation due to bone resorption. However, levels of the large N-terminal

mid-fragment are not altered by bisphosphonate therapy in patients with metabolic bone disease. 2. UNDER CARBOXYLATED OSTEOCALCIN

Vitamin K is required for the post-translational γ-carboxylation of osteocalcin [65]. Assays have been developed that measure undercarboxylated osteocalcin by either hydroxyapatite precipitation or specific antibodies that recognize undercarboxylated osteocalcin [69, 70]. Undercarboxylated osteocalcin in the serum appears to increase with age, a change that is attributed to an agerelated decrease in vitamin K [71, 72]. Most available assays do not distinguish between the degree of carboxylation of osteocalcin and therefore may overestimate bone formation as undercarboxylated osteocalcin is less likely to be incorporated into the bone matrix. Vergnaud et al. [70] reported an increase in the undercarboxylated osteocalcin to be predictive of hip fracture risk in elderly women. This association was not attributed to increased bone formation, and it was suggested that undercarboxylated osteocalcin was an indicator of poor bone quality. Anticoagulants will potentially affect the proportion of undercarboxylated osteocalcin in the circulation.

B. Specificity and Clearance Osteocalcin has a half-life in the circulation of around 5 minutes and is cleared by the kidney. Proteolytic degradation of the intact osteocalcin molecule occurs both in vivo and in vitro. Therefore, when measuring intact osteocalcin, samples must be maintained at a low temperature to minimize degradation. The N-terminal mid-fragment molecule is more stable [73]. However, it has been suggested that specificity of commercial assays may be poor [74, 75]. The generation of osteocalcin fragment molecules is not fully understood. Osteocalcin fragments could be derived from osteoblastic protein synthesis, catabolism of the intact molecule in bone or proteolysis in blood or other tissues [41]. Generation of fragments may be influenced by metabolic bone disease or medications.

C. Assays

Figure 3

Proposed immunoreactive forms of osteocalcin in normal adult serum. (Adapted from reference [68] with permission.)

Serum osteocalcin has been shown to correlate with bone formation rate and calcium accretion determined by kinetic studies but there was no correlation with bone resorption [76, 77]. Osteocalcin can be measured by both in-house and commercially available methods. Immunoassays use antibodies directed against the C-terminal region of the bovine osteocalcin molecule, or antisera directed against

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human osteocalcin [78–81]. The human-specific assays are reported to be more sensitive than a bovine RIA for the clinical investigation of metabolic bone disease [79]. Automated assays for osteocalcin are now available.

D. Variability 1. CONTROLLABLE

When measuring osteocalcin the assay guidelines on sample collection and handling should be followed as these can vary depending on the type of assay. In general, samples should be kept at low temperature when processing and fresh samples used to avoid freeze–thaw cycles. Short-term storage at −20° is adequate but for long-term storage −70⬚C is advised [71]. Osteocalcin should not be measured in hemolyzed samples as enzymes released from the lyzed red blood cells cause proteolysis of the intact osteocalcin [82]. Lipemic samples should also be avoided as osteocalcin bound to lipid has reduced immunoreactivity. Anticoagulants can affect osteocalcin concentrations in plasma due to the vitamin K dependence [83]. If assays have antisera directed against conformational epitopes chelating agents should not be used for blood collection tubes, as the structural stability of osteocalcin is calcium-dependent. Improved stability has been observed for assays that measure both the intact and the large N-terminal mid-fragment of osteocalcin [84]. Osteocalcin has a circadian rhythm with a nocturnal peak and an afternoon nadir [85]; therefore, samples should be collected at the same time of day if repeat measurements are required. Nielsen et al. [86] proposed that endogenous serum cortisol controls the circadian rhythm of circulating osteocalcin. Osteocalcin levels have been reported to be higher in the winter and spring compared to summer and autumn [15, 51, 87]. There is a significant increase in osteocalcin levels during the luteal phase of the menstrual cycle [48]. 2. UNCONTROLLABLE

In children, osteocalcin levels are higher compared to adults, and correlate with growth velocity rather than age, with maximal values during puberty. Osteocalcin concentrations are higher in men compared to women with an age-related decline but a transient increase after the fifth decade in women [88]. Osteocalcin could increase with age due to kidney impairment. Serum osteocalcin is increased following fracture, remaining elevated for at least 3 months post fracture [89]. A study of Colles’ fracture reported an increase in osteocalcin of 44% (P < 0.001) within 1 week with a return to baseline at 9 months [20]. During pregnancy bone formation markers are elevated during the third trimester,

with the exception of osteocalcin [30]. This could be attributed to the degradation of osteocalcin by the placenta [90].

E. Evaluating Change in Bone Formation Glucocorticoid therapy affects osteocalcin levels in serum. In healthy young men, serum osteocalcin levels fall rapidly with glucocorticoid administration [91]. The effect was dose-dependent and attributed to the reduced biosynthesis of osteocalcin by osteoblasts. A placebocontrolled study of three osteocalcin assays for assessment of prednisolone-induced suppression of bone turnover found that the response of individual subjects to treatment was the same for each assay [92].

F. Evaluating Change in Bone Remodeling 1. ANTIRESORPTIVE THERAPY

Osteocalcin has been shown to decrease in postmenopausal osteoporotic women treated with alendronate therapy [60, 93]. The change in osteocalcin in patients with Paget’s disease of bone treated with bisphosphonates is of a smaller magnitude compared to other bone formation markers. Reid et al. reported a decrease of 33% in osteocalcin compared to an 80% decrease in bone ALP and PINP in patients with Paget’s disease of bone treated with ibandronate for 6 months. In Paget’s disease of bone serum osteocalcin is not increased by the same magnitude as other bone formation markers. There is an increase in osteoclast activity resulting in increased bone turnover and formation of woven bone, which has a disorganized structure. Osteocalcin is produced by mature osteoblasts and is negatively charged and therefore it has an affinity for the positively charged calcium ions in bone mineral. Woven bone has an irregular structure exposing more bone mineral for the osteocalcin to bind to. There is a greater abundance of osteocalcin incorporated into the matrix of woven bone and therefore less in the circulation [94, 95]. 2. ESTROGEN

Bone formation markers are significantly increased in women following surgical menopause, with the magnitude of change in osteocalcin similar to that of bone ALP but not as great as the increase in PINP [29]. 3. RENAL DISEASE

Osteocalcin is rapidly metabolized by the kidney. Therefore osteocalcin levels will be elevated in patients with impaired renal function.

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There is ongoing work to determine the usefulness of biochemical markers of bone formation as a tool for the assessment of bone involvement of cancer patients. Bone formation markers are significantly increased in patients with prostate cancer that have bone metastases compared to those without [34]. However osteocalcin does not appear to be a useful marker in this area of research.

VII. DISCUSSION Biochemical markers of bone turnover can be used to detect both short- and long-term changes in bone turnover. Unlike bone density measurements bone markers can detect changes within weeks. However they are not useful as a diagnostic tool in osteoporosis (Table II). Bone markers may be useful to monitor skeletal response or compliance to treatment of metabolic bone disease [8, 26, 59, 96] and to identify fast bone losers following the menopause [29]. There are several approaches to the identification of nonresponders to treatment. One method is to calculate least significant change, which is based on the within-subject variability. Both the biological and analytical variability are considered [97]. A true responder is identified as having a marker response that is greater than the least significant change. An alternative approach is to define an optimal threshold of change in bone marker using receiver-operating characteristics analysis. Cut-off values are calculated that provide a probability estimate of response to treatment [98]. The choice of analyte is important to provide the optimum information required. In bone disease with defects of collagen metabolism, the procollagen propeptides would be the marker of choice. If there were a dramatic effect on bone turnover, such as in Paget’s disease of bone, then the measurement of total ALP may be sufficient to monitor change in bone turnover [26]. However some report that serum bone ALP and PINP best reflect a significant change in activity of Paget’s disease [99]. It is important to consider confounding factors that may affect the bone formation markers. Sources of variability in individual patients should be identified and minimized (Tables IV and V) [52]. It is not possible to control some factors including age, gender, menopausal status, recent fracture, and renal function. These should be accounted for by using the appropriate reference ranges for the interpretation of results [100]. Some drugs, such as statins [101], affect bone formation and it is important to be aware of any medications that a patient may be taking which may affect bone metabolism. Other sources of variability include diet, circadian rhythm, and exercise, but these can be minimized by collecting fasting samples at a standardized time. Although there is no consensus on the

effect of exercise on bone formation markers it may be useful to be aware of the type and intensity of exercise carried out by the subject. It has been suggested that subjects should refrain from exercise for at least 24 hours before samples are collected [52]. It is useful to establish a sample collection and storage protocol with samples taken within specified time limits. Collecting multiple samples over 3–5 days and averaging the results can make a large improvement in variability [100]. This would be beneficial before the onset of treatment to obtain a true baseline. Osteocalcin and PINP are lower after feeding but the clinical impact of this is thought to be small [102]. In order to reduce variability to a minimum, particularly in a research setting, it would be advisable to collect fasting samples. It is important that appropriate reference data are used for clinical interpretation of results. A reference range from a local population is more desirable than the manufacturers’ reference data. The recent advent of automated assays has provided methods with superior precision and accuracy for bone formation markers [103]. These methods allow many more samples to be processed much faster than the manual assays and availability of bone marker measurements to become more widespread. Automation may, however, enforce some constraints on assays such as limitation of incubation times. It is important that new assay methods are fully validated before data are acceptable. A major concern with the measurement of biochemical markers of bone turnover is interlaboratory variation. There is a large discrepancy between results from different laboratories for the same analyte even when identical methods are used [104]. A standardized evaluation procedure is required to provide accurate results.

References 1. Owen, T. A., Aronow, M., Shalhoub, V., Barone, L. M., Wilming L., Tassinari, M. S., Kennedy, M. B., Pockwinse, S., Lian, J. B., and Stein, G. S. (1990). Progressive development of the rat osteoblast phenotype in vitro: reciprocal relationships in expression of genes associated with osteoblast proliferation and differentiation during formation of the bone extracellular matrix. J. Cell. Physiol. 143, 420–430. 2. Siggelkow, H., Rebenstorff, K., Kurre, W., Niedhart, C., Engel, I., Schulz, H., Atkinson, M. J., and Hufner, M. (1999). Development of the osteoblast phenotype in primary human osteoblasts in culture: comparison with rat calvarial cells in osteoblast differentiation. J. Cell. Biochem. 75, 22–35. 3. Stein, G. S., Lian, J. B., Stein, J. L., Van Wijnen, A. J., and Montecino, M. (1996). Transcriptional control of osteoblast growth and differentiation. Physiol. Rev. 76, 593–629. 4. Owen, T. A., Aronow, M. S., Barone, L. M., Bettencourt, B., Stein, G. S., and Lian, J. B. (1991). Pleiotropic effects of vitamin D on osteoblast gene expression are related to the proliferative and differentiated state of the bone cell phenotype: dependency upon basal levels of gene expression,

538

5.

6.

7.

8.

9.

10.

11.

12.

13.

14.

15.

16.

17.

18.

19.

20.

duration of exposure, and bone matrix competency in normal rat osteoblast cultures. Endocrinology 128, 1496–1504. Eastell, R., Robins, S. P., Colwell, T., Assiri, A. M., Riggs, B. L., and Russell, R. G. (1993). Evaluation of bone turnover in type I osteoporosis using biochemical markers specific for both bone formation and bone resorption. Osteoporos. Int. 3, 255–260. Smedsrod, B., Melkko, J., Risteli, L., and Risteli, J. (1990). Circulating C-terminal propeptide of type I procollagen is cleared mainly via the mannose receptor in liver endothelial cells. Biochem. J. 271, 345–350. Melkko, J., Hellevik, T., Risteli, L., Risteli, J., Smedsrod, B. (1994). Clearance of NH2-terminal propeptides of types I and III procollagen is a physiological function of the scavenger receptor in liver endothelial cells. J. Exp. Med. 179, 405–412. Komi, J., Heikkinen, J., Rutanen, E. M., Halonen, K., Lammintausta, R., and Ylikorkala, O. (2004). Effects of ospemifene, a novel SERM, on biochemical markers of bone turnover in healthy postmenopausal women. Gynecol. Endocrinol. 18, 152–158. Guanabens, N., Pares, A., Alvarez, L., Martinez de Osaba, M. J., Monegal, A., Peris, P., Ballesta, A. M., and Rodes, J. (1998). Collagenrelated markers of bone turnover reflect the severity of liver fibrosis in patients with primary biliary cirrhosis. J. Bone Miner. Res. 13, 731–738. Parfitt, A. M., Simon, L. S., Villanueva, A. R., and Krane, S. M. (1987). Procollagen type I carboxy-terminal extension peptide in serum as a marker of collagen biosynthesis in bone. Correlation with Iliac bone formation rates and comparison with total alkaline phosphatase. J. Bone Miner. Res. 2, 427–436. Dominguez, C. C., Sosa, H. M., Traba, M. L., Alvarez, V. E., and de la Piedra, C. (1998). Biochemical markers of bone formation in the study of postmenopausal osteoporosis. Osteoporos. Int. 8, 147–151. Orum, O., Hansen, M., Jensen, C. H., Sorensen, H. A., Jensen, L. B., Horslev-Petersen, K., and Teisner, B. (1996). Procollagen type I N-terminal propeptide (PINP) as an indicator of type I collagen metabolism: ELISA development, reference interval, and hypovitaminosis D induced hyperparathyroidism. Bone 19, 157–163. Jensen, C. H., Hansen, M., Brandt, J., Rasmussen, H. B., Jensen, P. B., and Teisner, B. (1998). Quantification of the N-terminal propeptide of human procollagen type I (PINP): comparison of ELISA and RIA with respect to different molecular forms. Clinica. Chimica. Acta. 269, 31–41. Brandt, J., Krogh, T. N., Jensen, C. H., Frederiksen, J. K., and Teisner, B. (1998). Thermal instability of the trimeric structure of the N-terminal propeptide of human procollagen type I in relation to assay technology. Clin. Chem. 41, 47–53. Woitge, H. W., Scheidt-Nave, C., Kissling, C., Leidig-Bruckner, G., Meyer, K., Grauer, A., Scharla, S. H., Ziegler, R., and Seibel, M. J. (1998). Seasonal variation of biochemical indexes of bone turnover: results of a population-based study. J. Clin. Endocrinol. Metab. 83, 68–75. Blumsohn, A., Naylor, K. E., Timm, W., Eagleton, A. C., Hannon, R. A., and Eastell, R. (2003). Absence of marked seasonal change in bone turnover: a longitudinal and multicenter cross-sectional study. J. Bone Miner. Res. 18, 1274–1281. Brahm, H., Strom, H., Piehl-Aulin, K., Mallmin, H., and Ljunghall, S. (1997). Bone metabolism in endurance trained athletes: a comparison to population-based controls based on DXA, SXA, quantitative ultrasound, and biochemical markers. Calcif. Tissue. Int. 61, 448–454. Bennell, K. L., Malcolm, S. A., Khan, K. M., Thomas, S. A., Reid, S. J., Brukner, P. D., Ebeling, P. R., and Wark, J. D. (1997). Bone mass and bone turnover in power athletes, endurance athletes, and controls: a 12-month longitudinal study. Bone 20, 477–484. Blumsohn, A., Hannon, R. A., Wrate, R., Barton, J., Al-Dehaimi, A. W., Colwell, A., and Eastell, R. (1994). Biochemical markers of bone turnover in girls during puberty. Clin. Endocrinol 40, 663–670. Joerring, S., Jensen, L. T., Anderson, G. R., and Johansen, J. S. (1992). Types I and III procollagen extension peptides in serum respond to fracture in humans. Arch. Orthop. Trauma Surg. 111, 265–267.

KIM E. NAYLOR AND RICHARD EASTELL 21. Ingle, B. M., Hay, S. M., Bottjer, H. M., and Eastell, R. (1999). Changes in bone mass and bone turnover following ankle fracture. Osteoporos. Int. 10, 408–415. 22. Ingle, B. M., Hay, S. M., Bottjer, H. M., and Eastell, R. (1999). Changes in bone mass and bone turnover following distal forearm fracture. Osteoporos. Int. 10, 399–407. 23. Hernandez, M. V., Guanabens, N., Alvarez, L., Monegal, A., Peris, P., Riba, J., Ercilla, G., Martinez de Osaba, M. J., and Munoz-Gomez, J. (2004). Immunocytochemical evidence on the effects of glucocorticoids on type I collagen synthesis in human osteoblastic cells. Calcif. Tissue. Int. 74, 284–293. 24. Cooper, M. S., Blumsohn, A., Goddard, P. E., Bartlett, W. A., Shackleton, C. H., Eastell, R., Hewison, M., and Stewart, P. M. (2003). 11 beta-hydroxysteroid dehydrogenase type 1 activity predicts the effects of glucocorticoids on bone. J. Clin. Endocrinol. Metab. 88, 3874–3877. 25. Winterbottom, N., Vernon, S., Freeman, K., Daniloff, G., and Seyedin, S. (1993). A serum immunoassay for the C-terminal propeptide of type I collagen. J. Bone Miner. Res. 8, S341. 26. Reid, I. R., Davidson, J. S., Wattie, D., Wu, F., Lucas, J., Gamble, G. D., Rutland, M. D., and Cundy, T. (2004). Comparative responses of bone turnover markers to bisphosphonate therapy in Paget’s disease of bone. Bone 35, 224–230. 27. Alvarez, L., Guanabens, N., Peris, P., Vidal, S., Ros, I., Monegal, A., Bedini, J. L., Deulofeu, R., Pons, F., Munoz-Gomez, J., and Ballesta, A. M. (2001). Usefulness of biochemical markers of bone turnover in assessing response to the treatment of Paget’s disease. Bone 29, 447–452. 28. Suvanto-Luukkonen, E., Risteli, L., Sundstrom, H., Penittinen, J., Kauppila, A., and Risteli, J. (1997). Comparison of three serum assays for bone collagen formation during postmenopausal estrogen-progestin therapy. Clinica. Chimica. Acta. 266, 105–116. 29. Peris, P., Alvarez, L., Monegal, A., Guanabens, N., Duran, M., Pons, F., Martinez de Osaba, M. J., Echevarria, M., Ballesta, A. M., and MunozGomez, J. (1999). Biochemical markers of bone turnover after surgical menopause and hormone replacement therapy. Bone 25, 349–353. 30. Naylor, K. E., Iqbal, P., Fledelius, C., Fraser, R., and Eastell, R. (2000). The effect of pregnancy on bone density and bone turnover. J. Bone Miner. Res. 15, 129–137. 31. Alvarez, L., Torregrosa, J. V., Peris, P., Monegal, A., Bedini, J. L., Martinez de Osaba, M. J., Filella, X., Martin, G., Ricos, C., Oppenheimer, F., and Ballesta, A. M. (2004). Effect of hemodialysis and renal failure on serum biochemical markers of bone turnover. J. Bone Miner. Metab. 22, 254–259. 32. Tsuchida, T., Ishimura, E., Miki, T., Matsumoto, N., Naka, H., Jono, S., Inaba, M., and Nishizawa, Y. (2005). The clinical significance of serum osteocalcin and N-terminal propeptide of type I collagen in predialysis patients with chronic renal failure. Osteoporos. Int. 16, 172–179. 33. de la Piedra, C., Castro-Errecaborde, N. A., Traba, M. L., MendezDavila, C., Garcia-Moreno, C., Rodriguez, D. A., and RodriguezMolina, J. (2003). Bone remodeling markers in the detection of bone metastases in prostate cancer. Clin. Chim. Acta. 331, 45–53. 34. Jung, K., Lein, M., Stephan, C., Von Hosslin, K., Semjonow, A., Sinha, P., Loening, S. A., and Schnorr, D. (2004). Comparison of 10 serum bone turnover markers in prostate carcinoma patients with bone metastatic spread: Diagnostic and prognostic implications. Int. J. Cancer 111, 783–791. 35. Simojoki, M., Santala, M., Risteli, J., and Kauppila, A. (2003). Discrepant expression of carboxy- and aminoterminal propeptides of type I procollagen predicts poor clinical outcome in epithelial ovarian cancer. Gynecol. Oncol. 88, 358–362. 36. Charles, P., Poser, J. W., Mosekilde, L., and Jensen, F. T. (1985). Estimation of bone turnover evaluated by 47Ca-kinetics. Efficiency of serum bone gamma-carboxyglutamic acid-containing protein, serum alkaline phosphatase, and urinary hydroxyproline excretion. J. Clin. Invest. 76, 2254–2258.

Chapter 32 Measurement of Biochemical Markers of Bone Formation 37. Delmas, P. D. (1988). Biochemical markers of bone turnover in osteoporosis. In (B. L., Riggs, and L. J., Melton III, eds). Osteoporosis, 297–316. Raven Press, New York. 38. Fishman, W. H. (1990). Alkaline phosphatase isoenzymes: recent progress. Clin. Biochem. 23, 99–104. 39. Van Hoof, V.O., and De Broe, M. E. (1994). Interpretation and clinical significance of alkaline phosphatase isoenzyme patterns. Crit. Rev. Clin. Lab. Sci. 31, 197–293. 40. Moss, D. W., and Whitby, L. G. (1975). A simplified heat-inactivation method for investigating alkaline phosphatase isoenzyme in serum. Clin. Chim. Acta. 61, 63–71. 41. Calvo, M. S., Eyre, D. R., and Gundberg, C. M. (1996). Molecular basis and clinical application of biological markers of bone turnover. Endocrine Rev. 17, 333–368. 42. Lawson, G. M., Katzman, J. A., Kimlinger, T. K., and O’Brien, J. F. (1985). Isolation and preliminary characterization of a monoclonal antibody that interacts preferentially with the liver isoenzyme of human alkaline phosphatase. Clin. Chem. 31, 381–385. 43. Garnero, P., and Delmas, P. D. (1993). Assessment of the serum levels of bone alkaline phosphatase with a new immunoradiometric assay in patients with metabolic bone disease. J. Clin. Endocrinol. Metab. 77, 1046–1053. 44. Gomez, B. J., Ardakani, S., Ju, J., Jenkins, D., Cerelli, M. J., Daniloff, G. Y., and Kung, V. T. (1995). Monoclonal antibody assay for measuring bone-specific alkaline phosphatase activity in serum. Clin. Chem. 41, 1560–1566. 45. Hata, K., Tokuhiro, H., Nakatsuka, K., Miki, T., Nishizawa, Y., Morii, H., and Miura, M. (1996). Measurement of bone-specific alkaline phosphatase by an immunoselective enzyme assay method. Ann. Clin. Biochem. 33, 127–131. 46. Hill, C. S., and Wolfert, R. L. (1989). The preparation of monoclonal antibodies which react preferentially with human bone alkaline phosphatase and not liver alkaline phosphatase. Clin. Chim. Acta. 186, 315–320. 47. Crofton, P. M. (1982). Biochemistry of the alkaline phosphatase isoenzymes. Crit. Rev. Clin. Lab. Sci. 16, 161–194. 48. Nielsen, H. K., Brixen, K., Bouillon, R., and Mosekilde, L. (1990). Changes in biochemical markers of osteoblastic activity during the menstrual cycle. J. Clin. Endocrinol. Metab. 70, 1431–1437. 49. Greenspan, S. L., Dresner-Pollak, R., Parker, R. A., London, D., and Ferguson, L. (1997). Diurnal variation of bone mineral turnover in elderly men and women. Calcif. Tissue Int. 60, 419–423. 50. Devgun, M. S., Paterson, C. R., and Martin, B. T. (1981). Seasonal changes in the activity of serum alkaline phosphatase. Enzyme 26, 301–305. 51. Douglas, A. S., Miller, M. H., Reid, D. M., Hutchison, J. D., Porter, R. W., and Robins, S. P. (1996). Seasonal differences in biochemical parameters of bone remodelling. J. Clin. Pathol. 49, 284–289. 52. Hannon, R., and Eastell, R. (2000). Preanalytical variability of biochemical markers of bone turnover. Osteoporos. Int. 11 Suppl 6, S30–S44. 53. Crofton, P. M. (1992). Wheat-germ lectin affinity electrophoresis for alkaline phosphatase isoforms in children: age-dependent reference ranges and changes in liver and bone disease. Clin. Chem. 38, 663–670. 54. Cross, N. A., Hillman, L. S., Allen, S. H., Krause, G. F., and Vieira, N. E. (1995). Calcium homeostasis and bone metabolism during pregnancy, lactation and postweaning: a longitudinal study. Am. J. Clin. Nutr. 61, 514–523. 55. Black, A. J., Topping, J., Durham, B., Farquharson, R. G., and Fraser, W. D. (2000). A detailed assessment of alterations in bone turnover, calcium homeostasis, and bone density in normal pregnancy. J. Bone Miner. Res. 15, 557–563. 56. Bowles, S. A., Kurdy, N., Davis, A. M., France, M. W., and Marsh, D. R. (1996). Serum osteocalcin, total and bone specific alkaline phosphatase

57.

58.

59.

60. 61.

62.

63.

64.

65. 66.

67. 68.

69.

70.

71. 72.

73.

74.

75.

76.

539

following isolated tibial shaft fracture. Ann. Clin. Biochem. 33, 196–200. Bowles, S. A., Kurdy, N., Davis, A. M., and France, M. W. (1997). Changes in serum bone-specific alkaline phosphatase following tibial fracture. Ann. Clin. Biochem. 34, 690–691. Woitge, H. W., Seibel, M. J., and Ziegler, R. (1996). Comparison of total and bone specific alkaline phosphatase in patients with nonskeletal disorders or metabolic bone disease. Clin. Chem. 42, 1796–1804. Kress, B. C., Mizrahi, I. A., Armour, K. W., Marcus, R., Emkey, R. D., and Santora, A. C. (1999). Use of bone alkaline phosphatase to monitor alendronate therapy in individual postmenopausal osteoporotic women. Clin. Chem. 45, 1009–1017. Braga de Castro, M. A., Hannon, R., and Eastell, R. (1999). Monitoring alendronate therapy for osteoporosis. J. Bone Miner. Res. 14, 602–608. Rogers, A., Saleh, G., Hannon, R. A., Greenfield, D., and Eastell, R. (2002). Circulating estradiol and osteoprotegerin as determinants of bone turnover and bone density in postmenopausal women. J. Clin. Endocrinol. Metab. 87, 4470–4475. Eastell, R., Hannon, R. A., Cuzick, J., Clack, G., and Adams, J. E. (2002). Effect of anastrozole on bone density and bone turnover: Results of the ‘Arimidex’ (Anastrozole), Tamoxifen, Alone or in Combination (ATAC) Study. J. Bone Miner. Res. 17, S165. Woitge, H.W., Horn, E., Keck, A. V., Auler, B., Seibel, M. J., and Pecherstorfer, M. (2001). Biochemical markers of bone formation in patients with plasma cell dyscrasias and benign osteoporosis. Clin. Chem. 47, 686–693. Abildgaard, N., Brixen, K., Eriksen, E. F., Kristensen, J. E., Nielsen, J. L., and Heickendorff, L. (2004). Sequential analysis of biochemical markers of bone resorption and bone densitometry in multiple myeloma. Haematologica 89, 567–577. Lee, A. J., Hodges, S., and Eastell, R. (2000). Measurement of osteocalcin. Ann. Clin. Biochem. 37 ( Pt 4), 432–446. Taylor, A. K., Linkhart, S., Mohan, S., Chrinstenson, R. A., Singer, F. R., and Baylink, D. (1990). Multiple osteocalcin fragments in human urine and serum as detected by a midmolecule osteocalcin radioimmunoassay. J. Clin. Endocrinol. Metab. 70, 467–472. Gundberg, C., and Weinstein, R. S. (1986). Multiple immunoreactive forms in uremic serum. J. Clin. Invest. 77, 1762–1767. Garnero, P., Grimaux, M., Seguin, P., and Delmas, P. D. (1994). Characterization of immunoreactive forms of human osteocalcin generated in vivo and in vitro. J. Bone Miner. Res. 9, 255–264. Merle, B., and Delmas, P. D. (1990). Normal carboxylation of circulating osteocalcin (bone Gla protein) in Paget’s disease of bone. Bone Miner. 11, 237–245. Vergnaud, P., Garnero, P., Meunier, P. J., Breart, G., Kamihagi, K., and Delmas, P. D. (1997). Undercarboxylated osteocalcin measured with a specific immunoassay predicts hip fracture in elderly women: the EPIDOS study. J. Clin. Endocrinol. Metab. 82, 719–724. Power, M. J., and Fottrell, P. F. (1991). Osteocalcin: diagnostic methods and clinical applications. Crit. Rev. Clin. Lab. Sci. 28, 287–335. Szulc, P., Arlot, M., Chapuy, M.-C., Duboeuf, F., Meunier, P. J., and Delmas, P. D. (1994). Serum undercarboxylated osteocalcin correlates with hip bone mineral density in elderly women. J. Bone Miner. Res. 9, 1591–1595. Rosenquist, C., Qvist, P., Bjarnason, N., and Christiansen, C. (1995). Measurement of a more stable region of osteocalcin in serum by ELISA with two monoclonal antibodies. Clin. Chem. 41, 1439–1445. Colford, J., Sailer, D., and Langman, C. (1997). Five osteocalcin assays compared: tracer specificity, fragment interference, and calibration. Clin. Chem. 43, 1240–1241. Souberbielle, J. C., Marque, D., Bonnet, P., Herviaux, P., and Sachs, C. (1997). Simple method to evaluate specificity of osteocalcin immunoassays. Clin. Chem. 43, 1663–1664. Eastell, R., Delmas, P. D., Hodgson, S. F., Eriksen, E. P., and Mann, K. G. (1988). Bone formation rate in older normal women: concurrent

540

77.

78. 79.

80.

81.

82.

83.

84.

85.

86.

87.

88.

89. 90.

assessment with bone histomorphometry, calcium kinetics, and biochemical markers. J. Clin. Endocrinol. Metab. 67, 741–748. Brown, J. P., Delmas, P. D., Malaval, L., Edouard, C., Chapuy, M. C., and Meunier, P. J. (1984). Serum bone Gla-protein: a specific marker for bone formation in postmenopausal osteoporosis. Lancet 1, 1091–1093. Diego, E. M. D., Guerrero, R., and de la Piedra, C. (1994). Six osteocalcin assays compared. Clin. Chem. 40, 2071–2077. Garnero, P., Grimaux, M., Demiaux, B., Preaudat, C., Seguin, P., and Delmas, P. D. (1992). Measurement of serum osteocalcin with a human-specific two-site immunoradiometric assay. J. Bone Miner. Res. 7, 1389–1398. Taylor, A. K., Linkhart, S. G., Mohan, S., and Baylink, D. J. (1988). Development of a new radioimmunoassay for human osteocalcin: evidence for a mid molecule epitope. Metabolism 37, 872–877. Masters, P. W., Jones, R. G., Purves, D. A., Cooper, E. H., and Cooney, J. M. (1994). Commercial assays for serum osteocalcin give clinically discordant results. Clin. Chem. 40, 358–363. Tracy, R. P., Andrianorivo, A., Riggs, B. L., and Mann, K. G. (1990). Comparison of monoclonal and polyclonal antibody-based immunoassays for osteocalcin: a study of sources of variation in assay results. J. Bone Miner. Res. 5, 451–461. Power, M. J., O’Dwyer, B., Breen, E., and Fottrell, P. F. (1991). Osteocalcin concentrations in plasma prepared with different anticoagulants. Clin. Chem. 37, 281–284. Blumsohn, A., Hannon, R. A., and Eastell, R. (1995). Apparent instability of osteocalcin in serum as measured with different commercially available immunoassays. Clin. Chem. 41, 318–319. Eastell, R., Calvo, M. S., Burritt, M. F., Offord, K. P., Russell, R. G. G., and Riggs, B. L. (1992). Abnormalities in circadian patterns of bone resorption and renal calcium conservation in type I osteoporosis. J. Clin. Endocrinol. Metab. 74, 487–494. Nielsen, H. K., Brixen, K., Kassem, M., Charles, P., and Mosekilde, L. (1992). Inhibition of the morning cortisol peak abolishes the expected morning decrease in serum osteocalcin in normal males: evidence of a controlling effect of serum cortisol on the circadian rhythm in serum osteocalcin. J. Clin. Endocrinol. Metab. 74, 1410–1414. Thomsen, K., Eriksen, E. F., Jorgensen, J. C. R., Charles, P., and Mosekilde, L. (1989). Seasonal variation of bone-Gla protein. Scand. J. Clin. Lab. Invest. 49, 605–611. Vanderschueren, D., Gevers, G., Raymaekers, G., Devos, P., and Dequeker, J. (1990). Sex and age-related changes in bone and serum osteocalcin. Calcif. Tissue. Int. 46, 179–182. Obrant, K. J., Merle, B., Bejui, J., and Delmas, P. D. (1990). Serum bone-gla protein after fracture. Clin. Orthop. 258, 300–303. Rodin, A., Duncan, A., Quartero, H. W. P., Pistofidis, G., Mashiter, G., Whitaker, K., Crook, D., Stevenson, J. C., Chapman, M. G., and Fogelman, I. (1989). Serum concentrations of alkaline phosphatase isoenzymes and osteocalcin in normal pregnancy. J. Clin. Endocrinol. Metab. 68, 1123–1127.

KIM E. NAYLOR AND RICHARD EASTELL 91. Godschalk, M. F., and Downs, R. W. (1988). Effect of short-term glucocorticoids on serum osteocalcin in healthy young men. J. Bone Miner. Res. 3, 113–115. 92. Heuck, C., and Wolthers, O. D. (1998). A placebo-controlled study of three osteocalcin assays for assessment of prednisolone-induced suppression of bone turnover. J. Endocrinol. 159, 127–131. 93. Garnero, P., Shih, W. J., Gineyts, E., Karpf, D. B., and Delmas, P. D. (1994). Comparison of new biochemical markers of bone turnover in late postmenopausal osteoporotic women in response to alendronate treatment. J. Clin. Endocrinol. Metab. 79, 1693–1700. 94. Delmas, P. D., Demiaux, B., Malaval, L., Chapuy, M.-C., and Meunier, P. J. (1986). Serum bone GLA-protein is not a sensitive marker of bone turnover in Paget’s disease of bone. Calcif. Tiss. Int. 38, 60–61. 95. Ingram, R. T., Park, Y. K., Clarke, B. L., and Fitzpatrick, L. A. (1994). Age- and gender-related changes in the distribution of osteocalcin in the extracellular matrix of normal male and female bone. Possible involvement of osteocalcin in bone remodeling. J. Clin. Invest. 93, 989–997. 96. Alvarez, L., Peris, P., Guanabens, N., Vidal, S., Quinto, L., Monegal, A., Pons, F., Ballesta, A. M., and Munoz-Gomez, J. (2004). Long-term biochemical response after bisphosphonate therapy in Paget’s disease of bone. Proposed intervals for monitoring treatment. Rheumatology (Oxford) 43, 869–874. 97. Hannon, R. A., Blumsohn, A., Naylor, K. E., and Eastell, R. (1998). Response of biochemical markers of bone turnover to hormone replacement therpay: impact of biological variability. J. Bone Miner. Res. 13, 1124–1133. 98. Riggs, B.L. (2000). Are biochemical markers for bone turnover clinically useful for monitoring therapy in individual osteoporotic patients? Bone 26, 551–552. 99. Alvarez, L., Ricos, C., Peris, P., Guanabens, N., Monegal, A., Pons, F., and Ballesta, A. M. (2000). Components of biological variation of biochemical markers of bone turnover in Paget’s bone disease. Bone 26, 571–576. 100. Bernardi, D., Zaninotto, M., and Plebani, M. (2004). Requirements for improving quality in the measurement of bone markers. Clin. Chim. Acta. 346, 79–86. 101. Mundy, G., Garrett, R., Harris, S., Chan, J., Chen, D., Rossini, G., Boyce, B., Zhao, M., and Gutierrez, G. (1999). Stimulation of bone formation in vitro and in rodents by statins. Science 286, 1946–1949. 102. Clowes, J. A., Hannon, R. A., Yap, T. S., Hoyle, N. R., Blumsohn, A., and Eastell, R. (2002). Effect of feeding on bone turnover markers and its impact on biological variability of measurements. Bone 30, 886–890. 103. Schmidt-Gayk, H., Spanuth, E., Kotting, J., Bartl, R., Felsenberg, D., Pfeilschifter, J., Raue, F., and Roth, H. J. (2004). Performance evaluation of automated assays for beta-CrossLaps, N-MID-Osteocalcin and intact parathyroid hormone (BIOROSE Multicenter Study). Clin. Chem. Lab. Med. 42, 90–95. 104. Seibel, M. J., Lang, M., and Geilenkeuser, W. J. (2001). Interlaboratory variation of biochemical markers of bone turnover. Clin. Chem. 47, 1443–1450.

Chapter 33

Measurement of Biochemical Markers of Bone Resorption Marius E. Kraenzlin Markus J. Seibel

Division of Endocrinology, Diabetology and Clinical Nutrition, University Hospital Basel, Petersgraben 4, CH-4031 Basel, Switzerland Bone Research Program, ANZAC Research Institute, University of Sydney and Concord Hospital, Sydney, Australia

I. Introduction II. Collagen Related Markers III. Noncollagenous Protein of the Bone Matrix

IV. Osteclast Enzymes References

I. INTRODUCTION

The bone matrix is composed of approximately 70% mineral (mainly hydroxylapatite crystals) and 30% organic matter. The latter consists of collagen, noncollagenous proteins, and proteoglycans. The main structural protein of bone is type I collagen, which constitutes approximately 90% of the organic matrix (see Chapters 1 and 12). During the process of bone resorption, collagen and the other constituents of the organic matrix are degraded by the action of various proteases. The resulting breakdown products, as well as the enzymes produced by active osteoclasts and the other components involved in osteoclast regulation and function, are consecutively released into the circulation and partly eliminated by the kidney. In recent years, considerable progress has been made in the isolation and characterization of cellular and extracellular components of the skeletal matrix, which in turn have facilitated the development of biochemical markers that specifically reflect either bone formation or bone resorption. These biochemical indices have greatly enriched the spectrum of analytes used in the assessment

Bone is a metabolically active tissue and undergoes continuous remodeling, a process that largely relies on the activity of osteoblasts (bone formation) and osteoclasts (bone resorption). Under normal conditions and through the action of systemic hormones and local mediators, bone formation and bone resorption are coupled to each other, maintaining a finely tuned balance in skeletal turnover and metabolism (see Chapters 15–20). In contrast, metabolic bone diseases are characterized by more or less pronounced imbalances in, and uncoupling of, bone turnover. For several reasons, disorders of bone and mineral metabolism have become increasingly relevant to both basic research and clinical practice. Consequently, the interest in, and the need for effective measures to be used in experimental research, and in the screening, diagnosis, and follow-up of patients suffering form bone disorders has markedly grown. Dynamics of Bone and Cartilage Metabolism

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Copyright © 2006 by Academic Press. All rights of reproduction in any form reserved.

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of skeletal pathologies. They are noninvasive, comparatively inexpensive and, when applied and interpreted correctly, helpful tools in the diagnostic and therapeutic assessment of metabolic bone disease. The bone resorption markers currently available or under investigation, and some of their characteristics, are summarized in Table I. Bone resorption may be assessed by measurement of any of the following components in urine and/or serum. 1. Collagen degradation products: hydroxyproline (OHPr), hydroxlysine glycosides, the 3-hydroxy-pyridinium crosslinks pyridinoline (PYD), and deoxypyridinoline (DPD) or their higher-molecular-weight derivates originating from the nonhelical (i.e. amino-terminal Table I. Marker (abbreviation)

cross-linked telopeptide (NTX-I), carboxy-terminal cross-linked telopeptides (CTX-I) and helical (helical peptide; HELP)) region of the collagen type I molecule. 2. Noncollagenous proteins of the bone matrix: bone sialoprotein, osteocalcin. 3. Enzymes expressed and secreted by osteoclasts during active bone resorption: tartrate resistant acid phosphatase 5b, some cathepsins. Efforts have been made to demonstrate the clinical utility of biochemical markers of bone turnover in the identification and definition of osteoporotic fracture risk (see Chapter 37), early diagnosis of metabolic bone disease (see Chapters 41–52), determination of effective therapy

Biochemical Markers of Bone Resorption

Tissue of origin

Analytical specimen

Analytical method

Remarks/specificity

Collagen related markers Hydroxyproline, total and dialyzable (Hyp)

Bone, cartilage, soft tissue, skin

Urine

Colorimetry HPLC

Hydroxylysine-glycosides

Bone, soft tissue, skin, serum complement

Urine (serum)

HPLC ELISA

Pyridinoline (PYD)

Bone, cartilage, tendon, blood vessels

Urine Serum

HPLC ELISA

Deoxypyridinoline (DPD)

Bone, dentin

Urine Serum

HPLC ELISA

Carboxy-terminal crosslinked telopeptide of type I collagen (ICTP, CTX-MMP) Carboxy-terminal crosslinked telopeptide of type I collagen (CTX-I) Amino-terminal crosslinked telopeptide of type I collagen (NTX-I) Collagen I alpha 1 helicoidal peptide (HELP)

Bone, skin

Serum

RIA

All tissues containing type I collagen

Urine (a-/β) Serum (β only) Urine Serum

ELISA RIA

All tissues containing type I collagen All tissues containing type I collagen

Urine

ELISA CLIA RIA ELISA

Present in all fibrillar collagens and partly collagenous proteins, including C1q and elastin. Present in newly synthesized and mature collagen, i.e. both collagen synthesis and tissue breakdown contribute to urinary hydroxyproline Hydroxylysine in collagen is glycosylated to varying degrees, depending on tissue type. Glycosylgalactosyl-OHLys in high proportion in collagens of soft tissues, and C1q; Galyctosyl-OHLys in high proportion in skeletal collagens Collagens, with highest concentrations in cartilage and bone; absent from skin; present in mature collagen only Collagens, with highest concentration in bone; absent from cartilage or skin; present in mature collagen only Collagen type I, with highest contribution probably from bone; may be derived from newly synthesized collagen Collagen type I, with highest contribution probably from bone. Isomerization of aspartyl to β-aspartyl occurs with aging of collagen molecule. Collagen type I, with highest contribution from bone

Degradation fragment derived from the helical part of type I collagen (α1 chain, AA 620–633). Correlates highly with other markers of collagen degradation, no specific advantage or difference in regards to clinical outcomes

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Chapter 33 Measurement of Biochemical Markers of Bone Resorption Table I. Marker (abbreviation) Noncollagenous proteins Bone sialoprotein (BSP)

Osteocalcin fragments (ufOC, U-Mid-OC, U-LongOC)

Biochemical Markers of Bone Resorption—cont’d

Tissue of origin Bone, dentin, hypertrophic cartilage

Analytical specimen Serum

Analytical method RIA ELISA

Bone

Urine

ELISA

Tartrate-resistant acid phosphatase (TRACP)

Bone Blood

Plasma Serum

Colorimetry RIA ELISA

Cathepsins (e.g. K, L)

K: Primarily in osteoclasts L: Macrophages, Osteocalsts

Plasma, serum

ELISA

Remarks/specificity Acidic, phosphorylated glycoprotein, synthesized by osteoblasts and osteoclastic-like cells, laid down in bone extracellular matrix. Appears to be associated with osteoclast function Certain age-modified OC fragments are released during osteoclastic bone resorption and may be considered an index of bone resorption

Osteoclast enzymes

for those with established disease, and monitoring the effects of these therapies (see Chapters 38 and 39). The value of a biochemical marker of bone turnover depends on several criteria. Firstly, the ideal marker should be tissue specific. In the case of bone, this applies to osteocalcin, but not, for example, to hydroxyproline and many other “bone markers”. Consequently, the serum or urine concentrations of these markers are also affected by the turnover of nonskeletal tissues. Secondly, a marker should be specific for either bone formation or bone resorption. However, some of the components used as bone turnover markers reflect, at least in part, both bone formation and resorption (e.g., urinary hydroxyproline, certain osteocalcin fragments). Thirdly, changes in bone markers are usually not disease specific, but reflect alterations in skeletal metabolism independent of the underlying cause. Finally, a marker should be easily measurable in serum or urine by specific and sensitive techniques, and the factors that control its synthesis, metabolism, elimination, and variation should be well known. With most markers, however, there is still only limited information available about their metabolism and the factors that influence their production and degradation.

Six isoenzymes found in human tissues (osteoclasts, platelets, erythrocytes) Band 5b predominant in bone (osteoclasts). Enzyme identified in both the ruffled border of the osteoclast membrane and the secretions in the resorptive space Cathepsin K, cysteine protease, plays an essential role in osteoclast-mediated bone matrix degradation by cleaving helical and telopeptide regions of collagen type I Cathepsin K and L cleave the loop domain of TRACP and activate the latent enzyme. Cathepsin L has a similar function in macrophages.Tests for measurement of cathepsins in blood are presently under evaluation

This chapter will review both the basics of, and some recent progress made in the development of markers of bone resorption. The clinical application of the bone resorption assays will be reviewed in other chapters.

II. COLLAGEN RELATED MARKERS A. Hydroxyproline Until a few years ago, the best-established and most widely used marker of bone resorption was urinary hydroxyproline (OHPr). Hydroxyproline occurs mainly in fibrillar collagens, where it accounts for 13–14% of the total amino acid content. Hydroxyproline is produced as a result of the post-translational modifications of proline by prolyl hydroxylase during collagen synthesis. The amino acid occurs almost exclusively in collagen and therefore its urinary levels are considered to reflect collagen turnover in bone and other tissues [1]. There are, however, several problems with using hydroxyproline as a marker for bone resorption. First, hydroxyproline is not specific for bone. It is found in all

544 collagens, not just type I [1]. Second, the use of hydroxyproline as a quantitative measure of collagen breakdown is hampered by the fact that the free amino acid is reabsorbed by the kidneys and eventually degraded in the liver into products that can not be detected by the hydroxyproline assay. Furthermore, it has been suggested that about 10% of the total urinary hydroxyproline pool is released during the proteolytic processing of newly synthesized procollagen and therefore indicative of collagen formation [2]. Thus, urinary hydroxyproline is derived from breakdown of all types of collagen in the body, and at least in part also from collagen-containing foods in the diet. Measurement of urinary hydroxyproline involves acid hydrolysis of the OHPr-containing peptides by adding an equal volume of 12 N HCl to the urine and heating at 100–105°C for 10–18 hours. This hydrolysis step is required to measure total amounts of OHPr, as more than 90% of urinary OHPr is peptide-bound. The acid hydrolysis is followed by oxidation of free hydroxyproline to pyrrole with chloramine T, and the colorimetric reaction of the pyrrole with p-dimethyl-aminobenzaldehyde (Ehrlich reagent) is measured. The urinary OHPr concentrations are read from a standard curve based on aqueous OHPr solutions which are run simultaneously with the samples [3–7]. The intra- and interassay variation reported for the colorimetric assay ranges from 2–10% and 8–15%, respectively. The recovery ranges from 70–90% and the sensitivity is in the order of 5 umol/L [4, 5, 8]. Injection of iodinated contrast medium is known to interfere with hydroxyproline determination [9]. The iothalamic acid contained in contrast medium reduces the optical extinction by 12–84%, depending on the amount of iodinated contrast medium. The iodide ions are probably released during thermal hydrolysis and reduced to iodine by chloramine T, at the same time the chloramine T is inactivated, and thus not available to oxidize OHPr [9]. Further to the colorimetric assay, several HPLC methods for urinary hydroxyproline have been published. Hydroxyproline, like most imino and amino acids, absorbs only weakly in the ultraviolet-visible region and possesses no native fluorescence. Therefore, a hydroxyproline derivative is needed to allow photometric or fluorimetric detection. Dimethylaminoazobenzene-4-sulfonyl chloride, phenylisothiocyanate, o-phthaladehyde and 4-chlor-7nitrobenzo-2-oxa-1,3-diazole have been used as derivatization agents [10–12]. The separation is then performed on a reverse-phase column followed by UV-detection. Quantitation of OHPr is done by using an internal or external standard (Fig. 1). The intra- and interassay variation, as well as recovery and sensitivity described for the HPLC, are considerably better than for the

Marius E. Kraenzlin and Markus J. Seibel

Figure 1 Separation of urinary hydroxyproline (OHPr) by highperformance liquid chromatography (IS: internal standard; FMOC: 9-fluorenylmethyl-chlorformate).

colorimetric method; the interassay variation being 4–5% and analytical recovery ranging from 95–110%. OHPr measurements are usually performed in a fasting urine sample collected over 2 hours in the morning and expressed as OHPr/creatinine ratio. Since OHPr is absorbed through the GI tract, the intake of collagen-rich foods needs to be restricted for at least 12 hours before sample collection. In adults, a higher 24-hour OHPr excretion has been reported for men than for women, reflecting the difference in total body collagen mass. This difference disappears when corrections are made for creatinine clearance [7]. Urinary OHPr levels follow a circadian rhythm, which appears to be fairly constant from day to day [8].

B. Glycosylated Hydroxylysine Hydroxylysine is another amino acid unique to collagen and collagen-like peptides. Similar to OHPr, hydroxylysine is produced by a post-translational modification of lysine during collagen synthesis. Unlike OHPr, these amino acids can then undergo further modification by glycosylation, giving rise to galactosyl hydroxylysine (GH) and glucosylgalactosyl hydroxylysine (GGH) [13, 14]. This subsequent glycosylation of hydroxylysine differs in collagens from different tissues. The monoglycosylated, galactosyl-hydroxylysine is enriched in bone compared with the disaccharide form, glucosyl-galactosylhydroxylysine, which is the major form in skin. Although hydroxylysine is found in all collagens, GH is fiveto sevenfold more concentrated in type I collagen

Chapter 33 Measurement of Biochemical Markers of Bone Resorption

of bone than in type I collagen of skin. Measurements of GH are therefore thought to reflect mainly resorption of bone collagen [13, 14]. In addition, hydroxylysine glycosides, unlike hydroxyproline, are apparently not significantly metabolized prior to urinary excretion and are not significantly affected by diet. Because of this, it has been estimated that urinary hydroxylysine glycosides can account for 50–100% of collagen breakdown and not the 10–25% estimated for OHPr [13]. Under normal conditions hydroxylysine glycosides represent 80% of the total urinary hydroxylysine. Ten percent of the hydroxylysine is present in the free form and the remainder is peptide-bound, suggesting that free hydroxylysine is largely metabolized and not excreted [15]. This ratio of free to peptide-bound forms varies with age, the free and peptide-bound hydroxylysine being most prominent in children [15]. The major difficulty with measuring urinary hydroxylysine glycosides has been the methods involved in their determination. The procedure generally consists of separation by ion-exchange chromatography followed by amino acid analysis [16, 17]. Another method for measuring these glycosides has been introduced, comprising the conversion of the amino groups into fluorescent derivatives prior to their separation by reversed phase high-performance liquid chromatography (HPLC) [18]. GH is derivatized by adding dansyl chloride to the urine samples. A hydrolytic pretreatment is not required. The separation is carried out by reversed-phase HPLC involving a gradient elution system. Quantification of the measured analyte is performed using an external standard extracted from urine or with commercially available, purified lysine [19]. Intra- and interassay variations are in the order of 5–8% and 6–7%, respectively [20, 21]. The sensitivity appears to be in the order of picomoles [18]. Although the introduction of the HPLC method for measurement of the glycosylated hydroxylysine is an improvement, the method still remains technically demanding and slow.

Figure 2

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An immunoassay for urinary galactosyl-hydroxylysine has been developed based on polyclonal antibodies using GH purified from sea sponges as antigen [22]. The assay showed no significant cross-reactivity to GGH, hydroxylysine, galactose, or related sugars. The detection limit of the assay is 1.25 µM. The intra- and interassay variations were less than 10% and 14%, respectively [22]. In contrast to the HPLC method, which is specific for the free forms of GH, the immunoassay is believed to measure both the free and peptide forms of GH. In general, GH is measured in urine after an overnight fast. The stability properties of glycosylated hydroxylysine have not been fully characterized, but it may be necessary to add boric acid to preserve the urine samples [23]. There has been one report measuring GH in serum [24]. To date, there have been some reports on the value of GH in the evaluation of metabolic and metastatic bone disease. However, more research is required to determine the usefulness of this marker in clinical practice [18, 20, 22, 25, 26].

C. The 3-Hydroxy-Pyridinium Crosslinks of Collagen The pyridinium crosslinks, hydroxylysyl-pyridinoline or pyridinoline (PYD) and lysyl-pyridinoline or deoxypyridinoline (DPD) have received considerable attention over the past decade as specific and highly sensitive markers of bone resorption. PYD and DPD are derived by enzyme-mediated oxidation at specific hydroxylysine and lysine residues within the collagen molecule (Fig. 2) and found in mature, but not immature or newly synthesized type I, II, and III collagens [27, 28]. The crosslinks covalently link collagen molecules between two telopeptides and a triple-helical sequence at two intermolecular

Molecular structure of pyridinoline crosslinks. R1 and R2 are telopeptide sequences and R3 represents a helical sequence; for the free crosslinks, R1, R2, and R3 = -CH(COOH)NH2.

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Figure 3 Schematic representation of the intermolecular crosslinking in type I collagen and the locations of the peptide sequence selected for the N-terminal (NTX) and C-terminal (CTX) immunoassays (modified and adapted from references [28, 80, 82]).

sites and therefore stabilize collagen fibrils (Fig. 3). Of the two forms, PYD and DPD, PYD is the major crosslink in all connective tissues, whereas DPD is found in high amounts in mineralized tissue. In human bone, for example, the DPD:PYD ratio is 1:3.5, as compared to a ratio of 1:10 in human cartilage. Significant amounts of DPD are found in bone and dentine only, with very small amounts in the aorta and ligaments. DPD is therefore considered to be specific for bone collagen degradation. PYD and DPD, in contrast to OHPr, are completely absent from the collagen of normal skin (see also Chapter 2). During collagen breakdown, the pyridinoline crosslinks are released and cleared by the kidneys. In urine, PYD and DPD are present as both free amino acid derivatives (about 40%) and as oligopeptide-bound fractions (about 60%) [29]. The free form can be measured directly, whereas the conjugated form has to be hydrolyzed before assay, in which case the total amount of PYD and DPD is determined [30]. The relatively normal values for DPD excretion in patients with renal dysfunction support the notion that crosslink excretion is generally unaffected by changes in renal function [31]. They are apparently not metabolized within the body and their levels are, in contrast to OHPr, not influenced by diet. However, there is some evidence that some free PYD and DPD excreted into the urine is produced by the kidney [32].

There are two principal methods for the measurement of urinary pyridinoline crosslink concentrations: HPLC analyses with or without hydrolysis of the urine, and immunoassay detection. Although the HPLC assays are considered the reference methods, they are cumbersome and labor-intensive. At present, most laboratories use a number of different immunoassays either urinary-free pyridinolines, or the “crosslinked” N- and C-terminal telopeptides in urine or serum [33]. 1. Measurement of 3-Hydroxy-pyridinium Crosslinks of collagen by HPLC

Urine samples are usually hydrolyzed with 6N HCl at 107°C for 16 hours in order to convert all crosslink components into the peptide-free form. By omitting the hydrolysis step, only the peptide-free fraction of PYD and DPD is measured. Following a partition chromatography step (CF1 cellulose), the pyridinoline crosslink components are separated by reversed-phase ion-paired HPLC. Eluting fluorescence peaks are quantified using an external PYD and/or DPD standard extracted from bone or urine (Fig. 4) [34, 35]. A problem in the interpretation of results obtained by HPLC was the lack of a functional internal standard. Although recovery for PYD and DPD from hydrolysis to identification is usually between 80–90% or higher, the addition of an internal standard is required in order to improve the precision of the overall method. This internal

Chapter 33 Measurement of Biochemical Markers of Bone Resorption

Figure 4

Separation of urinary pyridinoline crosslinks, pyridinoline (PYD) and deoxypypridinoline (DPD) by reverse-phase high-performance liquid chromatography.

standard is particularly necessary to correct for betweensample variation in recovery [30]. During recent years, a number of reports have described the successful use of either pyridoxine, isodesmosine, derived from elastin, or acetyl-pyrididinoline, a semisynthetic derivative, as internal standards (Fig. 5) [30, 36–38]. Pyridoxine shows a chromatographic behavior that is different from that of pyridinium crosslinks. While isodesmosine may occur endogenously in urine, acetylated PYD does not have these limitations [38, 39]. However, acetylated PYD

547

hydrolyzes quickly under acidic conditions and can therefore only be added to the samples after hydrolysis [39]. Mean recoveries of PYD and DPD, monitored using the internal standard isodesmosine, were 97% and 93% respectively, and interassay variation was lowered from 15.1% to 5.3% for PYD and from 20.8% to 4.6% for DPD [30, 36]. Although the use of an internal standard has considerably improved analytical precision, the HPLC procedure is not yet standardized, and results from different laboratories are not necessarily comparable [39–42]. Laboratories using the same standard usually provide values with a correlation coefficient of r > 0.95, but values may differ in absolute terms. Therefore, an accepted international reference material is needed to enable comparison between different laboratories and centers [39, 40, 42, 43]. Methods to chemically synthesize PYD and DPD have been described and synthetic PYD and DPD have been used recently to characterize and validate an HPLC assay for total and free PYD and DPD [39, 44, 45]. The coefficient of variation for the determination of PYD varies between 4–8%, and for DPD between 6–10%, with a detection limit of 1 pmol. Calculated as a coefficient of variation, the overall reproducibility of the complete analysis, including CF1 column fractionation, was found to be 12–16% [34, 46, 47]. The pyridinoline measurements are usually performed on a fasting urine sample collected over 2 hours in the morning and are expressed as a crosslink/creatinine ratio. The values in fasting morning urine samples correlate well with the 24-hour output of PYD and DPD [30, 31]. Sulfasalazine (salicyloazosulfapyridine), a compound used in rheumatoid arthritis or chronic inflammatory bowel disease, has been reported to interfere with the determination of pyridinoline in treated individuals [48]. Sulfasalazine is metabolized into sulfapyridine and 5-aminosalicylic acid by intestinal bacteria. These metabolites are absorbed and then excreted in the urine. Sulfapyridine and 5-aminosalicylic acid peaks elute at a retention time similar to that of the peaks for PYD and DPD (Fig. 6). The retention time of sulfapyridine or its derivative can be delayed, and thereby separated from the pyridinoline peaks, by increasing the solvent heptabutyric acid (HFBA) [48]. 2. Measurement of Pyridinium Crosslinks by Immunoassay

Figure 5

Separation of urinary pyridinoline crosslinks, pyridinoline (PYD) and deoxypypridinoline (DPD) by reverse phase high performance liquid chromatography, with the addition of an internal standard (IS), acetyl-pyridinoline (reproduced from reference [214], with permission).

Immunoassays for pyridinium crosslinks utilize antibodies directed against either the free amino acids or the small peptides containing the crosslink. All of these assays offer considerable advantages in the ease and speed of analysis, but usually at the expense of precision and reproducibility.

548

Figure 6

High-performance liquid chromatogram of pyridinoline crosslinks illustrating the interference by sulfapyridine.

An ELISA for the direct measurements of free crosslink components in urine was developed initially using polyclonal, and later monoclonal, antibodies [49, 50]. This assay did not, however, distinguish between PYD and DPD, as both crosslinks showed similar reactivity with the antibodies. An intraassay variation of approximately 7% and interassay variation of 11%, as well as a sensitivity of 25 nmol/L have been reported [49]. An ELISA based on a monoclonal antibody has been described which shows 100% reactivity with free DPD and exhibits less than 2% cross-reactivity with PYD [51]. The intra- and interassay variations were 4–8% and 4–13.7%, respectively, with a sensitivity of 2 nM DPD. Another immunoassay developed by the same group measures both free PYD and free DPD in urine, with the monoclonal antibody employed showing similar affinity for each crosslink [50]. Peptides larger than approximately 1000 Da exhibited minimal cross-reactivity in this assay. In early 1998, the immunoassay for free DPD in urine was for the first time established on an automated system, making this assay now also feasible for large-scale routine analyses [52, 53]. The results obtained with the two immunoassays for free crosslinks, including the automated version, correlated well with values for total DPD measured by HPLC. The total assay precision for the automated chemiluminescence immunoassay (CLIA) was <10% at DPD concentrations as low as 30 nmol/L [52, 53]. PYD and DPD are generally analyzed in a fasting urine sample and corrected for creatinine (see below). As an

Marius E. Kraenzlin and Markus J. Seibel

alternative option, measurement of PYD in sweat has been described [54]. Sweat is collected with a special patch, applied to the abdomen or lower back, over 5 days. Free PYD is measured in extracted analyte and corrected for potassium, a marker for sweat output. The total (analytical and biological) variabilities for PYD/K were 16.2%, 6.8%, and 14.7%, respectively [54]. However, clinical data on these measurements are sparse [55]. An important consideration when using the immunoassays for pyridinium crosslinks in urine is the knowledge of the normal physiological variations that occur in the excretion of the free crosslink fraction. It has been shown that the proportion of free pyridinolines is smaller in elderly subjects than in adolescents and in healthy adults. In particular, elderly subjects with vitamin D deficiency seem to excrete a lesser amount of free crosslink components than healthy subjects. In the elderly, a change in the molecular distribution of pyridinoline crosslinks is observed, that is an increased proportion of the largest peptide bound forms (particularly >10000 Da), and a decreased proportion of free pyridinolines and smallest peptide bound forms [56]. Consequently, the increase in urinary crosslinks often seen in subjects beyond the age of 70 years is mainly due to a increase in the peptidebound (i.e. high-molecular-weight) fraction [57]. This seems to indicate that changes in crosslink metabolism are at least partly responsible for the age-related increase in urinary crosslink concentration. Furthermore, in patients treated with bisphosphonates or hormone replacement therapy, it has been suggested that changes in the free fraction of immunoreactive crosslinks do not reflect the changes in the excretion of total pyridinoline crosslinks. In a short-term study on intravenous pamidronate treatment of Paget’s disease, no reduction of the excretion of free DPD, but a markedly decreased excretion of the peptide bound fraction, could be observed [58, 59]. This finding, however, has not been confirmed by others [60]. To circumvent the problem of the variable excretion of free pyridinolines, prior to immunoassay, a hydrolysis step may be performed such that the total DPD concentration (i.e. the sum of free and peptide-bound fractions) can be measured [60].

D. Collagen Peptides The main molecular sites involved in collagen crosslinking are the short non-helical peptides at both ends of the collagen molecule, termed amino- (N) and carboxy- (C) terminal telopeptides. In normal collagen, these telopeptides are each linked via pyridinium or, as shown more recently, pyrrole compounds to the helical portion of

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Figure 7 Schematic presentation of the type I collagen monomer showing the location of the CTX epitope and the G site susceptible to racemization/isomerization. Type I collagen consists of two α1 chains and one α2 chain. The intact C-telopeptide α1 chain consists of 26 amino acid residues (1193–1217). The lysine residue, K1208, can be involved in intermolecular crosslinking. An attack by a peptide backbone nitrogen of the glycine G1212 on the side chain carbonyl group of an adjacent aspartyl residue D1211 can result in the formation of a succinimide ring (A → B). The succinimide ring is prone to hydrolysis and racemization producing peptides and β-aspartyl peptides in both the D and L configurations. Racemization is thought to proceed primarily through the succinimide pathway (B → E), but other pathways as direct proton abstraction (A ↔ D and C ↔ F) may also contribute to the formation of D-aspartyl (reproduced with permission from reference [69]).

neighboring collagen molecules (Fig. 3) [28, 61–64]. During collagen breakdown, N- and C-terminal telopeptide fragments of various sizes, still attached to the helical portions of a nearby molecule by a pyridinium or pyrrole crosslink, are released into the circulation. The majority of these fragments are relatively small and readily cleared by the kidneys. It has been demonstrated that type I collagen molecules can undergo racemization and isomerization at the aspartic acid residue D1211 within the C-telopeptide [65–67]. As a consequence of these spontaneous, nonenzymatic posttranslational modifications, the CTX-I molecule may occur in one of four different forms: as a native linear peptide (αL), as the isomerized beta form (βL); as the racemized

linear peptide (αD); and finally as the isomerized and racemized beta form (βD) (Fig. 7) [65, 67, 68]. The processes of racemization and isomerization have been attributed to protein aging [65, 67–69]. Newly synthesized type I collagen is incorporated into the bone matrix exclusively in the αL form. There is evidence suggesting that the relative proportions at which age-related CTX forms occur in bone is βL > βD > αD. Measurement of the relative amounts of newly synthesized (αL) and agemodified CTX may possibly provide information about the age of resorbed bone and the metabolic activity of bone [65, 67–69]. For example, in children, the αL/βL CTX-I ratio is more than three-fold higher than in adults. In Paget’s disease, which is characterized by (focal) high

550 bone turnover (see Chapter 47), the ratio is even more elevated [65–67, 70]. In postmenopausal women a small increase in the αL/βL CTX-I ratio has been observed and it appears that the ratio may be an independent predictor of fracture risk [71, 72]. In contrast, the ratio decreases after bisphosphonate treatment [70]. Over the years, several immunoassays have been developed for the quantification of telopeptides derived from both termini of the type I collagen. 1. Carboxyterminal Crosslinked Telopeptides of Type I Collagen

a. Assay for the Detection of Large C-telopeptide Fragments of Type I Collagen in Serum (ICTP/CTX-MMP) The first assay to be developed for the measurement of collagen telopeptides in serum was a radioimmunoassay (RIA) for the C-telopeptide crosslink domain generated by metalloproteinase (MMP) treatment [73]. This assay is based on a polyclonal antibody against purified crosslinked peptides of 8.5 kDa (ICTP or CTX-MMP), prepared by digesting human bone collagen with bacterial collagenase, or with trypsin, and purified by reversedphase separations on HPLC [73]. The same material is also used for the calibration of the assay. The antigenic determinant detectable in serum requires a trivalent crosslink, including two phenylalanine-rich domains of the teleopeptide region of the α1 chain of type I collagen. Divalently and noncrosslinked peptides do not react with the antibody, nor do peptides isolated from skin. The assay also detects other crosslink forms such as DPD or pyrrole. Treatment with cathepsin K cleaves the telopeptide structure between the phenylalanine-rich region and the crosslink, and therefore abolishes immunoreactivity in the assay [74]. The intra- and interassay coefficients of variation are 5–8% and 6–9%, respectively [75, 76]. A nonspecific binding of approximately 10% has been reported [75, 76]. The detection limit is in the order of 0.34 µg/L [73]. The main mechanism of its removal from blood is via the kidney because of the small size of the CTX-MMP antigen. Glomerular filtration rates of ≤50 ml/min allow for a significant retention of this peptide [73]. Lipemic serum also appears to interfere with the assay. The assay appears to be sensitive for pathological bone resorption as seen in multiple myeloma, metastatic bone disease and other degradation processes involving hasty breakdown of bone and collagen type I. In contrast, serum ICTP levels have been reported to change rather subtly in postmenopausal osteoporosis [77]. However, other reports describe serum ICTP as a reliable and robust marker of bone resorption. For example, following oophorectomy (OE, n = 18), serum ICTP levels

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increased significantly at 7 days and peaked between 1 and 3 months post OE. Bone loss at the lumbar BMD at 6 months and at 12 months post OE were significantly correlated with serum ICTP, but not with changes in bone formation markers [78]. More recent data by Meier and coworkers, investigating a healthy male population from the Australian Dubbo study, have demonstrated serum ICTP levels to be a reliable predictor of future fracture risk in men [79]. b. Assay for the Detection of Small C-telopeptide Fragments of Type I Collagen in Urine and Serum (CTX-I) The initial ELISA for measurement of the crosslinked part of the C-terminal telopeptide region of type I collagen in urine was based on a polyclonal antibody directed against an octapeptide sequence specific for the C-telopeptide α-1 chain (Glu-Lys-Ala-His-βAsp-GlyGly-Arg) [80] (Fig. 3). The antibody shows specificity for an amino acid sequence common to all type I collagen molecules and recognizes only the isomerized (β) form (EKAHD-β-GGR, or β-CTX-I) [81]. This particular sequence of amino acids was chosen as the antigen because it contains the lysine of the C-telopeptide domain, which participates in intermolecular crosslinking. Due to the compact structure of this peptide, protection from further degradation when excreted in the urine was anticipated [80, 82]. The intra- and interassay coefficients of variation are 6–8% and 8%, respectively [80, 83, 84]. At low concentrations, however, the assay exhibits poor precision [84]. The detection limit is 0.2–0.5 µg/ml. A radioimmunoassay based on monoclonal antibodies recognizing the linear, nonisomerized octapeptide form (EKAHD-αD-GGR or α-CTX-I) of the same amino acid sequence has also been developed [85]. This assay shows less than 2% crossreactivity with the β-isomerized form (β-CTX-I). Intra- and interassay variations for this assay are 3–5% and 4–6.5%, respectively [85]. Both the α-CTX-I and the β-CTX-I assays only require one CTX-I chain to react, however the lysine residue within the CTX-I epitope can participate in interand intramolecular covalent crosslinks joining two epitopes [65]. Hence, the α-CTX-I RIA measures molecules composed of two C-telopeptide fragments, where one of the two epitopes is in α form, whereas the other epitope may be any of the two forms (α or β) [86]. Correspondingly, the urine β-CTX-I measures β-βCTX-I and β-αCTX-I fragments, in contrast to the assay measuring β-CTX-I in serum, which employs two β-CTX-I specific antibodies and therefore measures only β-βCTXI (see below). Recently an ELISA was developed using a specific antibody to α-CTX-I, measuring exclusively ααCTX-I with a detection limit of 0.21 ng/ml [86].

Chapter 33 Measurement of Biochemical Markers of Bone Resorption

The intra- and interassay variations for this ELISA are 3% and 9%, respectively. In addition to the assay recognizing the native (αL) and the isomerized (βL) forms of CTX-I, specific assays recognizing the racemized (αD) and the isomerized/ racemized (βD) isoforms of CTX-I have recently been developed [67]. Antisera were raised against the peptide conjugates αD-EKAHDGGR or βD-EKAHDGGR. Intraand interassay variations for the αD-CTX-I assay are 8.5% and 10.4%, respectively. Intra- and interassay variations for βD-CTX-I are 7.5% and 11.4%, respectively [67]. The detection limit is 1.07 ng/ml for αD-CTX-I, and 0.06 ng/ml for βD-CTX-I. In 1997, a sandwich assay for the measurement of β-CTX-I in serum was developed. Initially, a polyclonal antiserum against collagenase-digested collagen type I, together with the β-isomerized C-terminal octapeptide attached to the ELISA plate, was used [87]. The intraand interassay coefficients for this assay are 6–7% and 9%, respectively, the detection limit 10 ng/ml [87]. The newer assay uses two monoclonal antibodies, thus recognizing only the β-βCTX-I [88]. The intra- and interassay variations for this assay are 4.7–4.9% and 5.4–7.9%, respectively. This assay has also been adapted to an automated analyzer system [89, 90]. In 2001, Srivastava and coworkers decribed the development of an ELISA for the detection of C-telopeptide fragments of type I collagen in urine, using a rat monoclonal antibody against a synthetic linear peptide, DFSFLPQPPQEKAHDGGR, which contains a slightly different epitope than the CTX-I assay. This longer sequence was isolated from the urine of patients with Paget’s disease of bone. The measuring range of this ELISA was 10–1000 ng/ml with a detection limit of 10 ng/ml. Average intra- and interassay CVs were <7% and <11%, respectively. Studying healthy pre- and postmenopausal as well as osteoporotic women, the assay performed comparable to similar, commercially available assays for the C-telopeptide, and showed strong correlations with urinary DPD levels [91]. c. Assay for the Detection of N-telopeptide Fragments of Type I Collagen in Urine and Serum (NTX-I) In the early 1990s, a competitive inhibition ELISA was developed for the measurement of N-terminal crosslinked telopeptide (NTX-I) fragments in the urine [82]. This assay is based on a monoclonal antibody that specifically recognizes an epitope embedded in the α-2 chain of the N-telopeptide fragment (Fig. 3). The peptide has the sequence QYDGKGVG, where K is involved in a trivalent crosslinking site. The compound still contains the pyridinium crosslink, but the antibody does not recognize the pyridinoline or deoxypyridinoline per se. Thus, the

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antibody appears to react only with a conformational epitope, which is specific for the crosslinking of bone collagen and the presence of a pyridinium crosslink is not essential for reactivity [82]. Collagen must be broken down to small crosslinked peptides that contain the exact sequence before antibody binding occurs with the NTX-I antigen. Furthermore, the antibody also recognizes such peptides in culture medium conditioned by osteoclasts that are resorbing human bone particles in vitro [92, 93]. These data suggest that the NTX-I and peptide is a direct product of osteoclastic proteolysis and appears not to be metabolized further [82]. There is some contention about this specificity, however. Although the antibody recognizes a crosslink containing sequence from the α-2 chain N-telopeptide, the crosslink is not involved in the epitope, and cross-reaction with skin-derived peptides has been reported [61, 94]. The NTX-I assay is calibrated using standard amounts of human bone collagen digested with bacterial collagenases. The intra- and interassay variability, expressed as a coefficient of variation, is on average 8% and 10% respectively, or higher at low concentration, and the sensitivity is 20–25 nM [83, 84, 95]. It has also been reported that the assay exhibits nonlinear recovery for sample dilutions prepared in assay buffer. This nonlinear recovery, however, could be corrected for by sample dilutions made in urine at low analyte concentrations [84]. This assay requires no hydrolysis or pretreatment of the urine. Generally, measurement is performed in second morning spot urine. Results are reported as bone collagen equivalents (BCE), in nM, corrected for creatinine excretion to compensate for differences in urine dilution. An assay for the measurement of type I collagen N-telopeptides in serum had been developed in the late 1990s [96]. Experimental and clinical data demonstrated this assay to provide a useful index of bone resorption, however, mostly for technical reasons, the serum assay has not become as widely used as the assay for NTX-I in urine [97–107]. More recently, a point of care device (POCD) for NTX has been developed. This device provides rapid results, shows a reasonable correlation with the conventional immunoassay, but has a higher degree of variability [108]. d. Assay for the Detection of Type I Collagen Helical Peptide in Urine (HELP) Recently, a new monoclonal antibody-based ELISA detecting a fragment of the helical part of type I collagen in urine has become available. The antibody recognizes an epitope localized to amino acids 620–633 of the collagen type I alpha1 chain. Pilot studies in healthy pre- and postmenopausal women demonstrated an intra- and interassay variability of

552 7.3 and 8.7%, respectively. The long-term intraindividual variability assessed over 6 months in 18 untreated postmenopausal women was 24%. In postmenopausal women, levels of the urinary helical peptide were 42% higher than in premenopausal women. This change was very similar to the increase seen in urinary CTX-I levels (47%) [109]. While this new assay appears to demonstrate adequate analytical performance, very similar results have been obtained with established markers of bone resorption. Further studies are therefore required in order to determine whether measurement of the helical peptide in urine confers any advantage over existing and well-researched assays.

E. Collagen-related Markers of Bone Resorption: Pre-analytical Issues Most, if not all, collagen-related markers of bone resorption are stable to normal conditions of sample handling. Both the free and conjugated forms of PYD and DPD have been shown to be stable in urine samples kept at room temperature for several weeks [30, 110]. Several reports show that pyridinium crosslinks can be stored at —20oC for years [30, 110–112]. Repeated freezing and thawing of urine samples up to ten times had no effect on the concentrations of the pyridinolines [110]. However, pyridinium crosslinks are unstable when subjected to intensive UV irradiation. After exposure to natural (UV) light for 3 hours, pyridinolines in an aqueous solution were rapidly degraded, while no changes were noted in the concentrations of PYD and DPD present in urine [113–115]. The fact that photolysis was more pronounced in aqueous solutions than in urine may be attributable to the higher intensity of UV light in the clear aqueous solution. The rate of pyridinoline photolysis is also pH dependent (the rate increases with increasing pH) [113], and the sensitivity to UV light seems to be greater for free pyridinoline than for total pyridinoline [115]. Photolysis of the pyridinoline standard solution is prevented by storage in amber vials [114], Urine samples meant to be analyzed for PYD or DPD should be placed in the dark within 2 hours of collection [113]. The telopeptides NTX-I and CTX-I are not affected by UV light exposure [115]. The N- and C-terminal telopeptides are usually stable at room temperature. However, the large serum C-terminal telopeptide, ICTP, has been described to lose up to 12% in immunoreactivity when stored at room temperature for 5 days [73, 80, 84]. Most collagen-related markers of bone resorption show significant diurnal variation [31, 75, 84, 116–125] (see also Chapter 34). Although some authors have reported diurnal changes as high as 200% of the daily mean

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(e.g. s-CTX-I) [126], most studies found amplitudes between ±15–30%. Diurnal variation appears not to be affected by age, menopause, bed rest, physical activity, or cortisol production [119, 124, 127, 128]. Although in postmenopausal women bone turnover is higher than in premenopausal women, the circadian variation is similar for both pre- and postmenopausal women and, thus, is not influenced by sex hormones [119, 121]. In osteoporotic women, it has been suggested that the increase in bone resorption at night may persist longer into the morning and that this may account in part for the greater bone loss in osteoporotic women [116]. However, this could not be confirmed for elderly women with osteopenia [119]. The circadian rhythm also does not appear to be influenced by seasonal changes [129]. The only factor with a pronounced influence on circadian variation seems to be fasting, which can significantly reduce the diurnal variation of some markers by about one fourth [124]. Thus, it has been reported that food intake strongly affects the serum levels of CTX-I, but not those of NTX-I or ICTP [130]. The etiology of these diurnal variations remains unknown. Many hormones such as parathyroid hormone, growth hormone, and cortisol exhibit circadian variations, and may therefore be implicated in the daily ups and downs of bone turnover [131–134]. There is some evidence that gastrointestinal hormones released following food intake are involved in the variation of CTX-I serum levels. Intake of glucose, protein or fat resulted in a decrease in bone resorption independent of gender or menopausal status [128]. Part of this response appears to be due to variation in serum insulin or intestinal proglucagonderived peptides such as glucagon-like peptide-2 (GLP-2). A recent study in humans found that parenteral administration of GLP-2 caused a statistically significant and dose-dependent reduction in serum CTX-I levels, as compared to baseline and placebo. No such effect was observed when glucose-dependent insulinotropic polypeptides (GIP) or GLP-1 were administered [135, 136]. It has been recommended to perform measurements of collagen-related bone resorption markers in fasting spot samples, either as a first or second morning void, and to correct the result for urinary creatinine. Alternatively, the excretion rate of the marker may be determined in a timed 24-hour urine collection. Both approaches are subject to some difficulties. Creatinine output has been reported to be fairly constant with time (variations within 10%) and to correlate with lean body mass [137], but there are also reports suggesting that the correction for creatinine in a urine spot sample could be misleading. On the other hand, timed 24-hour urine collections are subject to inevitable inaccuracies due to collection errors.

Chapter 33 Measurement of Biochemical Markers of Bone Resorption

In general, similar results are obtained from either 24hour, 2-hour, or spot urine collections. It should be borne in mind however, that the slope of diurnal changes is steepest during the morning hours (see diurnal variation), which is usually the time at which urine samples are collected. Therefore, if spot urine is used for measurement of bone resorption markers, the time of collection should be controlled as closely as possible. In addition to diurnal variations, most markers of bone resorption exhibit considerable variations between consecutive days [30, 41, 84]. This intraindividual, dayto-day variability has been studied extensively for the collagen-related markers and is apparently due to genuine variations in marker levels and not to analytical imprecision. Expressed as a coefficient of variation, the day-to-day variation in crosslink excretion measured by HPLC was 16–26% [30, 31]. A similar variability has been reported for the free pyridinolines (7–25%), NTX-I (13–35%), and serum (8–10%) or urinary CTX-I (12–35%) [84, 138–142]. In general, serum markers show less day-to-day variability than urine markers, which may be related to the introduction of an additional test (urine creatinine) and/or to the difficulties associated with collecting accurately timed urine samples. Mild to moderate renal impairment appears to have no significant influence on total excretion of the pyridinoline crosslinks, and urinary values usually do not change until a creatinine clearance of <25 ml/min is reached [31, 57, 143]. In contrast, the serum levels of the C-terminal telopeptide ICTP are strongly affected by renal function and tend to increase with a glomerular filtration rate of 50 ml/min or less [73]. Urinary CTX-I is probably mostly cleared through glomerular filtration and therefore increased in end-stage renal failure [125].

III. NON-COLLAGENOUS PROTEINS OF THE BONE MATRIX A. Bone Sialoprotein Bone sialoprotein (BSP) is a glycosylated protein with an apparent MW of 70–80 kDa (core protein: 34 kDa) that accounts for approximately 5–10% of the noncollageneous proteins of the bone extracellular matrix [144, 145]. High amounts of BSP are expressed in active osteoblasts and, to a lesser degree, in osteoclast-like cell lines [146–148]. Compared to other noncollageneous proteins, the tissue distribution of BSP is relatively restricted; the protein or its mRNA is detected mainly in mineralized tissues such as bone and dentin, and at the interface of calcifying cartilage and bone. BSP contains an Arg-Gly-Asp (RGD)

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integrin recognition sequence, improves the attachment of osteoblasts and osteoclasts to surfaces, binds preferentially to the alpha2-chain of collagen type I, nucleates hydroxyapatite crystal formation in vitro and appears to enhance osteoclast-mediated bone resorption [149–151]. The protein is therefore considered to play an important role in cell–matrix adhesion processes and in the supramolecular organization of the extracellular matrix of mineralized tissues. Interestingly, BSP-related immunoreactivity has also been demonstrated in placental trophoblasts, platelets [152], and certain malignancies such as breast, prostate, and thyroid cancer, or multiple myeloma [153]. All of these tumors metastasize to bone, and recent in vitro and animal studies have demonstrated that transfection of MDA-MB-231 human breast carcinoma cells with BSP stimulates migration and invasion of cancer cells in vitro and growth of primary and secondary tumors in nude mice. Therefore, BSP appears to play an important tole in the metastasis of ceratin cancers to bone [154, 155]. Vice versa, decreased levels of BSP were shown to correlate with reduced proliferation, colony formation, and migration of MDA-MB-231 cancer cells [156]. Several immunoassays for the determination of bone sialoprotein in serum have been developed [157–159]. All of these assays are based upon polyclonal antisera against BSP, although attempts have been made to develop monoclonal-based assays [160]. In terms of normal ranges, biological and pathological variability, and response to osteotropic treatment, most experience has been collected using a homologous radioimmunoassay [159, 161]. This assay uses bone sialoprotein isolated from human bone by standard extraction procedures and final purification by wide pore reversed-phase HPLC. The anti-BSP antibodies against human BSP were raised in chicken. In this particular assay, the coefficients of variation were 6.1–7.0% for intra-assay variability, and 9.2–9.4% for interassay variability. The lower detection limit is 0.7 ng/ml, and recovery after spiking of human samples with purified BSP ranged between 92–108%. Importantly, no cross-reactivity was observed for a number of noncollageneous protein, such as osteocalcin, osteopontin, and osteonectin, or with bone alkaline phosphatase [159] (Fig. 8). Employing the aforementioned radioimmunoassay, normal values for serum BSP ranged between 4.0 and 23.5 ng/ml (mean 11.8 ± 4.6 ng/ml). In healthy adults, serum BSP values were generally found unrelated to body measures, liver or kidney function, or to lumbar bone density. However, serum BSP levels were significantly lower in premenopausal women than in males or postmenopausal females. Interestingly, in postmenopausal

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Figure 8

Crossreactivity of anti-BSP antibody with noncollagenous proteins (䊉 bone sialoprotein (BSP), 䉬 bone alkaline phosphatase, 䊏 osteocalcin, 䉲 osteonectin) (reproduced from reference [159]).

females, serum BSP levels were correlated with chronological age (r = 0.41, P < 0.001) (Fig. 9) but not with age at menopause or with the duration of menopause. In contrast, no relation with age was seen in males and premenopausal females [161]. So far, the published literature indicates that serum BSP reflects processes associated with osteoclast activity and bone resorption, rather than osteoblast activity. Serum BSP levels appear to be of particular interest in malignant diseases involving bone, where they seem to be associated with tumor cell burden. Thus, depending on the disease stage, serum BSP is greatly elevated in patients with multiple myeloma, where BSP levels correlated with the bone marrow plasma cell content and serum beta2-microglobulin. After chemotherapy, BSP reflected the response to treatment and correlated with the change in monoclonal protein. Patients with multiple myeloma and normal baseline BSP levels survived longer than patients with initially elevated BSP values, and

Figure 9

Serum bone sialoprotein (BSP) levels in healthy female subjects according to menopausal status (reproduced from reference [161]).

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serum monoclonal protein and BSP were independent predictors of survival [162]. Similar observations have been made in a recent study on prostate cancer, where serum BSP was shown to be independent of prognostic factors for cancer-associated mortality [163]. In patients with hypercalcemia of malignancy, treatment with intravenous bisphosphonates results in a rapid fall of serum BSP values, paralleling the changes seen in urinary crosslink concentrations [159, 164]. Compared to healthy perimenopausal controls, serum BSP levels were found to be significantly elevated in patients with postmenopausal osteoporosis. In this study, serum BSP levels were positively correlated with other markers of bone resorption such as PYD, DPD, and NTX-I, and with cytokines (IL-11, TGFbeta2) involved in bone turnover. Serum BSP levels decreased during antiresorptive treatment and this decrease paralleled the change in other bone resorption markers [160]. Bone sialoprotein was found to be significantly increased in patients with active Crohn’s disease [165]. As seen with many other biomarkers of bone metabolism, serum BSP shows typical changes related to biological influences, such as diurnal, day-to-day, and menstrual variability, as well as changes with somatic growth and season [123, 164]. In patients with renal failure, serum BSP levels tend to increase, and in this group highest values are seen in patients with renal insufficiency and secondary hyperparathyroidism [161]. Following phlebotomy, BSP levels remain stable for several hours both at room temperature and at −20°C, and have been shown not to change significantly during repeated freeze–thaw cycles.

B. Osteocalcin Fragments in Urine Osteocalcin (OC) is a small noncollagenous protein synthesized by mature osteoblasts, odontoblasts, and hypertrophic chondrocytes. Osteocalcin measured in serum is considered a sensitive index of osteoblastic activity and thus of bone formation (see Chapter 32). Serum osteocalcin is usually measured by immunoassay, although with considerable inconsistency of results between various assays. These discrepancies in osteocalcin values obtained from assays using different antibodies are thought to be due to the presence of a number of different OC fragments in serum [166]. Since OC is incorporated into the bone matrix upon bone formation, it has long been argued that proteolytic fragments of this peptide should be released during bone matrix resorption. Thus, depending on the assay used, measurement of OC in serum would capture not only

Chapter 33 Measurement of Biochemical Markers of Bone Resorption

those molecules derived from osteoblasts, but also OC fragments released during bone resorption. More recently, several groups have been investigating the significance of OC fragments in urine as a marker of bone resorption. Using mass spectrometry and N-terminal sequencing, Ivaska and coworkers have found multiple proteolytic fragments of osteocalcin in human urine [167]. Most of these fragments contained less than 30 residues and originated from the mid-molecular region of the intact OC molecule. Interestingly, levels of OC fragments inversely correlated with bone mineral density in elderly women. The same group later demonstrated that in vitro osteocalcin is released from bone slices during bone resorption both in intact and fragmented form [168]. Another study, utilizing a monoclonal antibody-based inhibition ELISA to quantify fragments derived from the OC mid-region (residues 21–29) in bone cell culture supernatants, demonstrated that the release of OC fragments was closely associated with osteoclast-mediated pit formation. When the same assay was used to detect mid-OC fragments in urine, values decreased following antiresorptive treatment with bisphosphonates. Further characterization of the fragments found in urine demonstrated most of the antigenic material to be of small size (< 15 amino acids in length) and to contain modified aspartyl residues typical of aged proteins [169]. Recent clinical studies suggest that the quantification of specific OC fragments in urine may provide an index of bone resorption [170, 171].

IV. OSTECLAST ENZYMES A. Tartrate-Resistant Acid Phosphatase The acid phosphatases are a heterogenous group of enzymes that hydrolyze phosphomonoesters at low pH with the release of phosphoric acid. There are five isoenzymes of acid phosphatase in serum, the major sources of which are bone, prostate, platelets, erythrocytes, and the spleen [172] (see also Chapter 10 [173–175]). These isoenzymes differ in molecular size, sensitivity towards various inhibitors, and electrophoretic mobility in acidic native polyacrylamic gels [172–174]. According to their relative electrophoretic mobility, serum acid phosphatase isoenzymes have been classified as type 0, 1, 2, 3a, 4, 5a, and 5b isoenzymes [174, 175]. Based on their sensitivity to inhibition by L(+)-tartrate, serum acid phosphatase isoenzymes are also divided into two classes, namely tartrate-sensitive (TSAP) and tartrate-resistant acid phosphatase (TRACP). However, one should bear in mind that the susceptibility to L(+)-tartrate is not completely tissue

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specific [172–175]. The only TRACP isoenzymes present in serum are the type 5a and 5b acid phosphatases. The two isoforms 5a and 5b are different in that 5a contains sialic acid, whereas 5b does not. The skeleton is rich in acid phosphatase activity [176–178]. While osteoclasts show the greatest abundance in type 5b TRACP, some activity has also been shown in osteoblasts and osteocytes [179–181]. Generally, TRACP is used as a cytochemical marker to distinguish osteoclasts from other bone cells [182–184]. In serum, TRACP can be measured either by spectrophotometric techniques or, more recently, by immunoassay. A variety of spectrophotometric and kinetic assays for serum TRACP have been developed [173, 174, 185–194]. These assays differ in many aspects, including the type and concentration of substrate, assay buffer, and pH of the reaction. The most widely used kinetic assay measures the hydrolysis of p-nitrophenol phosphate in the presence of sodium tartrate [185]. This assay takes into account several possible interferences. To minimize the interferences of noncompetitive inhibitors, the serum samples are diluted five-fold and the substrate concentration is increased. The diluted serum samples are then preincubated at 37oC for 1 hour before assay to inactivate erythrocytic TRACP activity released by hemolysis. To minimize the effect of platelet-derived TRACP activity, which may be released during clotting, the clotting time should be kept to a minimum, i.e. 1–2 hours at room temperature [185]. The reported intra- and interassay variations are 5% and 10% respectively and the recovery of exogenous added purified TRACP activity to human serum is 110 ± 10% [185]. The major drawback of the kinetic assays currently in use is the fact that these methods do not distinguish between TRACP activity derived from osteoclasts or from other sources. Consequently, they lack specificity and sensitivity for the osteoclastic isoenzyme. To improve the specificity and sensitivity of serum TRACP measurements, several immunoassays have been recently developed. The antibodies for these assays were against material isolated from spleen of patients with hairy-cell leukemia, against 5b-TRACP isolated from chord plasma, or more recently from human bone [195–198]. The first reported immunoassays used an immunoprecipitation approach based upon a polyclonal antibody directed against a partially purified osteoclastic-like 5b TRACP recovered from the spleen of a patient with hairy-cell leukemia, or alternatively, antibodies against porcine uteroferrin, which resembles osteoclastic TRACP, were used [195–197]. TRACP activity of the immune complex was then determined using a kinetic assay. However, the specificity of the antisera employed in these

556 immunoprecipitation assays has not been evaluated, and it seems of further disadvantage that serum TRACP “levels” are still determined on the basis of the enzyme activity. Immunoassays measuring protein concentration rather than enzyme activity have been developed [198–202]. Thus, an ELISA for serum type 5b osteoclastic TRACP using polyclonal antibodies against partially purified hairy cell leukemia spleen type 5b TRACP has been described [199]. The antibody reacted with the lysosomal type 5b TRACP in osteoclasts, but showed no crossreactivity with partially purified acid phosphatases from normal human spleen, prostatic acid phosphatase, or acid phosphatase extracted from osteoblasts [199]. Intra- and interassay variations are both less than 10% for this assay. Another ELISA is based on polyclonal antibodies against human cord plasma TRACP [198]. The antibody used in this assay crossreacted with extracts of bone, but not with extracts obtained from spleen, erythrocytes, platelets, prostate, or osteoblasts. Intra- and interassay variations are reported less than 12.5%, analytical recovery varied from 94–106%. In another ELISA-purified TRACP isolated from cord, plasma was used as antigen [198]. The antiserum crossreacted with an extract of bone, but not with extracts of spleen, erythrocytes, platelets, osteoblasts, or with prostatic acid phosphatase. The interassay variation for this assay is 8.5%. During the past years, the first assays based on polyand monoclonal antibodies against TRACP purified from human bone have been described [201, 203, 204]. The polyclonal antisera were employed in a competitive fluorescence immunoassay and were shown to immunohistochemically recognize osteoclasts from human bone, and alveolar macrophages from human lung tissue, but not cells from human spleen tissue [203]. The two monoclonal antibodies raised against human osteoclastic type 5b TRACP were found to recognize different epitopes and therefore allowed the development of a two-site fluorescence immunoassay [201]. In this latter assay, the intra- and interassay coefficients of variation are both less than 5%, analytical recovery is 98.9 ± 3.3%, and a sensitivity of less than 0.1 µg/L has been demonstrated. However, all these assays measure total TRACP (5a and 5b) in serum. A new assay was developed using monoclonal antibodies to bind total TRACP (both 5a and 5b) to microtiter plate and then to measure bound enzyme activity using pNPP as substrate. The best pH value to detect specifically 5b TRACP was 6.1, at this pH 5a TRACP is almost inactive and 5b is almost as active as in the optimum range (5.7–5.9) [204]. The antibody binds to enzymes

Marius E. Kraenzlin and Markus J. Seibel

from erythrocytes and platelets and only recognized 32-kDa type 5 TRACP band from human bone and human serum in Western analysis [204]. Intra- and interassay variations are 3.2% and 6.9%, analytical sensitivity 0.06 U/L [125, 204]. Using this assay, within-subject variability of TRACP5b was reported as 6.6%, which is lower than that of many other urinary and serum telopeptide markers. Also, diurnal variation for TRACP 5b appeared to be minimal compared to that of bCTX in serum (14% vs. 137%), with day-today variability around 10–12%. Furthermore, in contrast to serum CTX-I, TRACP 5b was not affected by food intake or hemodialysis. The absolute response to antiresorptive therapy was less pronounced with serum TRACP 5b than with other markers of bone resorption, but the signal-to-noise ratio was better for serum TRACP5b than for CTX-I [125]. TRACP levels correlate well with histological indices of osteoclasts [205]. TRACP activity measured by immunoassay appears to be stable up to 8 hours, whereas for the conventional kinetic assays, the enzyme loses more than 20% of its activity per hour at room temperature [185, 195, 204, 206]. The addition of citrate buffer to the sample can stabilize the enzyme. TRACP activity declines slowly during frozen storage at −20oC, but retains its activity for up to 1 year when stored at −70o C or lower [185, 204]. In our own hands, up to three freeze–thaw cycles did not appear to have a significant effect on TRACP serum activity [122], but repeated freeze–thaw cycles should be avoided. Serum TRACP accumulates in renal failure [207]. Taken together, serum TRACP5b is a marker of bone resorption that carries great promise for routine clinical application. It has the added advantage of reflecting osteoclast, i.e. actual cellular activity, rather then collagen degradation, which is always secondary to this activity.

B. Cathepsins The name cathepsin is generic for endopeptidases and exopeptidases active at acidic pHs. Cathepsin K is a member of the cysteine protease family that, unlike other cathepsins, has the unique ability to cleave both helical and telopeptide regions of collagen I. Its clinical relevance was only appreciated in 1996, when Gelb et al. discovered that pycnodysostosis (osteopetrosis and short stature) was the result of nonsense, missense, and stop codon mutations in the cathepsin K gene [208]. Another key observation was that the cathepsin K null mouse demonstrates osteopetrosis [209, 210]. Immunocytochemical studies have confirmed the localization of cathepsin K in osteoclasts being

Chapter 33 Measurement of Biochemical Markers of Bone Resorption

extracellularly along the bone and cartilage resorption lacunae, and intracellularly in vesicles, granules, and vacuoles throughout the cytoplasm of multinucleated osteoclasts attached to the surface of bone and cartilage [211]. Therefore, cathepsin K is functionally involved in osteoclastic bone resorption, and most likely a key player in normal bone matrix degradation/bone resorption. Recently, an enzyme-linked immunosorbant assay for cathepsin K measurement in serum and cell culture (Cathepsin K ELISA BI-20432, Biomedica, Austria) has been developed using a polyclonal sheep antibody. So far, only preliminary clinical studies in healthy men and preand postmenopausal women, as well as in patients with postmenopausal osteoporosis or Paget’s disease of bone are available [212]. Mean (±SD) serum cathepsin K values in healthy subjects ranged between 3.20 ± 1.41 pmol/L and 2.65 ± 1.29 pmol/L, respectively. Mixed-effect analysis of variance suggested that the intrasubject variance of cathepsin K was 1.81 ± 0.90 pmol/L of which biological and interassay variance accounted for 68% (1.28 ± 0.87 pmol/L) and analytical variance accounted for 32% (0.58 ± 0.28 pmol/L). Serum cathepsin K levels were not affected by food intake and did not differ between the healthy men and women. In contrast, women with postmenopausal osteoporosis (11.3 ± 13.1 pmol/L, P = 0.01) and patients with Paget’s disease (6.2 ± 4.4 pmol/L, P = 0.01) had significantly higher cathepsin K levels at baseline as compared to controls matched for age and menopausal status. In patients with osteoporosis, bisphosphonate treatment resulted in a significant 37.8% decrease in serum cathepsin K levels, with nadir levels at 1 month [212]. Another recent study evaluated circulating cathepsin L levels in patients with low bone density and/or osteoporosis. Cathepsin L was measured using a commercially available enzyme-linked immunosorbent assay, and serum levels were compared to other bone markers such as bonespecific ALP, osteocalcin, or NTX-I [213]. Patients with osteoporosis had significantly higher serum levels of cathepsin L. Treatment with antiresorptive agents significantly reduced serum cathepsin L levels. Taken together, these results indicate that measurement of cathepsin K or L in serum holds promise as further cellular markers of osteoclast activity with potential for clinical use.

References 1. Prockop, D. J., and Kivirikko, K. I. (1984). Heritable diseases of collagen. N. Engl. J. Med. 311, 376–386. 2. Smith, R. (1980). Collagen and disorders of bone. Clin. Sci. 59, 215–221.

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3. Woessner, J. F. (1961). The determination of hydroxyproline in tissue and protein samples containing small proportions of this imino acid. Arch. Biochem. Biophys. 93, 440–447. 4. Kivirikko, K. I., Laitinen, O., and Prockop, D. J. (1967). Modifications of a specific assay for hydroxyproline in urine. Analyt. Biochem. 19, 249–255. 5. Podenphant, J., Larsen, N. E., and Christiansen, C. (1984). An easy and reliable method for determination of urinary hydroxyproline. Clinica. Chimica. Acta. 142, 145–148. 6. Sela, B. A., and Doolman, R. (1991). Refinement of a specific assay for hydroxyproline in urine. Clin. Chem. Acta. 203, 91–94. 7. Hodgkinson, A., and Thompson, T. (1982). Measurement of the fasting urinary hydroxyproline: creatinine ratio in normal adults and its variation with age and sex. J. Clin. Pathol. 35, 807–811. 8. Mautalen, C. A. (1970). Circadian rhythm of urinary total and free hydroxyproline excretion and its relation to creatinine excretion. Cli. Experimental. 75, 11–18. 9. Scalella, P. (1982). Interference by iodinated contrast media in total urinary hydroxyproline measurement. Lancet, 1(8277), 907. 10. Reed, P., Holbrock, I. B., Gardner, M. L. G., and McMurray, J. R. (1991). Simple, optimized liquid-chromatographic method for measuring total hydroxyproline in urine evaluated. Clin. Chem. 37, 285–290. 11. Casari, E., Ferrero, C. A., Grazioli, V., and Murone, M. (1992). Performance of a modified HPLC method to determine total hydroxyprolinuria in 2-h urine samples. Clin. Chem. 38, 2337–2338. 12. Dawson, C. D., Jewett, S., and Driskoll, W. J. (1988). Liquidchormatographic determination of total hydroxyproline in urine. Clin. Chem., 34, 1572–1574. 13. Cunningham, L. W., Ford, J. D., and Segrest, J. P. (1967). The isolation of identical hydroxylysyl glycosides from hydrolysates of soluble collagen and from human urine. J. Biol. Chem. 242, 2570–2571. 14. Krane, S. M., Kantrowitz, F. G., Byrns, M., Pinnell, S. R., and Singer, F. R. (1977). Urinary excretion of hydroxylysine and its glycosides as an index of collagen degradation. J. Clin. Invest. 39, 819–827. 15. Askenasi, R. (1975). Urinary excretion of freee hydroxylysine, peptide-bound hydroxylysine and hydroxylysyl glycosides in physiological conditions. Clin. Chim. Acta. 59, 87–92. 16. Segrest, J. P., and Cunningham, L. W. (1970). Variations in human urinary O-hydroxylysyl glycoside levels and their relationship to collagen metabolism. J. Clin. Invest. 49, 1497–1509. 17. Krane, S. M., Kantrowittz, F. G., Byrne, M., Pinnell, S. R., and Singer, F. R. (1977). Urinary excretion of hydroxylysine and its glycosides as an index of collagen degradation. J. Clin. Invest. 59, 819–827. 18. Moro, L., Modricky, C., Stagni, N., Vittur, F., and Bernard, B. (1984). High-performance liquid chromatographic analysis of urinary hydroxylysyl glycosides as indicator of collagen turnover. Analyst 109, 1621–1622. 19. Moro, L., Battista, C., Modricky, C., Rovis, L., and de Bernard, B. (1989). High-performance liquid chromatographic preparation of galactosyl-hydroylysine, a specific bone collagen marker. J. Chromat. 490, 285–292. 20. Moro, L., Modricky, C., Rovis, L., and de Bernard, B. (1988). Determination of galactosyl hydroxylysine in urine as a means for the identification of osteoporotic women. Bone Min. 3, 271–276. 21. Moro, L., Gazzarrini, C., Modricky, C., Rovis, L., de Bernard, B., Galligioni, E., Crivellari, D., Morassut, S., and Monfardini, S. (1990). High predictivity of galactosyl-hydroxylysine in urine as an indicator of bone metastases from breast cancer. Clin. Chem. 36, 772–774. 22. Leigh, S. D., Ju, H. S., Lundgard, R., Daniloff, G. Y., and Liu, V. (1998). Development of an immunoassay for urinary galactosylhydroxylysine. J. Immunol. Methods 220, 169–178. 23. Blumsohn, A., Hannon, R. A., and Eastell, R. (1995). Biochemical assessment of skeletal activity. Phys. Med. and Rehab. Clin. North Am. 6, 483–505.

558 24. al-Dehaimi, A. W., Blumsohn, A., and Eastell, R. (1999). Serum galactosyl hydroxylysine as a biochemical marker of bone resorption. Clin. Chem. 45, 676–681. 25. Bettica, P., Moro, L., Robins, S. P., Taylor, A. K., Talbot, J., Singer, F. R., and Baylink, D. J. (1992). Bone-resorption markers galactosyl hydroxylysine, pyridinium crosslinks, and hydroxyproline compared. Clin. Chem. 38, 2313–2318. 26. Moro, L., Mucelli, R. S., Gazzarrini, C., Modricky, C., Marotti, F., and de Bernard, B. (1988). Urinary beta-1-galactosyl-0-hydroxylysine (GH) as a marker of collagen turnover of bone. Calc. Tiss. Int. 42, 87–90. 27. Robins, S. P. (1983). Cross-linking of collagen. Isolation, structural characterization and glycosylation of pyridinoline. J. Biochem. 215, 167–173. 28. Eyre, D. R., Paz, M. A., and Gallop, P. M. (1984). Cross-linking in collagen and elastin. Ann. Rev. Biochem. 53, 717–748. 29. Delmas, P. D., Gineyts, E., Bertholin, A., Garnero, P., and Marchand, F. (1993). Immunoassay of pyridinoline crosslink excretion in normal adults and in Paget’s disease. J. Bone Min. Metab. 8, 643–648. 30. Colwell, A., Russell, R. G. G., and Estell, R. (1993). Factors affecting the assay of urinary 3-hydroxy pyridinium crosslinks of collagen as markers of bone resorption. Europ. J. Clin. Invest. 23, 341–349. 31. McLaren, A. M., Isdale, A. H., Whitings, P. H., Bird, H. A., and Robins, S. P. (1993). Physiological variations in the urinary excretion of pyridinium crosslinks of collagen. Br. J. Rheumatol. 32, 307–312. 32. Colwell, A., and Eastell, R. (1996). The renal clearance of free and conjugated pyridinium cross-links of collagen. J. Bone Miner. Res. 11, 1976–1980. 33. Huber, F., Traber, L., Roth, H. J., Heckel, V., and Schmidt-gayk, H. (2003). Markers of bone resorption – measurement in serum, plasma or urine? Clin. Lab. 49, 203–207. 34. Black, D., Duncan, A., and Robins, S. P. (1988). Quantitative analysis of the pyrdinium crosslinks of collagen in urine using ion-paired reversed-phase high-performance liquid chromatography. Anal. Biochem. 169, 197–203. 35. Uebelhart, D., Gineyts, E., Chapuy, M.-C., and Delmas, P. D. (1990). Urinary excretion of pyridinium crosslinks: A new marker of bone resorption in metabolic bone disease. Bone Mineral 8, 87–96. 36. Pratt, D. A., Daniloff, Y. N., Duncan, A., and Robins, S. P. (1992). Automated analysis of the pyridinium crosslinks of collagen in tissue and urine using solid-phase extraction and reversed-phase highperformance liquid chromatography. Anal. Biochem. 207, 168–175. 37. Robins, S. P., Stead, D. A., and Duncan, A. (1994). Precautions in using internal standard to measure pyridinoline and deoxypyrinoline in urine. Clin. Chem. 40, 2322–2323. 38. Bank, R. A., Beekman, B., Verzijl, N., de Roos, J. A., Sakkee, A. N., and TeKoppele, J. M. (1997). Sensitive fluorimetric quantitation of pyridinium and pentosidine crosslinks in biological samples in a single high-performance liquid chromatographic run. J. Chromatogr. B. Biomed. Sci. Appl. 703, 37–44. 39. Vesper, H. W., Audain, C., Woolfitt, A., Ospina, M., Barr, J., Robins, S. P., and Myers, G. L. (2003). High-performance liquid chromatography method to analyze free and total urinary pyridinoline and deoxypyridinoline. Analyt. Biochem. 318, 204–211. 40. Calabresi, E., Lasagni, L., Franceschelli, F., Bartolini, L., Serio, M., and Brandi, M. L. (1994). Use of an internal standard to measure pyridinoline and deoxypyridinoline in urine. Clin. Chem. 40, 336–337. 41. Jensen, J. B., Kollerup, G., Sorensen, A., and Sorensen, O. H. (1997). Intraindividual variability of bone markers in the urine. Scand. J. Clin. Lab. Invest. 57, 29–34. 42. Vesper, H. W., Smith, S. J., Audain, C., and Myers, G. L. (2001). Comparison study of urinary pyridinoline and deoxypyridinoline. Clin. Chem. 47, 2029–2031. (GENERIC) Ref Type: Generic

Marius E. Kraenzlin and Markus J. Seibel 43. Robins, S. P., Duncan, A., Wilson, N., and Evans, B. J. (1996). Standarization of pyridinium crosslinks, pyridinoline and deoxypyridinoline, for use as biochemical markers of collagen degradation. Clin. Chem. 42, 1621–1626. 44. Waelchli, R., Beerli, C., Meigel, H., and Revesz, L. (1997). First total synthesis of the bone resorption markers deoxypyridinoline and hydroxypyridinoline. Chem. Lett. 7, 2831–2836. 45. Allevi, P., Longo, A., and Anastasia, M. (1999). Total synthesis of deoxypyridinoline, a biochemical marker of collagen turnover. Chem. Commun. 32, 559–560. 46. Seibel, M. Duncan, A., and Robins, S. P. (1989) Urinary hydroxypyridinium crosslinks provide indices of cartilage and bone involvement in arthritic disease. J. Rheumatol. 16, 964–970. 47. Seibel, M. J., Baylink, D. J., Farley, J. R., Epstein, S., Yamauchi, M., Eastell, R., Pols, H. A., Raisz, L. G., and Gundberg, C. M. (1997). Basic science and clinical utility of biochemical markers of bone turnover—a Congress report. Exp. Clin Endocrinol. Diabetes. 105, 125–133. 48. Peel, N. F. A., al-Dehaimi, A. W., Colwell, A., Russel, R. G. G., and Eastell, R. (1994). Sulfasalazine may interfere with HPLC assay of urinary pyridinium crosslinks. Clin. Chem. 40, 167–168. 49. Seyedin, S. M., Kung, V. T., Daniloff, Y. N., Hesley, R. P., Gomez, B., Nielsen, L. A., Rosen, H. N., and Zuk, R. F. (1993). Immunoassay for urinary pyridinoline: The new marker of bone resorption. J. Bone Miner. Res. 8, 635–641. 50. Gomez, B., Ardakani, S., Evans, B. J., Merrel, L. D., Jenkins, D. K., and Kung, V. T. (1996). Monoclonal antibody assay for free urinary pyridinium cross-links. Clin. Chem. 42, 1168–1175. 51. Robins, S. P., Woitge, H., Hesley, R., Ju, J., Seyedin, S., and Seibel, M. J. (1994). Direct enzyme-linked immunoassay for urinary deoxypyridiniloine as a specific marker for measuring bone resorption. J. Bone Miner. Res. 9, 1643–1649. 52. Seibel, M. J., Woitge, H., Auler, B., Kissling, C., and Ziegler, R. (1998). Urinary free deoxypyridinoline: comparison of a new automated chemiluminescence immunoassay with HPLC and ELISA techniques. Clin. Lab. 44, 129–135. 53. Rosano, T. G., Peaston, R. T., Bone, H. G., Woitge, H. W., Francis, R. M., and Seibel, M. (1998). Urinary free deoxypyridioline by chemiluminescence immunoassay: analytical and clinical evaluation. Clin. Chem. 44, 2126–2132. 54. Sarno, M., Powell, H., Tjersland, G., Schoendorfer, D., Harris, H., Adams, K., Ogata, P., and Warnick, G. R. (1999). A collection method and high-sensitivity enzyme immunoassay for sweat pyridinoline and deoxypyridinoline cross-links. Clin. Chem. 45, 1501–1509. 55. Sarno, M., Sarno, L., Baylink, D., Drinkwater, B., Farley, S., Kleerekoper, M., Lang, R., Lappe, J., Licata, A., McClung, M., Miller, P., Nattrass, S., Recker, R., Schwartz, E. N., Tucci, J. R., Wolf, S., Powell, H., Tjersland, G., and Warnick, G. R. (2001). Excretion of sweat and urine pyridinoline crosslinks in healthy controls and subjects with established metabolic bone disease. Clin. Chem. Lab. Med. 39, 223–228. 56. Kamel, S., Brazier, J., Neri, V., Picard, C., Samson, L., Desmet, G., and Sebert, J. L. (1995). Multiple molecular form of pyridinoline crosslinks excreted in human urine evaluated by chromatographic and immunoassay methods. J. Bone Miner. Res. 10, 1385–1392. 57. Seibel, M. J. (1997). Clinical use of pyridinium crosslinks. In: Calcium Regulating Hormones and Markers of bone Metabolism: Measurement and Interpretation. (H. Schmidt-gayk, et al, ed.), pp. 157–169. Clin. Lab. Publications, Heidelberg, Germany. 58. Garnero, P., Shih. W. J., Gineyts, E., Karpf, D. B., and Delmas, P. D. (1994). Comparison of new biochemical markers of bone turnover in late postmenopausal osteoporotic women in response to alendronate treatment. J. Clin. Endocrinol. Metab. 79, 1693–1700. 59. Randall, A. G., Kent, G. N., Garcia-Webb, P., Bhagat, C. I., Pearce, D. J., Gutteridge, D. H., Prince, R. L., Stewart, G., Stuckey, B., Will, R. K.,

Chapter 33 Measurement of Biochemical Markers of Bone Resorption

60.

61.

62.

63.

64.

65.

66.

67.

68.

69.

70.

71.

72.

73.

74.

Retallack, R. W., Price, R. I., and Ward, L. (1996). Comparison of biochemical markers of bone turnover in Paget disease treated with pamidronate and a proposed model for the relationships between measurements of different forms of pyridinoline cross-links. J. Bone Miner. Res. 11, 1176–1184. Blumsohn, A., Naylor, K. E., Assiri, A. M., and Eastell, R. (1995). Different responses of biochemical markers of bone resorption to bisphosphonate therapy in Paget disease. Clin. Chem. 41, 1592–1598. Knott, L. and Bailey, A. J. (1998). Collagen cross-links in mineralizing tissues: a review of their chemistry, function and clinical relevance. Bone, 22, 181–187. Hanson, D., and Eyre, D. R. (1996). Molecular site specificity of pyridinoline and pyrrole crosslinks in type I collagen of human bone. J. Biol. Chem. 247, 26508–26516. Knott, L., Tarlton, J. F., and Bailey, A. J. (1997). Chemistry of collagen cross-linking: Biochemical changes in collagen during the partial mineralization of turkey leg tendon. Biochem. J. 322, 535–542. Kuypers, R., Tyler, M., Kurth, L. B., Jenkins, I. D., and Horgan, D. J. (1992). Identification of the loci of the collagen-associated Ehrlich chromagen in type I collagen confirms its role as a trivalent crosslink. Biochem. J. 283, 129–136. Fledelius, C., Johnsen, A. H., Cloos, P. C., Bonde, M., and Qvist, P. (1997). Characterization of urinary degradation products derived from type I collagen. Identification of a beta-isomerized Asp-Gly sequence within the C-terminal telopeptide (alpha1) region. J. Biol. Chem. 272, 9755–9763. Garnero, P., Fledelius, C., Gineyts, E., Serre, C. M., Vignot, E., and Delmas, P. D. (1997). Decreased beta-isomerization of the C-terminal telopeptide of type I collagen alpha 1 chain in Paget’s disease of bone. J. Bone Miner. Res. 12, 1407–1415. Cloos, P. A., and Fledelius, C. (2000). Collagen fragments in urine derived from bone resorption are highly racemized and isomerized: a biological clock of protein aging with clinical potential. Biochem. J. 345, Pt 3, 473–480. Cloos, P. A., and Christgau, S. (2002). Non-enzymatic covalent modifications of proteins: mechanisms, physiological consequences and clinical applications. Matrix Biol. 21, 39–52. Cloos, P. A. C., Fledelius, C., Christgau, S., Engsig, M., Delmas, P., Body, J. J., and Garnero, P. (2003). Investigation of bone disease unsing isomerized and racemized fragments of type 1 collagen. Calcified Tissue Int. 72, 8–17. Garnero, P., Gineyts, E., Schaffer, A. V., Seaman, J., and Delmas, P. D. (1998). Measurement of urinary excretion of nonisomerized and betaisomerized forms of type I collagen breakdown products to monitor the effects of the bisphosphonate zoledronate in Paget’s disease. Arthritis Rheum. 41, 354–360. Reginster, J.-Y., Henrotin, Y., Christiansen, C., Gamwell-Henriksen, E., Bruyere, O., Collette, J., and Christgau, S. (2001). Bone resorption in post-menopausal women with normal and low BMD assessed with biochemical markers specific for telopeptide derived degradation products of collagen type 1. Calcif. Tissue Int. 69, 130–137. Garnero, P., Cloos, P., Sornay-Rendu, E., Qvist, P., and Delmas, P. D. (2002). Type I collagen racemization and isomerization and the risk of fracture in postmenopausal women: the OFELY prospective study. J. Bone Miner. Res. 17, 826–833. Risteli, J., Elomaa, I., Niemi, S., Novamo, A., and Risteli, L. (1993). Radioimmunoassay for the pyridinoline cross-linked carboxy-terminal telopeptide of type I collagen: A new serum marker of bone collagen degradation. Clin. Chem. 39, 635–640. Garnero, P., Ferreras, M., Karsdal, M. A., Nicamhlaoibh, R., Risteli, J., Borel, O., Qvist, P., Delmas, P. D., Foged, N. T., and Delaisse, J. M. (2003). The type I collagen fragments ICTP and CTX reveal distinct enzymatic pathways of bone collagen degradation. J. Bone Miner. Res. 18, 859–867.

559

75. Hassager, C., Risteli, J., Risteli, L., Jensen, S. B., and Christiansen, C. (1992). The diurnal variation in serum markers of type I collagen synthesis and degradation in healthy premenopausal women. J. Bone Miner. Res. 7, 1307–1311. 76. Eriksen, E. F., Charles, P., Melsen, F., Mosekilde, L., Risteli, L., and Risteli, J. (1993). Serum markers of type I collagen formation and degradation in metabolic bone disease: Correlation with bone histomorphometry. J. Bone Miner. Res. 8, 127–132. 77. Okabe, R., Inaba, M., Nakatsuka, K., Miki, T., Naka, H., Moriguchi, A., and Nishizawa, Y. (2004). Significance of serum CrossLaps as a predictor of changes in bone mineral density during estrogen replacement therapy; comparison with serum carboxyterminal telopeptide of type I collagen and urinary deoxypyridinoline. J. Bone Miner. Metab. 22, 127–131. 78. Yasumizu, T., Fukada, Y., and Hoshi, K. (1999). Changes in serum levels of type I collagen-related proteins after surgically induced menopause and correlations with bone loss in the lumbar spine. Endocr. J. 46, 337–343. 79. Meier, C., Nguyen, T. V., Center, J. R., Seibel, M. J., and Eisman, J. A. (2005). Bone resorption and osteoporotic fractures in elderly men: the dubbo osteoporosis epidemiology study. J. Bone Miner. Res. 20, 579–587. 80. Bonde, M., Qvist, P., Fledelius, C., Riis, B. J., and Christiansen, C. (1994). Immunoassay for quantifying type I collagen degradation products in urine evaluated. Clin. Chem. 40, 2022–2025. 81. Fledelius, C., Kolding, I., Bonde, M., Cloos, P., and Christgau, S. (1996). Specificity of the Crosslaps ELISA and the MabA7 ELISA. Osteoporosis Int., 6(Suppl.1), PMO440. 82. Hanson, D. A., Weis, M. A., Bollen, A. M., Maslan, S. L., Singer, F. R., and Eyre, D. R. (1992). A specific immunoassay for monitoring human bone resorption: quantitation of type I collagen cross-linked N-telopeptides in urine. J. Bone Miner. Res. 7, 1251–1258. 83. Garnero, P., Gineyts, E., Arbault, P., Christiansen, C., and Delmas, P. D. (1995). Different effects of bisphosphonate and estrogen therapy on free and peptide-bound cross-links excretion. J. Bone Miner. Res. 10, 641–649. 84. Ju, H.-S.J., Leung, S., Brown, B., Stringer, M. A., Leigh, S., Scherrer, C., Shepard, K., Jenkins, D., Knudsen, J., and Cannon, R. (1997). Comparison of analytical performance and biological variability of three bone resorption assays. Clin. Chem. 43, 1570–1576. 85. Bonde, M., Fledelius, C., Qvist, P., and Christiansen, C. (1996). Coated tube radioimmunoassay for c-telopeptides of type I collagen to assess bone resorption. Clin. Chem. 42, 1639–1644. 86. Cloos, P. A., Lyubimova, N., Solberg, H., Qvist, P., Christiansen, C., Byrjalsen, I., and Christgau, S. (2004) An immunoassay for measuring fragments of newly synthesized collagen type I produced during metastatic invasion of bone. Clin. Lab. 50, 279–289. 87. Bonde, M., Garnero, P., Fledelius, C., Qvist, P., Delmas, P. D., and Christiansen, C. (1997). Measurement of bone degradation products in serum using antibodies reactive with an isomerized form of an 8 amino acid sequence of the c-telopeptide of type I collagen. J. Bone Miner. Res. 12, 1028–1034. 88. Rosenquist, C., Fledelius, C., Christgau, S., Pedersen, B. J., Bonde, M., Qvist, P., and Christiansen, C. (1998). Serum CrossLaps One Step ELISA. First application of monoclonal antibodies for measurement in serum of bone-related degradation products from C-terminal telopeptides of type I collagen. Clin. Chem. 44, 2281–2289. 89. Garnero, P., Borel, O., and Delmas, P. (2001). Evaluation of a fully automated serum assay for c-terminal cross-linking teleopeptide of type I collagen in osteoporosis. Clin. Chem. 47, 694–702. 90. Schmidt-gayk, H., Spanuth, E., Kotting, J., Bartl, R., Felsenberg, D., Pfeilschifter, J., Raue, F., and Roth, H. J. (2004). Performance evaluation of automated assays for beta-CrossLaps, N-MID-Osteocalcin and intact parathyroid hormone (BIOROSE Multicenter Study). Clin. Chem. Lab. Med. 42, 90–95.

560 91. Srivastava, A. K., MacFarlane, G., Srivastava, V. P., Mohan, S., and Baylink, D. J. (2001). A new monoclonal antibody ELISA for detection and characterization of C-telopeptide fragments of type l collagen in urine. Calcif. Tissue. Int. 69, 327–336. 92. Eyre, D. R. (1995). The specificity of collagen crosslinks as markers of bone and connective tissue degradation. Acta. Orthop. Scand. 66, 166–170. 93. Apone, S., Fevold, K., Lee, M., and Eyre, D. R. (1998). A rapid method for quantifying osteoclast activity in vitro. J. Bone Miner. Res. 9, S178(abstract). 94. Robins, S. P. (1995). Collagen cross-links in metabolic bone disease. Acta. Orhop. Scand. 66, 95. Gertz, B. J., Hanson, D. A., Quan, H., Harris, S. T., Genant, H. K., Chesnut III, CH. H., and Eyre, D. R. (1994). Monitoring bone resorption in early postmenopausal women by an immunoassay for crosslinked collagen peptides in urine. J. Bone Min. Res. 9, 135–142. 96. Clemens, J. D., Herrick, M. V., Singer, F. R., and Eyre, D. R. (1997). Evidence that serum NTx (collagen-type I N-telopeptides) can act as an immunochemical marker of bone resorption. Clin. Chem. 43, 2058–2063. 97. Bekker, P. J., Holloway, D. L., Rasmussen, A. S., Murphy, R., Martin, S. W., Leese, P. T., Holmes, G. B., Dunstan, C. R., and DePaoli, A. M. (2004). A single-dose placebo-controlled study of AMG 162, a fully human monoclonal antibody to RANKL, in postmenopausal women. J. Bone Miner. Res. 19, 1059–1066. 98. Ali, S. M., Demers, L. M., Leitzel, K., Harvey, H. A., Clemens, D., Mallinak, N., Engle, L., Chinchilli, V., Costa, L., Brady, C., Seaman, J., and Lipton, A. (2004). Baseline serum NTx levels are prognostic in metastatic breast cancer patients with bone-only metastasis. Ann. Oncol. 15, 455–459. 99. Guillemant, J. A., Accarie, C. M., de la Gueronniere, V., and Guillemant, S. E. (2003). Different acute responses of serum type I collagen telopeptides, CTX, NTX and ICTP, after repeated ingestion of calcium. Clin. Chim. Acta. 337, 35–41. 100. Scariano, J. K., Garry, P. J., Montoya, G. D., Duran-Valdez, E., and Baumgartner, R. N. (2002). Diagnostic efficacy of serum cross-linked N-telopeptide (NTx) and aminoterminal procollagen extension propeptide (PINP) measurements for identifying elderly women with decreased bone mineral density. Scand. J. Clin. Lab. Invest. 62, 237–243. 101. Greenspan, S. L., Rosen, H. N., and Parker, R. A. (2000). Early changes in serum N-telopeptide and C-telopeptide cross-linked collagen type I predict long-term response to alendronate therapy in elderly women. J. Clin. Endocrinol. Metab. 85, 3537–3540. 102. Fall, P. M., Kennedy, D., Smith, J. A., Seibel, M. J., and Raisz, L. G. (2000). Comparison of serum and urine assays for biochemical markers of bone resorption in postmenopausal women with and without hormone replacement therapy and in men. Osteoporos Int. 11, 481–485. 103. Chandani, A. K., Scariano, J. K., Glew, R. H., Clemens, J. D., Garry, P. J., and Baumgartner, R. N. (2000). Bone mineral density and serum levels of aminoterminal propeptides and cross-linked n-telopeptides of type 1 collagen in elderly men. Bone 26, 513–518. 104. Eastell, R., Mallinak, N., Weiss, S., Ettinger, M., Pettinger, M., Cain, D., Flessland, K., and Chesnutt lll, C. (2000). Biological variability of serum and urinary N-telopeptides of type l collagen in postmenopausal women. J. Bone Miner. Res. 15, 594–598. 105. Scariano, J. K., Glew, R. H., Bou-Serhal, C. E., Clemens, J. D., Garry, P. J., and Baumgartner, R. N. (1998). Serum levels of crosslinked N-telopeptides and aminoterminal propeptides of type I collagen indicate low bone mineral density in elderly women. Bone 23, 471–477. 106. Scariano, J. K., Vanderjagt, D. J., Thacher, T., Isichei, C. O., Hollis, B. W., and Glew, R. H. (1998). Calcium supplements increase the serum levels of crosslinked N-telopeptides of bone collagen and

Marius E. Kraenzlin and Markus J. Seibel

107.

108.

109.

110.

111.

112.

113.

114. 115.

116.

117.

118. 119.

120.

121.

122. 123.

124.

parathyroid hormone in rachitic Nigerian children. Clin. Biochem. 31, 421–427. Gertz, B. J., Clemens, J. D., Holland, S. D., Yuan, W., and Greenspan, S. (1998). Application of a new serum assay for type I collagen crosslinked N-telopeptides: assessment of diurnal changes in bone turnover with and without alendronate treatment. Calcif. Tissue Int. 63, 102–106. Hannon, R. A., Boreham, J. V., Porter, J. A., Karnik, K., Eagleton, A. C., Eastell, R., and Blumsohn, A. (2004) Technical and clinical evaluation of a point of care device for measurement of N-telopeptide fragment of type I collagen in urine. J. Bone Miner. Res. 19 (Suppl.1), SU147. Garnero, P., and Delmas, P. D. (2003). An immunoassay for type I collagen alpha1 helicoidal peptide 620-633, a new marker of bone resorption in osteoporosis. Bone 32, 20–26. Gerrits, M. I., Thijssen, J. H., and van Rijn, H. (1995). Determination of pyridinoline and deoxypyridinoline in urine, with special attention to retaining their stability. Clin. Chem. 41, 571–574. Beardsworth, L. J., Eyre, D. R., and Dickson, I. R. (1990). Changes with age in the urinary excretion of lysyl- and hydroxylysylpyridinoline, two new markers of bone collagen turnover. J. Bone Min. Res. 5, 671–676. Seibel, M. J., Gartenberg, F., Silverberg, S. J., Ratcliffe, A., Robins, S. P., and Bilezikian, J. P. (1992). Urinary hydroxypyridinium cross-links of collagen in primary hyperparathyroidism. J. Clin. Endocrinol. Metab. 74, 481–486. Colwell, A., Hamer, A., Blumsohn, A., and Eastell, R. (1996). To determine the effects of ultraviolet light, natural light and ionizing radiation on pyridinium cross-links in bone and urine using highperformance liquid chromatography. Europ. J. Clin. Invest. 26, 1107–1114. Walne, A. J., James, I. T., and Perret, D. (1995). The stability of pyridinium crosslinks in urine and serum. Clin. Chem. Acta. 240, 95–97. Blumsohn, A., Colwell, A., Naylor, K. E., and Eastell, R. (1995). Effect of light and gamma-irradiation on pyridinolines and telopeptides of type I collagen in urine. Clin. Chem. 41, 1195–1197. Eastell, R., Calvo, M. S., Burritt, M. F., Offord, K. P., Russell, R. G. G., and Riggs, B. L. (1992). Abnormalities in circadian patterns of bone resorption and renal calcium conservation in type I osteoporosis. J. Clin. Endocrinol. Metab. 74, 487–494. Schlemmer, A., Hassager, C., Jensen, S. B., and Christiansen, C. (1992). Marked diurnal variation in urinary excretion of pyridinium cross-links in premenopausal women. J. Clin. Endocrinol. Metab. 74, 476–480. Pagani, F., and Panteghini, M. (1994). Diurnal rhythm in urinary excretion of pyridinium crosslinks. Clin. Chem. 40, 952–953. Schlemmer, A., Hassager, C., Pedersen, B., and Christiansen, C. (1994). Posture, age, menopause, and osteopenia do not influence the circadian variation in the urinary excretion of pyridinium crosslinks. J. Bone Miner. Res. 9, 1883–1888. Bollen, A. M., Martin, M. D., Leroux, B. G., and Eyre, D. R. (1995). Circadian variation in urinary excretion on bone collagen cross-links. J. Bone Miner. Res. 10, 1885–1890. Eastell, R., Simmons, P. S., Assiri, A. M., Burritt, M. F., Russel, R. G. G., and Riggs, B. L. (1992). Nyctohemeral changes in bone turnover assessed by serum bone gla-protein concentration and urinary deoxypyridinoline excretion: effect of growth and aging. Clin. Sci. 83, 375–382. Schmeller, N. T., and Bauer, H. W. (1983). Circadian variation of different fractions of serum acid phosphatase. Prostate 4, 391–395. Li, Y., Lang, M., Kissling, C., Woitge, H. W., Farahmand, I., Oberwittler, H., Karmatschek, M., Armbuster, F. P., Ziegler, R., and Seibel, M. J. (1998). Biological variability in serum immunoreactive bone sialoprotein. Clin. Lab. 44, 553–555. Qvist, P., Christgau, S., Pedersen, B. J., Schlemmer, A., and Christiansen, C. (2002). Circadian variation in the serum concentration

Chapter 33 Measurement of Biochemical Markers of Bone Resorption

125.

126.

127.

128.

129.

130. 131.

132.

133.

134.

135.

136.

137.

138.

139.

140.

141.

of C-terminal telopeptide of type I collagen (serum CTx): effects of gender, age, menopausal status, posture, daylight, serum cortisol, and fasting. Bone 31, 57–61. Hannon, R., Clowes, J., Eagleton, A. C., Al Hadari, A., Eastell, R., and Blumsohn, A. (2004). Clinical performance of immunoreactive tartrate resistent acid phosphatase isoform 5b as a marker of bone resorption. Bone 34, 187–194. Wichers, M., Schmidt, E., Bidlingmaier, M., and Klingmüller, D. (1999). Diurnal rhythm of CrossLaps in human serum. Clin. Chem. 45, 1858–1860. Caillot-Augusseau, A., Vico, L., Heer, M., Voroviev, D., Souberbielle, J. C., Zitterman, A., Alexandre, C., and LafageProust, M. H. (2000). Space flight is associated with rapid decreases of undercarboxylated osteocalcin and increases of markers of bone resorption without changes in their circadian variation: observations in two cosmonauts. Clin. Chem. 46, 1136–1143. Bjarnason, N. H., Henriksen, E. E., Alexandersen, P., Christgau, S., Henriksen, D. B., and Christiansen, C. (2002). Mechanism of circadian variation in bone resorption. Bone 30, 307–313. Nielsen, H. K., Brixen, K., and Mosekilde, L. (1990). Diurnal rhythm and 24-hour integrated concentrations of serum osteocalcin in normals: Influence of age, sex, season, and smoking habits. Calcif. Tissue Int. 47, 284–290. Christgau, S. (2000). Circadian variation in serum CrossLaps concentration is reduced in fasting individuals. Clin. Chem. 46, 431. Markowitz, M. E., Arnaud, S., Rosen, J. F., Thorpy, M., and Laximinarayan, S. (1988). Temporal interrelationships between the circadian rhythms of serum parathyroid hormone and calcium concentrations. J. Clin. Endocrinol. Metab. 67, 1068–1073. Calvo, M. S., Eastell, R., Offord, K. P., Bergstralh, E. J., and Burritt, M. F. (1991). Circadian variation in ionized calcium and intact parathyroid hormone: Evidence for sex differences in calcium homeostasis. J. Clin. Endocrinol. Metab. 72, 69–76. Nielsen, H. K., Laurberg, P., Brixen, K., and Mosekilde, L. (1991). Relations between diurnal variations in serum osteocalcin, cortisol, parathyroid hormone, and ionized calcium in normal individuals. Acta. Endocrinol. (Copenh) 124, 391–398. Ebeling, P. R., Butler, P. C., Eastell, R., Rizza, R. A., and Riggs, B. L. (1991). The nocturnal increase in growth hormone is not the cause of the nocturnal increase in serum osteocalcin. J. Clin. Endocrinol. Metab. 73, 368–372. Henriksen, D. B., Alexandersen, P., Bjarnason, N. H., Vilsboll, T., Hartmann, B., Henriksen, E. E., Byrjalsen, I., Krarup, T., Holst, J. J., and Christiansen, C. (2003). Role of gastrointestinal hormones in postprandial reduction of bone resorption. J. Bone Miner. Res. 18, 2180–2189. Henriksen, D. B., Alexandersen, P., Byrjalsen, I., Hartmann, B., Bone, H. G., Christiansen, C., and Holst, J. J. (2004). Reduction of nocturnal rise in bone resorption by subcutaneous GLP-2. Bone 34, 140–147. Heymsfield, S. B., Artega, C., McManus, B. S., Smith, J., and Moffitt, S. (1983). Measurements of muscle mass in humans: validity of the 24 hour urinary creatinine method. Am. J. Clin. Nutr. 37, 478–494. Popp-Snijders, C., Lips, P., and Netelenbos, J. C. (1996). Intraindividual variation in bone resorption markers in urine. Ann. Clin. Biochem. 33, 347–348. Blumsohn, A., Hannon, R. A., al-Dehaimi, A. W., and Eastell, R. (1994). Short-term intraindividual variability of markers of bone turnover in healthy adults. J. Bone Miner. Res. 9(Suppl.1), S153. Panteghini, M., and Pagani, F. (1995). Biological variation in bonederived biochemical markers in serum. Scand. J. Clin. Lab. Invest. 55, 609–616. Garnero, P., Mulleman, D., Munoz, F., Sornay-Rendu, E., and Delmas, P. D. (2003). Long-term variability of markers of bone

142.

143.

144.

145.

146.

147.

148.

149.

150.

151.

152.

153. 154.

155.

156.

157.

158.

159.

561

turnover in postmenopausal women and implications for their clinical use: the OFELY study. J. Bone Miner. Res. 18, 1789–1794. Christgau, S., Bitsch-Jensen, O., Bjarnason, N., Henriksen, E. G., Qvist, P., Alexandersen, P., and Henriksen, D. B. (2000). Serum CrossLaps for monitoring the response in individuals undergoing antiresorptive therapy. Bone 26, 505–511. Robins, S. P. (1990). Collagen markers in urine in human arthritis. In: Methods in Cartilage Research (A. Maroudas, et al, ed.), pp. 348–352. Academic Press, London. Fisher, L. W., McBride, O. W., Termine, J. D., and Young, M. F. (1990). Human bone sialoprotein: deduced protein sequence and chromosomal localization. J. Biol.Chem. 265, 2347–2351. Oldberg, A., Franzen, A., and Heinegard, D. (1985). Isolation and characterization of two sialoproteins present only in bone. Biochem. J. 232, 715–724. Chen, J., Shapiro, H. S., and Wrana, J. L. (1991). Localization of bone sialoprotein (BSP) expression to sites of mineral tissue formation in fetal rat tissue by in-situ hybridization. Matrix 11, 133–143. Shapiro, H. S., Chen, J., Wrana, J. L., Zhang, Q., Blum, M., and Sodek, J. (1993). Characterization of porcine bone sialoprotein: primary structure and cellular expression. Matrix 13, 431–440. Fujisawa, R., Butler, W. T., Brunn, J. C., Zhou, H. Y., and Kuboki, Y. (1993). Differences in composition of cell-attachment sialoproteins between dentin and bone. J. Dent. Res. 72, 1222–1226. Ross, F. P., Chappel, J., Alvarez, J. I., Sander, D., Butler, W. T., Farach-Carson, M. C., Mintz, K. A., Robey, P. G., Teitelbaum, S. L., and Cheresh, D. A. (1993). Interactions between the bone matrix proteins osteopontin and bone sialoprotein and the osteoclast integrin alpha v beta 3 potentiate bone resorption. J. Biol. Chem. 268, 9901–9907. Fujisawa, R., Nodasaka, Y., and Kuboki, Y. (1995). Further characterization of interaction between bone sialoprotein (BSP) and collagen. Calcif. Tissue Int. 56, 140–144. Hunter, G. K., Kyle, C. L., and Goldberg, H. A. (1994). Modulation of crystal formation by bone phosphoproteins: structural specificity of the osteopontin-mediated inhibition of hydroxyapatite formation. Biochem. J. 300, 723–728. Young, M. F., Kerr, J. M., Ibaraki, K., Heegaard, A. M., and Robey, P. G. (1992). Structure, expression, and regulation of the major noncollagenous matrix proteins of bone. Clin. Orthop. 281, 275–294. Chenu, C. and Delmas, P. D. (1992). Platelets contrubute to circulating levels of bone sialoprotein in human. J. Bone Miner. Res. 7, 47–54. Zhang, J. H., Tang, J., Wang, J., Ma, W., Zheng, W., Yoneda, T., and Chen, J. (2003). Over-expression of bone sialoprotein enhances bone metastasis of human breast cancer cells in a mouse model. Int. J. Oncol. 23, 1043–1048. Sharp, J. A., Waltham, M., Williams, E. D., Henderson, M. A., and Thompson, E. W. (2004). Transfection of MDA-MB-231 human breast carcinoma cells with bone sialoprotein (BSP) stimulates migration and invasion in vitro and growth of primary and secondary tumors in nude mice. Clin. Exp. Metastasis. 21, 19–29. Adwan, H., Bauerle, T., Najajreh, Y., Elazer, V., Golomb, G., and Berger, M. R. (2004). Decreased levels of osteopontin and bone sialoprotein II are correlated with reduced proliferation, colony formation, and migration of GFP-MDA-MB-231 cells. Int. J. Oncol. 24, 1235–1244. Saxne, T., Zunino, L., and Heinegard, D. (1995). Increased release of bone sialoprotein into synovial fluid reflects tissue destruction in rheumatoid arthritis. Arthritis Rheum. 38, 82–90. Ohno, U., Matsuyama, T., Ishii, S., Hata, H., and Kuboki, Y. (1995). Measurement of human immunoreactive bone sialoprotein (BSP) in normal adults and in pregnant women. J. Bone Miner. Res. 10 (Suppl 1): S476(Abstract). Karmatschek, M., Maier, I., Seibel, M. J., Woitge, H. W., Ziegler, R., and Armbruster, F. P. (1997). Improved purification of human bone

562

160.

161.

162.

163.

164.

165.

166.

167.

168.

169. 170.

171.

172. 173. 174. 175.

176. 177.

Marius E. Kraenzlin and Markus J. Seibel sialoprotein and development of a homologous radioimmunoassay. Clin. Chem. 43, 2076–2082. Cogan, G., Bansal, A. K., Ibrahim, S., Zhu, B., Goldberg, H. A., Ganss, B., Cheifetz, S., Armbruster, F. P., and Sodek, J. (2004). Analysis of human bone sialoprotein in normal and pathological tissues using a monoclonal antibody (BSP 1.2 mab). Connect. Tissue Res. 45, 60–71. Seibel, M. J., Woitge, H. W., Pecherstorfer, M., Karmatschek, M., Horn, E., Ludwig, H., Armbruster, F. P., and Ziegler, R. (1996). Serum immunoreactive bone sialoprotein as a new marker of bone turnover in metabolic and malignant bone disease. J. Clin. Endocrinol. Metab. 81, 3289–3294. Woitge, H. W., Pecherstorfer, M., Horn, E., Keck, A. V., Diel, I. J., Bayer, P., Ludwig, H., Ziegler, R., and Seibel, M. J. (2001). Serum bone sialoprotein as a marker of tumour burden and neoplastic bone involvement and as a prognostic factor in multiple myeloma. Br. J. Cancer 84, 344–351. Jung, K., Lein, M., Stephan, C., Von, H. K., Semjonow, A., Sinha, P., Loening, S. A., and Schnorr, D. (2004). Comparison of 10 serum bone turnover markers in prostate carcinoma patients with bone metastatic spread: diagnostic and prognostic implications. Int. J. Cancer 111, 783–791. Woitge, H. W., Knothe, A., Witte, K., Schmidt-gayk, H., Ziegler, R., Lemmer, B., and Seibel, M. J. (2000). Circannual rhythms and interactions of vitamin D metabolites, parathyroid hormone, and biochemical markers of skeletal homeostasis: A prospective study. J. Bone Miner. Res. 15, 2443–2450. Faust, D., Menge, F., Armbruster, F. P., Lembcke, B., and Stein, J. (2003). Increased serum bone sialoprotein concentrations in patients with Crohn’s disease. Z. Gastroenterol. 41, 243–247. Taylor, A. K., Linkhart, S., Mohan, S., Christenson, R. A., Singer, F. R., and Baylink, D. J. (1990). Multiple osteocalcin fragments in human urine and serum as detected by a midmolecule osteocalcin radioimmunoassay. J. Clin. Endocrinol. Metab. 70, 467–472. Ivaska, K. K., Hellman, J., Likojarvi, J., Kakonen, S. M., Gerdhem, P., Akesson, K., Obrant, K. J., Pettersson, K., and Vaananen, H. K. (2003). Identification of novel proteolytic forms of osteocalcin in human urine. Biochem. Biophys. Res. Commun. 306, 973–980. Ivaska, K. K., Hentunen, T. A., Vaaraniemi, J., Ylipahkala, H., Pettersson, K., and Vaananen, H. K. (2004). Release of intact and fragmented osteocalcin molecules from bone matrix during bone resorption in vitro. J. Biol. Chem. 279, 18361–18369. Cloos, P. A. and Christgau, S. (2004). Characterization of aged osteocalcin fragments derived from bone resorption. Clin. Lab. 50, 585–598. Gerdhem, P., Ivaska, K. K., Alatalo, S. L., Halleen, J. M., Hellman, J., Isaksson, A., Pettersson, K., Vaananen, H. K., Akesson, K., and Obrant, K. J. (2004). Biochemical markers of bone metabolism and prediction of fracture in elderly women. J. Bone Miner. Res. 19, 386–393. Ivaska, K. K., Kakonen, S. M., Gerdhem, P., Obrant, K. J., Pettersson, K., and Vaananen, H. K. (2005). Urinary osteocalcin as a marker of bone metabolism. Clin. Chem. 51, 618–628. Romas, N. A., Rose, N. R., and Tannenbaum, M. (1979). Acid phosphatase: New developments. Hum. Pathol. 10, 501–512. Yam, L. T. (1974). Clinical significance of human acid phosphatases. Am. J. Med. 56, 604–616. Li, C. Y., Chuda, R. A., Lam, K.-W., and Yam, L. T. (1973). Acid phosphatases in human plasma. J. Lab. Clin. Med. 82, 446–460. Lam, K. W, Lai, L., and Yam, L. T. (1978). Tartrate-resistant acid phosphatase activity measured by electrophoresis an acrylamid gel. Clin. Chem. 24, 309–312. Vaes, G., and Jacques, P. (1965) The assay of acid hydrolases and other enzymes in bone tissue. Biochem. J. 97, 380–388. Wergedal, J. E. (1970). Characterization of bone acid phosphatase activity. Proceeding of the Society for Experimental Biology, 134, 244–247.

178. Hammarstrom, L. E., Hanker, J. S., and Toverud, S. U. (1971). Cellular differences in acid phosphatase isoenzymes in bone and teeth. Clin. Orthop. 78, 151–155. 179. Lau, K.-H. W., Stepan, J. J., Yoo, A., Mohan, S., and Baylink, D. J. (1991). Evidence that tartrate-resistant acid phosphatases from osteoclastomas and hairy cell leukemia spleen are members of a multigene family. Int. J. Biochem. 23, 1237–1244. 180. Wergedal, J., Lau, K.-H. W., and Baylink, D. J. (1988). Fluoride and bovine bone extract influence cell proliferation and phosphatase activities in human bone cell cultures. Clin. Orthop. Rel. Res. 233, 274–282. 181. Bianco, P., Ballanti, P., and Bonucci, E. (1988). Tartrate resistant acid phosphatase activity in rat osteoblasts and osteocytes. (UnPub) 182. Minkin, C. (1982). Bone acid phosphatase: tartrate-resistant acid phosphatase as a marker of osteoclast function. Calc. Tiss. Int. 34, 285–290. 183. Wergedal, J. E. and Baylink, D. J. (1984). Characterization of cells isolated and cultured from human bone. Proceeding of the Society for Experimental Biology, 176, 27–31. 184. Lam, K. W., Lee, P., Li, C. Y., and Yam, L. T. (1980). Immunological and biochemical evidence for identity of tartrate-resistent isoenzymes of acid phosphatases from human spleen and tissues. Clin. Chem. 26, 420–422. 185. Lau, K. H. W., Onishi, T., Wergedal, J. E., Singer, F. R., and Baylink, D. J. (1987). Characterization and assay of tartrate-resistent acid phosphatase activity in serum: potential use to assess bone resorption. Clin. Chem. 33, 458–462. 186. Stepan, J. J., Pospichal, J., Presl, J., and Pacovsky, V. (1987). Bone loss and biochemical indices of bone remodelling in surgically induced postmenopausal women. Bone 8, 279–284. 187. Stepan, J. J., Tesarova, A., Havranek, T., Jodl, J., Normankova, J., and Pacovsky, V. (1985). Age and sex dependency of the biochemical indices of bone remodelling. Clin. Chem. Acta. 151, 273–283. 188. Stepan, J. J., Silinkova-Malkova, E., Havranek, T., Formankova, J., Zichova, M., Lachmanova, J., Strakova, M., Broulik, P., and Pacovsky, V. (1983). Relationship of plasma tartrate resistant acid phosphatase to the bone isoenzyme of serum alkaline phosphatase in hyperparathyroidism. Clin. Chem. Acta. 133, 189–200. 189. Cooper, J. D. H., Turnell, D. C., and Price, C. P. (1982). The estimation of serum acid phosphatase using alpha-naphthyl phosphate as substrate: observation on the use of fast red TR salt in the assay. Clin. Chem. 126, 297–306. 190. Bais, R. and Edwards, J. B. (1976). An optimized continuous-monotoring procedure for semiautomated determination of serum acid phosphatase activity. Clin. Chem. 22, 2025–2028. 191. Lam, K. W., Burke, D. S., Siemens, M., Cipperly, V., Li, C. Y., and Yam, L. T. (1982). Characterization of serum acid phosphatase associated with dengue hemorrhagic fever. Clin. Chem. 28, 2296–2299. 192. Warren, R. J. and Moss, D. W. (1977). An automated continuousmonitoring procedure for the determination of acid phosphatase activity in serum. Chim. Acta. 77, 179–188. 193. Omene, J. A., Glew, R. H., Baig, H. A., Robinson, D. B., Brock, W., Chamber, J. P. (1981). Determination of serum acid and alkaline phosphatase using 4-methylumbelliferyl phosphate. Afr. J. Med. Sci. 10, 9–18. 194. Nakanishi, M., Yoh, K., Uchida, K., Maruo, S., and Matsuoka, A. (1998). Improved method for measuring tartrate-resistant acid phosphatase activity in serum. Clin. Chem. 44, 221–225. 195. Lam, K. W., Siemens, M., Sun, T., Li, C.Y., Yam, L.T. (1982). Enzyme-immunoassay for tartrate-resistent acid phosphatase. Clin. Chem. 28, 467–470. 196. Echetebu, Z. O., Cox, T. M., and Moss, D. W. (1987). Antibodies to porcine uteroferrin used in measurement of human tartrate-resistant acid phosphatase. Clin. Chem. 33 No10, 1832–1836. 197. Whitaker, K. B., Cox, T. M., and Moss, D. W. (1989). An immunoassay of human band-5 (“tartrate-resistent”) acid phosphatase that

Chapter 33 Measurement of Biochemical Markers of Bone Resorption

198.

199.

200. 201.

202.

203.

204.

205.

involves the use of anti-porcine uteroferrin antibodies. Clin. Chem. 35, 86–89. Cheung, C. K., Panesar, N. S., Haines, C., Masarei, J., and Swaminathan, R. (1995). Immunoassay of a tartrate-resistant acid phosphatase in serum. Clin. Chem. 41, 679–686. Kraenzlin, M. E., Lau, K.-H. W., Liang, L., Freeman, T. K., Singer, F. R., Stepan, J., and Baylink, D. J. (1990). Development of an immunoassay for human serum osteoclastic tartrate-resistant acid phosphatase. J. Clin. Endocrinol. Metab. 71, 442–451. Ott, S. M. (1993). Clinical effects of bisphosphonates in involutional osteoporosis. J. Bone Miner. Res. 8(Suppl. 2), S597–S606. Halleen, J., Hentunen, T. A., Karp, M., Käkönen, S. M., Petterson, K., and Väänänen, H. K. (1998). Characterization of serum tartrate-resistant acid phosphatase and development of a direct two-site immunoassay. J. Bone Miner. Res. 13, 683–687. Pasco, J. A., Henry, M. J., Kotowicz, M. A., Sanders, K. M., Seeman, E., Pasco, J. R., Schneider, H. G., and Nicholson, G. C. (2004). Seasonal periodicity of serum vitamin D and parathyroid hormone, bone resorption, and fractures: the Geelong Osteoporosis Study. J. Bone Miner. Res. 19, 752–758. Halleen, J., Hentunen, T. A., Hellman, J., and Vaananen, H. K. (1996). Tartrate-resistant acid phosphatase from human bone: purification and development of an immunoassay. J. Bone Miner. Res. 11, 1444–1452. Halleen, J., Alatalo, S., Suominen, H., Cheng, S., Janckila, A. J., and Väänänen, H. K. (2000). Tartrate-resistent acid phosphatase 5b: a novel serum marker of bone resorption. J. Bone Miner. Res. 15, 1337–1345. Chu, P., Chao, T. Y., Lin, Y. F., Janckila, A. J., and Yam, L. T. (2003). Correlation between histomorphometric parameters of bone resorption and serum type 5b tartrate-resistant acid phosphatase in uremic patients on maintenance hemodialysis. Am. J. Kidney Dis. 41, 1052–1059.

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206. Seibel, M. J. (2000). Molecular marker on bone turnover: biochemical, technical and analytical aspects. Osteoporos Int, 11 (Suppl. 6), 18–29. 207. Stepan, J. J., Lachmanova, J., Strakova, M., and Pacovsky, V. (1987). Serum osteocalcin, bone alkaline phosphatase isoenzyme and plasma tartrate resistent acid phosphatase in patients on chronic maintenance hemodialysis. Bone Min. 3, 177–183. 208. Gelb, B. D., Shi, G. P., Chapman, H. A., and Desnick, R. J. (1996). Pycnodysostosis, a lysosomal disease caused by cathepsin K deficiency. Science 273, 1236–1238. 209. Saftig, P., Hunziker, E., Wehmeyer, O., Jones, S., Boyde, A., Rommerokirch, W., Moritz, J. D., Schu, P., and von Figura, K. (1998). Impaired osteoclastic bone resorption leads to osteopetrosis in cathepsinK-deficient mice. Proc. Natl Acad. Sci. USA 95, 13453–13458. 210. Gowen, M., Lazner, F., Dodds, R., Kapadia, R., Feild, J., Tavaria, M., Bertoncello, I., Drake, F., Zavarselk, S., Tellis, I., Hertzog, P., Debouck, C., and Kola, I. (1999). Cathepsin K knockout mice develop osteopetrosis due to a deficit in matrix degradation but not demineralization. J. Bone Miner. Res. 14, 1654–1663. 211. Goto, T., Yamaza, T., and Tanaka, T. (2003). Cathepsins in the osteoclast. J. Electron. Microsc. (Tokyo), 52, 551–558. 212. Meier, C., Meinhardt, U., Greenfield, J. R., Nguyen, T.V., de Winter, J., Dunstan, C., and Seibel, M. J. (2006). Serum cathepsin K levels reflect osteoclastic activity in postmenoopausal osteoporosis and Paget’s disease of bone. Clin. Lab. (in press). 213. Lang, T. H., Willinger, U., and Holzer, G. (2004). Soluble cathepsin-L: a marker of bone resorption and bone density? J. Lab. Clin. Med. 144, 163–166. 214. Robins, S. P. (1997). Measurement of pyridinium crosslinks by HPLC and immunoassay. In: Calcium Regulating Hormones and Markers of Bone Metabolism: Measurement and Interpretation (H. Schmidt-gayk, et al, ed.), pp. 135–140. Clin Lab Publications, Heidelberg, Germany.

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Chapter 34

Variability in the Measurement of Biochemical Markers of Bone Turnover Tuan V. Nguyen

Bone and Mineral Research Program, Garvan Institute of Medical Research, St. Vincent’s Hospital and University of NSW Research Program

Christian Meier

Endokrinologische Praxis & Labor, University Hospital Basel, Switzerland

Markus J. Seibel

Bone Research Program, ANZAC Research Institute, The University of Sydney and Concord Hospital, Sydney, Australia

I. Introduction II. Sources of Pre-Analytical Variability in the Measurement of Biochemical Markers of Bone Turnover

III. Statistical Consideration of Variability IV. Summary References

I. INTRODUCTION

However, in reality, measurements of markers of turnover are subject to considerable nonspecific within-subject variability, and this problem presents a major issue in the management of osteoporosis in clinical setting. When a change is observed in an individual patient after an invention, the change must be interpreted against the background of the respective marker’s total variability. It is, therefore, important to examine the sources of variability and analytic techniques to help the interpretation of change. Because measures of bone turnover markers are often used in epidemiologic studies as risk factors (or measures of exposure), the within-subject variability also presents problems in the interpretation of results from such studies. The reason is that as markers of bone resorption and bone

Variability in the measurement of chemical analytes is a major issue in clinical chemistry. Consideration and, wherever possible, tight control of factors not related to the specific process in question, is essential for the correct interpretation of laboratory results. At present, these caveats seem of particular relevance for the various molecular markers of bone turnover, as the degree of variability of some of these analytes has been shown to be rather substantial. Ideally, biochemical markers of turnover, and the assays used for their quantification, should have minimal variability, so that its change can be interpreted with confidence. Dynamics of Bone and Cartilage Metabolism

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formation fluctuate over time, a single measurement may not be very representative of a subject’s “true” underlying value, and can complicate the estimation of an etiologic association. It has been well established in epidemiology that when an unreliable measurement for a continuous risk factor is analyzed for an association with an outcome, then the association can be over- or underestimated [1]. More specifically, in the case of most osteoporosis studies, measures of bone turnover markers can dilute or attenuate the observed association between bone resorption/formation and the outcome of interest, thereby reducing the power (and the replicability) of the study to detect an etiologic association. The ability to quantify the reliability of an exposure measurement is thus an important consideration in the design and statistical analysis of epidemiologic studies. Conceptually and functionally, sources of variability can be classified into two broad groups: between-subject differences and within-subject variation. The betweensubject differences are attributable to differences in genetic make-up, age, gender, and clinico-demographic factors. The within-subject variation can be further classified into two sources: technical variability and biologic variability (Table I). Among the pre-analytical factors affecting biochemical measurements are firstly technical issues such as dietary influences, the type of specimen collected, and the mode of sample processing and storage. Biological variability includes effects such as diurnal or day-to-day

Table I.

Sources and Categories of Pre-analytical Variability

– Technical (sample and assay related) – Type of specimen (serum vs. urine) – Type of urine sample (24-h vs. FMV vs. SMV) – Sample processing – Sample storage (analyte stability) – Diet – Biological (subject related) – Intraindividual – Diurnal variability – Day-to-day variability – Menstrual rhythms – Seasonal rhythms – Exercise – Interindividual – Age – Gender – Race – Pregnancy and lactation – Menopausal status – Growth – Nonskeletal diseases – Skeletal disease (including fractures) – Immobility – Drugs (including oral contraceptives) – Clearance-related factors

variation, seasonal rhythms, and menstrual rhythms. However, the deciphering of the two sources of variability is not an easy matter, because of the interaction between genetic and environmental factors in the determination of bone turnover markers, and as a result, certain betweensubject differences can also be attributed to the withinsubject biologic variability. This chapter is mainly concerned with issues of withinsubject variability. It reviews the sources of variability and then presents a number of statistical models to estimate the variability and characterize the change in bone turnover markers in an individual patient.

II. SOURCES OF PRE-ANALYTICAL VARIABILITY IN THE MEASUREMENT OF BIOCHEMICAL MARKERS OF BONE TURNOVER A. Technical Aspects of Pre-analytical Variability The choice of sample (i.e. serum versus urine), the mode of urine collection (i.e. 24-hour collection versus first or second morning void), the appropriate preparation of the patient (i.e. minimizing the effects of diet or exercise before phlebotomy) and the correct processing and storage of specimens all influence the final analytical result. Therefore, special care must be taken of these more technical issues, in particular as they are modifiable and controllable [2]. 1. Handling, Processing and Storage of Samples

Some, but not all biochemical components used as markers of bone turnover are sensitive to ambient conditions such as temperature or UV radiation. In components susceptible to these influences, thermodegradation or photolysis will lead to a reduction in the measured signal and, if not noted, to a misinterpretation of the laboratory result. While the alkaline phosphatases and the procollagen propeptides have been shown to be rather stable at room temperature, serum osteocalcin is readily degraded even at temperatures as low as 4⬚C [3–5]. When using assays for intact OC(1-49), rapid enzymatic cleavage of the peptide into smaller fragments will lead to significant signal losses if the serum sample is kept at room temperature for more than 1–2 h. Adding protease inhibitors will delay, but by no means prevent this process [4]. Signal reduction is less pronounced when measuring smaller (already degraded) fragments of OC, e.g. OC(1-43) [4] (Fig. 1A). In contrast, both the free and conjugated forms of PYD and DPD have been shown to be stable in urine samples

Chapter 34 Variability in the Measurement of Biochemical Markers of Bone Turnover

A

567

B

Figure 1 (A) Loss of OC immunoreactivity by storage of regular serum samples at room temperature (). Addition of a stabilizer () partially prevents epitope loss over 12 hours of storage. Results are mean and SD. From reference [4]. (B) Effects of storage temperature on serum TRAP activity. Five serum samples were stored at either –20°C () or –70°C (
) for the indicated times, then assayed. The slope of each line, calculated by unweighted least squares linear regression, was the basis of the determination of the relative enzyme stability. From reference [13].

kept at room temperature for several weeks. Several reports show that pyridinium crosslinks can be stored at –20⬚C for years and that repeated freeze–thaw cycles of urine samples have no effect on the concentrations of PYD and DPD [6–9]. Similar stability has been reported for the urinary N-terminal (NTx) and C-terminal (CTx) collagen type I telopeptides, while C-terminal telopeptide in serum (ICTP) loses up to 12% of the signal when stored at room temperature for 5 days [10–12]. The activity of serum tartrate-resistant acid phosphatase (TRAP) declines rapidly during storage at room temperature or even at −20⬚C but is stable when stored at −70⬚C or lower [13] (Fig. 1B). Multiple freezing–thaw cycles usually have a deleterious effect on the serum TRAP activity [14]. In contrast, serum levels of bone sialoprotein (BSP) appear rather stable, both at room temperature, 4⬚C and −20⬚C, and have been shown to not change significantly during repeated freeze– thaw cycles [5]. However, when samples are being exposed to temperatures above 30⬚C, an increase in signal is usually seen with the RIA [5]. This effect may be due to the generation of neo-epitopes during the degradation of BSP–protein complexes. Some assays and marker components are sensitive to hemolysis of the sample, giving results that are either too low or too high. This is usually the case for osteocalcin and bone sialoprotein, but has also been described for TRAP and some other serum markers. Pyridinium crosslinks in aqueous solutions are unstable when subjected to intensive UV irradiation [15–17].

The increased rate of photolysis in the aqueous standards, compared with pyridinoline in urinary samples, may be attributed to the fact that the intensity of UV light is higher in the aqueous solutions. Photolysis of the pyridinoline standard solution is prevented by storage in amber vials [16]. The rate of pyridinoline photolysis increases with rising pH [15]. It has also been shown that the sensitivity to UV light is greater for free pyridinoline than for total pyridinoline [17]. The collagen type I telopeptides in urine, NTx and CTx, are not affected by UV light exposure [17]. 2. Timing and Mode of Urine Collection

In general, random samples can be used for measurement of most urinary parameters (except urinary calcium, which always requires a 24-hour sample). For convenience, measurement of bone turnover markers is usually performed either in first or second morning voids, or in 2-hour collections. In each case, values need to be corrected for urinary creatinine, which introduces additional pre-analytical and analytical variability. This necessity, i.e. the introduction of a second analyte, is considered one of the major reasons why urine markers of bone turnover show a higher degree of variability than serum indices. Creatinine output has been reported to be fairly constant with time (variations within 10%), and to correlate with lean body mass [18], but there are also reports suggesting that the correction for creatinine in a urine spot sample could be misleading. As an alternative, a 24-hour urine collection can be ordered and the

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excretion rate of the marker determined [19]. However, 24-hour urine collections are subject to inevitable inaccuracies due to collection errors. Experienced clinical chemists consider the collection of a truly timed 24-hour urine sample one of the more demanding tasks in laboratory medicine. With most markers, similar results are obtained from either 24-hour, 2-hour, or spot urine (FMV, SMV) collections. It should be borne in mind, however, that the slope of diurnal changes is steepest during the morning hours (see below), which is usually the time at which urine samples are collected. Therefore, if spot urine samples are used for measurement of bone turnover markers, the time of collection should be controlled as closely as possible. 3. Influence of Diet or Acute Exercise

The only bone turnover marker that is markedly affected by dietary influences is urinary hydroxyproline, a degradation product of newly synthesized and mature collagens (see Chapter 35). Urinary levels of hydroxyproline will rise considerably after the ingestion of hydroxyproline-rich foods, such as meat or gelatine [20]. Therefore, it is necessary to instruct patients to keep a collagen-free diet for at least 24 h before collecting their urine for hydroxyproline measurements. While the fasting or nonfasting state has no significant influence on the levels of most serum and urine markers, recent results suggest that serum CTx may be influenced by a previous meal [21] (Fig. 2). Thus, although no general recommendation has been put forward so far, samples for bone turnover markers should be taken in the fasting state. The effect of acute exercise immediately or shortly before phlebotomy for bone turnover markers has been studied, but results are equivocal [22–24]. While some markers appear to rise by as much as 30–40% of their baseline value,

others seem to be unaffected by these activities. These short-term effects need to be distinguished from the welldocumented influences of long-term endurance exercise on bone turnover [25]. 4. Standardization

Although standardization is part of the quality control process and therefore a concern of analytical rather then pre-analytical variability, the striking lack in standardization of most bone marker assays and its impact on the practical utility of these parameters deserves a few words. Markers of bone turnover are now offered by a great number of commercial laboratories and are widely used among practicing physicians in many countries. Nevertheless, there are presently no routine proficiency testing programs for these parameters. A recent trial amongst laboratories in Europe showed marked variability of most commercialized test kits, with interlaboratory coefficients of variation up to 48%. Results obtained from identical blood and urine samples using the same assay and the same method differed up to 7.3-fold between laboratories. If ordered in a clinical context, the same sample would have returned with normal and highly abnormal results [26] (Table II). Although several companies have promised stricter QA programs, the authors have not seen any major progress in this area. Therefore, results from one laboratory cannot be compared to those of a different laboratory, even if the same method and the same reference range has been used. It is obvious that immunoassays for bone turnover markers need to undergo strict standardization and should be included into routine proficiency testing programs.

B. Biological Aspects of Pre-analytical Variability 1. Diurnal Variation

Figure 2 Serum CTx in premenopausal women on a normal diet and during fasting. The cyclical variation was statistically significant both on normal diet and during fasting. From reference [21].

Most biochemical markers show significant diurnal variations, with highest values in the early morning hours and lowest values during the afternoon and at night. More than 30 years ago, Mautalen demonstrated that the excretion of urinary hydroxyproline (OHP) follows a circadian rhythm and this rhythm is fairly constant from day to day [27]. Diurnal variation in pre- and postmenopausal women, as well as men, has now been reported by many investigators for all biochemical markers of bone turnover, including the pyridinium and the telopeptide markers, as well as for tartrate-resistant acid phosphatase and bone sialoprotein [2, 5, 10, 21, 28–43]. Serum markers usually show less pronounced changes during the day than urinebased indices. This seems also true for the new serum

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Chapter 34 Variability in the Measurement of Biochemical Markers of Bone Turnover Table II.

Marker

Minimum and Maximum Values and CVILs Reported for Identical Bone Markers and Methods (from: Seibel et al. 2001 [26]) Assay

Units

BAP

IRMA (n = 22)

µg/L

BAP

EIA (n = 16)

U/L

BAP

LP (n = 6)

U/L

OC

LIA (n = 8)

µg/L

OC

EIA (n = 5)

µg/L

OC

all RIAs (n = 18)

µg/L

OC, RIA subgroups

1 (n = 6)

µg/L

2 (n = 6)

µg/L

Total DPD

HPLC (n = 29)

nmol

Free DPD

EIA (n = 15)

nmol

Free DPD

CLIA (n = 12)

nmol

NTx

EIA (n = 10)

nmol

ICTP

RIA (n = 5)

µg/L

Hydroxyproline

HPLC (n = 7)

µmol/L

Sample pool A B A B A B A B A B A B A B A B A B A B A B A B A B A B

Max

MMR

Group mean for sampleb

4.8 21.0 13.0 27.5 15 79 1.8 10.2 2.3 14.9 0.1 2.6

15.8 39.9 24.0 63.6 111 149 3.4 16.0 5.8 24.6 13.0 79.6

148 49 134 61 147 69 1410 410 11.1 7.5 645 186

710 197 190 73 177 100 8056 1564 14 9.9 936 232

3.3 1.9 1.8 2.3 7.3 1.9 1.9 1.6 2.5 1.7 130c 30c 2.5 1.8 2.7 3.2 4.8 4.0 1.4 1.2 1.2 1.5 5.6 3.8 1.3 1.3 1.4 1.2

9.7 29.0 21.0 46.0 34.5 118 2.9 13.0 4.5 17.3 3.7c 20.7c 2.65 15 8.97 51.5 450.4 139.0 154.2 67.3 160.6 79.1 4500 820 88.0 156.0 843 111

Mina

SDb

CLIL%

2.4 4.6 3.3 8.7 16.7 25.4 0.6 1.9 1.4 4.1 2.8c 17.1c 0.42 2.76 3.81 21.7 121.0 37.0 18.0 4.3 10.0 8.8 1750 320 24.2 33.5 101 13

25 16 17 20 48 22 21 16 31 24 68c 71c 16 18 42 43 27 28 12 6.4 6.4 11 39 39 27 22 12 12

aMin,

minimum; Max, maximum; LP, lectin precipitation. concentrations for sample pools A and B represent the group mean ± SD obtained from all values reported for the specific marker and method. cResults from all RIAs included. bThe

assays for bone resorption markers [36, 39, 40], although largely discrepant results have been reported for serum CTX: Wichers et al. [36] reported a daily amplitude of serum CTX of up to 2200% (Fig. 3), while Rosen et al. observed a much lower degree of variability [40]. In our hands, diurnal changes of serum CTX of 200–300% were the rule (Seibel et al. unpublished results). Diurnal variation appears not to be affected by age, menopause, bed rest, physical activity, or season [21, 33, 44–47]. Although in postmenopausal women bone turnover is higher than in premenopausal women, the circadian variation is similar for both pre- and postmenopausal women and, thus, is not influenced by sex hormones [33, 35]. In osteoporotic women, the increase in bone resorption at night may persist longer into the morning, thus accounting in part for the greater bone loss in these individuals [30]. However, this could not be confirmed for elderly women with osteopenia [33]. The circadian rhythm is also not influenced by season [44]. The only factor with a pronounced

Figure 3

Individual serum CTX concentration profiles in a 24-h time period in six healthy male volunteers. From reference [36].

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influence on circadian variation is fasting, which can reduce the variation significantly by about 25% [21]. It has been reported that serum levels of CTx are influenced by food intake (Fig. 2), in contrast to the levels of NTx and ICTP [48]. Overall the etiology of diurnal variations is unknown. Several hormones, such as parathyroid hormone, growth hormone, or cortisol show diurnal changes and may therefore be involved in the generation of diurnal changes in bone metabolism [49–52]. There is some evidence that food intake is influencing the variation observed in biochemical markers of bone resorption over day and night and that this influence is independent of gender and menopause. Intake of glucose, protein, or fat resulted in a decrease in bone resorption, part of this response appears to be due to variation in serum insulin [46]. Independent of this, there is wide agreement that controlling the time of sampling is crucial in order to obtain clinically relevant information from bone markers. 2. Day-to-day Variability

Intraindividually, biochemical markers of bone turnover not only vary within a single day but in most cases also between consecutive days [6, 10, 29]. This phenomenon is called between-day or day-to-day variability and is apparently due to genuine variations in marker levels and not to analytical imprecision. In general, serum markers show less day-to-day variability than markers of bone turnover measured in urine [5, 38–40, 53]. This may be related to errors introduced by variations in urine collection, kidney function, urinary creatinine, etc. The dayto-day variation in the urinary excretion of PYD and DPD, measured by HPLC and corrected for creatinine, ranges between 16–26% [6, 10, 41, 54]. Similar results have been reported for free pyridinoline by EIA (7–25%), for the N-terminal collagen type I telopeptide (NTx, 13–35%), for the C-terminal collagen type I telopeptide (serum CTx, 8–10%; urinary CTx 12–35%), for tartrateresistant acid phosphatase (TRAP, 10–12%) [10, 39, 55–58]. Day-to-day variability adds considerably to the total variation of biochemical markers of bone turnover. In contrast to diurnal variations, day-to-day variability cannot be controlled. 3. Menstrual Variability

Bone turnover varies with the menstrual cycle with an overall amplitude of approximately 10–20% [5, 59–62]. Although not all available data show the same degree of change, there is enough evidence to support the suggestion that bone formation is higher during the luteal than the follicular phase [59], whereas bone resorption is higher during the mid-follicular, late-follicular, and early luteal phase [61]. Cyclical changes in bone turnover have

also been reported in postmenopausal women treated with sequential estrogen/gestagen regimens, showing decreases during estrogen treatment and increases during gestagen treatment [63, 64]. In clinical practice, however, variability due to the menstrual cycle is of little relevance, as most women evaluated for osteoporosis are postmenopausal. However, in premenopausal women with metabolic bone disease, menstrual variability should be taken into account, and the timing for sampling is probably best during the first 3–7 days of the menstrual cycle. 4. Seasonal Variability

Bone turnover and its regulation seem to vary with seasonal changes. Many studies have shown that serum 25-OH vitamin D and urinary calcium are higher in summer than in winter, and that parathyroid hormone levels tend to increase during winter [65–71]. Fewer data are available as to whether season influences bone turnover itself [66, 72–76]. Recently it has been shown, in most [71–73, 75, 77–80] but not all [66, 74, 81–83] studies that bone turnover is accelerated during winter, with more pronounced changes in females than in males [72, 78]. Seasonal high-turnover tends to coincide with a significant reduction in serum 25-OH vitamin D, suggesting that the increase in bone turnover during the winter period may be due, at least in part, to subclinical vitamin D deficiency and secondary hyperparathyroidism [72, 77] (Fig. 4). Interestingly it has been shown recently in a prospective observational study, that the frequency of falls and of hip and wrist fractures follow the same periodicity [77]. 5. Growth, Age, and Gender

During early childhood and then again during the pubertal growth spurt, biochemical markers of bone turnover are significantly higher than during adulthood because of the contribution of growth, and modeling as well as remodeling of bone [84–95] (Fig. 5). In girls, peak bone marker levels are observed approximately 2 years earlier than in boys, and estradiol seems to be the major determinant of the increase in bone turnover. In men between 20 and 30 years of age, bone turnover markers are usually higher than in women of the same age bracket. After the age of 50, most bone turnover markers tend to increase with further aging (Fig. 6), but less in men than in women. In the latter, the age-related increase in bone turnover is more pronounced due to the menopause, when both bone resorption and formation increase by about 50–100% [96–100]. 6. Nonskeletal Diseases and Clearance-Related Factors

A number of nonskeletal diseases have been shown to strongly affect bone turnover markers. These conditions

Chapter 34 Variability in the Measurement of Biochemical Markers of Bone Turnover

571

Figure 4

Circannual variation of calciotropic hormones and markers of bone turnover in controls (open circles, broken line, n =16) and subjects supplemented with oral vitamin D and calcium during the winter months (filled circles, line, n = 27; the intervention period is symbolized by an open bar). Data are presented as mean values calculated as the percentage of the respective mesor (rhythm-adjusted annual mean). From reference [80].

mostly relate to impairments in the clearance and/or metabolism of the components measured. Thus, even moderate impairment of renal function (GFR 50 ml/min) has been shown to have significant effects on the serum levels of osteocalcin [101, 102], of bone sialoprotein [96], and of the collagen type I telopeptides (NTx, CTx, ICTP) [12, 103]. In contrast, the excretion of pyridinium crosslinks usually does not change until a creatinine clearance of <25 ml/min is reached [41, 104, 105]. Serum levels of TRAP do not accumulate in renal failure [43, 106, 107]. Impairment of hepatic function usually leads to a nonspecific rise in the serum levels of the collagen type I propeptides (PINP, PICP), as these components are cleared by liver endothelial cells. 7. Fractures

Markers of bone formation and bone resorption increase during fracture healing [108–112]. The maximal

increase occurs in the first 2 months and may persist for 1 year (Fig. 7). The increase is greater for markers on bone formation (20–55%) than for markers of bone resorption (0–35%) [108–112]. The changes of the markers are dependent on the site of fracture.

III. STATISTICAL CONSIDERATION OF VARIABILITY A. Statistical Models for Estimation of Variability The partition of variability of a measurement is traditionally done by the classical theory of measurement [113], in which it is assumed that there is a true value of a measured trait in an individual, and that the observed value deviates from the true value by a random error. Accordingly, a measured bone turnover marker in an individual (denoted

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Figure 5

(A) Serum bone alkaline phosphatase (BAP) in normal male () and female (
) children as a function of age (from reference [95]). (B) Age-related changes in urinary DPyr excretion in healthy boys (▲) and girls () aged 3–18 years. Shaded area represents mean ± 2 SD for normal young adults. Asterisks indicate significant gender-related differences in corresponding age groups: *P < 0.05; **P < 0.01; ***P < 0.0001. From reference [94].

Figure 6 Age-related changes of biochemical markers of bone resorption (S-ICTP) and bone formation (S-PINP) in a cohort of communitydwelling healthy elderly men over 60 years. All men were participating in the Dubbo Osteoporosis Epidemiology Study.

by xi) can be considered to have two parts: a “true” value or “signal” (“error-free” value) denoted by m i, and a random error (“noise”) denoted by ei; that is: xi = mi + ei

(1)

Assuming that m i and ei are independently normally distributed with means of m T and 0, and variances of σ T2 2 and σ e , respectively, then the variance of the observed xi 2 2 2 is given by σ y = σ T + σ e . In other words, the total variance of a measurement is made up of two components: variance of differences among subjects in the “true” values, and variance of within-subject, including measurement 2 error. In this formulation, σ T is the variance due to differences among subjects, and σ e2 , is the variance due to within-subject variability.

Figure 7

Changes of biochemical markers during fracture healing in patients with osteoporosis. Mean percent changes from the initial values are plotted (*P < 0.05 compared with the initial values). Values of Pyr, Dpyr, CTx-ELISA at weeks 1, 2, and 4, Pyr, OCN-Mid at week 8 and OCN-Mid at week 24 were statistically significant compared with the initial values. From reference [109].

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For a sample of n representative subjects with ki multiple measurements over a period of time, it is possible to estimate the two sources of variance. Let the observed measure of a subject i taken at time j be xij (where i = 1, 2, 3, …, n, and j = 1, 2, 3, …, ki), then for each subject it is possible to calculate the sample mean and variance for the subject, k xi = 1/ ki ∑ ji= 1 xij and si2 = (1/ ki − 1) ∑ ( xij − xi )2 . The overall mean for the entire sample is simply: x = 1 / n ∑ in= 1 xi . An estimate of the within-subject vari2 ance σ e2 , denoted by Sw , is simply the average of individ2 ual variances, e.g.: Sw = [∑ in= 1 ( ki − 1)si2 ]/ ∑ in= 1 ( ki − 1). Furthermore, an estimate of the between-subject vari2 ance σ T , denoted by ST2 , is: ST2 = ( BMS − Sw2 )/ k , where BMS = ∑ in=1 ki ( xi − x )2 / ∑ in=1 ( ki − 1)n, and k is the average number of measurements per subject. Or, alternatively, ST2 and Sw2 can be obtained from a random-effect analysis of variance model via a state-of-the-art computer procedure such as PROC MIXED [114]. The within-subject variance Sw2 can further be partitioned into two components: one due to biological variation (Si2) and one due to analytical variation (Sa2 ). Traditionally, the within-subject coefficients of variation due to withinsubject (CVw), can be calculated as the square root of the within-subject variance component divided by the overall mean. The within-subject CVw can be further partitioned into within-subject biological variation (CVi) and analytical variation (CVa) using the relation CVi = CVw2 − CVa2 . Others [115] advocate the use of the within-subject standard deviation, Sw, as a measure of within-subject variability. Thus, a low value of CVw is considered as evidence of good measurement reliability. Previous studies have shown that the within-subject CVw differed significantly between markers (Table III). As can be seen from the table, serum

Table III.

markers of bone turnover generally have lower CVw than urinary markers. In short-term study (less than 1 year), serum markers of bone formation vary by around 10% within any given patient. However, measurements of urinary markers can differ by as much as 30% in one person, even on the same day [116]. For free cross-link determinations, the CVw ranges between 13% and 15%, although it is closer to 20% for urine N-telopeptides [117]. Although the coefficient of variation (CV) has been used extensively in the assessment of within-subject variability, it has certain limitations because its denominator (the sample mean) is very arbitrary [118, 119]. Consequently, the CV can be artificially small if the measured value of a measure is large such as the case of PINP in the above table. An alternative measure of within-subject variability is the coefficient of reliability (R) which is estimated as the ratio of variance of true value over total variance [113], i.e. R=

ST2

(2)

Sw2 + ST2

Because the variances are positive, it follows that the value of R is always between 0 (which indicates absolute unreliability) and 1 (which indicates absolute reliability). Thus, when the within-subject variability (as indicated by Sw) is low relative to the between-subject variability (ST), the value of R is usually large. As the formulation shows, R is not just a measure of within-subject variability, it is also in fact an estimate of the correlation between the observed and “true” (unmeasured) value for a subject. In common epidemiological studies, when the relationship between a categorical outcome (such as fracture) and a measure of bone turnover with a coefficient of reliability

Analytic and Biologic Coefficient of Variation (CV, %) in Selected Bone Formation and Resorption Markers

Marker

Within-subject CV (%)

Osteocalcin

7–27

PICP PINP

9–10 6–19

BSAP

7–9

urinary Pyr

10–26

urinary NTx

10–33

urinary CTx Serum CTx Serum ICTP

26–47 9.3 9–10

Biologic CV (%)

19.2

Analytic CV (%)

2.6

7.9 20.7

3.5

References Gertz [39], Jensen [130], Panteghini [56], Hannon [131] Hannon [131], Stevenson [132] Stevenson [132], Hannon [131], Scariano [133], Alvarez [134], Nguyen [120] Panteghini [56], Hannon [131], Scariano [133] Jensen [130], Jensen [29], McLaren[41], Ginty [135], Colwell [6] Gertz [39], Popp-Snijder [55], Hannon [131], Jensen [29], Alvarez [134], Scariano [133], Eastell [136] Alvarez [134], Hannon [131], Borderie [137] Alvarez [134] Hannon [131], Stevenson [132]

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of r, the magnitude of association will be a downward bias by 100(1 − r). It turns out that this coefficient is quite useful for the estimation of a subject’s “true” value (which is demonstrated below).

B. Regression Toward the Mean (RTM) A common problem associated with the within-subject variability is the phenomenon in which subjects with extreme measurements at baseline tend to regress toward the mean in subsequent visits. More than 100 years ago, this phenomenon was historically described by Francis Galton who noted that tall parents tended to have children who were shorter than they, although still above the average height, and conversely, that the children of short parents were still shorter than average, but taller than their parents [120]. The phenomenon of RTM is very common in virtually all medical studies and can pose serious problems for the interpretation of data, leading to erroneous conclusion with negative impact on the health care system [121]. In the field of osteoporosis research, this has been noted by Nguyen et al. [122], Chapurlat et al. [123], and Cummings et al. [124]. To understand the phenomenon of RTM and its relation to the within-subject variability, it is instructive to consider some theoretical aspects of the RTM. Consider a baseline measurement (x0) and a follow-up measurement (x1) for a group of individuals. If the measurements are errorless, and there is no real change during the period, then an individual’s measurement at baseline is exactly equal to the follow-up measurement (line of identity in Fig. 8). If, on the other hand, there is random within-subject variability, and under the same assumption of no change, the correlation between the baseline and follow-up measurements is less than 1, as represented by the dotted line. Because of this imperfect correlation, an individual with lower (x11) or higher (x12) than the baseline average will tend to be closer to the average in the follow-up measurement (d′1 < d1 and d′2 < d2). Consequently, the higher- or lower-value individuals tend to have smaller observed change. This theoretical result suggests that, for example, in clinical studies designed to select individuals with extreme (low or high) values of a bone turnover marker (or bone mineral density), it is expected that in the absence of any real change or therapeutic effect, the individuals will have a greater change in subsequent visits. For the sake of illustration, consider a hypothetical study with 10 000 subjects whose ICTPs were measured at baseline and follow-up. Assume that the mean (± standard deviation) of ICTP at baseline is 4.0 ± 1.2 µg/L, that there was no treatment effect

Figure 8

Illustration of regression to the mean phenomenon: Baseline values are represented in the x-axis; follow-up measurements are shown in the y-axis. The line of identity (perfect correlation) is shown by the solid line. The regression line is shown in the dotted line. When there is no measurement error, the baseline value is exactly equal to the follow-up value. When there is measurement and within-subject variability, individuals with lower/higher than average at baseline tend to be closer to the average during the subsequent measurement.

(i.e., average difference between baseline and follow-up is 0), and that the within-subject correlation in ICTP is 0.7, then the expected changes stratified by baseline quartile of ICTP are as shown in Table IV. However, even with uncontrolled and nonselected studies, empirical evidence of the presence of RTM has also been noted. In a sample of 909 healthy men and women aged between 18 and 61 years without any intervention, whose PINP and ICTP were repeatedly measured in 2-week intervals, the average change was, as expected, minimal (0.5 ± 4.5% for PINP and 0.9 ± 10.7% for ICTP). However, those in the lower quartile of baseline PINP and ICTP measurements had subsequently experienced increase in the measurements; in contrast, those with the highest quartile of baseline PINP and ICTP

Table IV. Quartile of ICTP at baseline 1 2 3 4

Effect of Regression Toward the Mean Expected mean ± SD of change from baseline (µg/L) 0.47 ± 0.88 0.16 ± 0.87 −0.12 ± 0.85 −0.45 ± 0.89

Note: Data were simulated for 10 000 pseudo-patients whose ICTP measurements are normally distributed with mean being 4.0 µg/L, standard deviation 1.2 µg/L, and within-subject correlation 0.7. The mean and SD of ICTP at baseline and follow-up are exactly the same.

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measurements had subsequently experienced decrease in the measurements (Table V).

C. Estimation of “True Value” for an Individual Thus, the presence of the within-subject variability in bone turnover measures leaves clinicians and researchers in a difficult position, which can be summarized into two questions: (i) Given an observed level of a bone marker for an individual, what is the individual’s likely true or long-term level?; and (ii) Given an observed change in the marker, to what extent does the change reflect a “true” change? From a common sense point of view, an individual’s true value of a bone turnover marker can be estimated as the average of a very large number of measurements over a relatively short period of time. Nevertheless, even with the large number of repeated measurements to capture the normal within-individual variability, one is still faced with the problem of having to make inference regarding that patient’s true level. Making correct inference requires some estimates of the RTM parameters. Suppose that the “true” values in the population have a mean m and variance σ T2 . If an individual’s single measurement of a marker is denoted by xi, under the Bayesian theory [125], the best estimate of his/her “true” value (mi), denoted by ti, can be obtained by combining the two sources of errors, the population and the measurement, with weighting equal to the inverse of their variances, e.g., ti =

µ / sT2 + xi / sw2 1/ sT2 + 1/ sw2

)

= Rxi + (1 − R µ

(3)

This is also known as the “empirical Bayes” estimate [126]. Comparison between the terms (1 − R)m and Rxi indicates that the coefficient reliability loaded at the individual’s observed value stronger than the population mean. When the individual observed value exceeds the population mean (xi > m), the individual’s estimated true value is less than the observed value (xi < ti); on the other hand,

when the individual observed value is less than the population mean, the estimated true value is higher than the observed value. The estimated variance of this estimate of the true value is: 2

2 ti

s

2

⎛ s2 ⎞ ⎛ s2 ⎞ = ⎜ 2 w 2 ⎟ sT2 + ⎜ 2 T 2 ⎟ sw2 = Rsw2 (4) ⎝ sT + sw ⎠ ⎝ sT + sw ⎠

and the two-sided 95% credible interval for mi can be estimated as between ti − 1.96sw R and ti + 1.96sw R . The above formulation is valid under the following assumptions that the observed value of the marker (xi) is normally distributed, and that the within-subject variance is normally distributed and is independent from the underlying measured levels. Under these assumptions, it can be stated that the difference between mi and m is normally distributed with mean R(xi – m) and variance of 2 Rσ w . Consequently, the estimated true value for an individual falls between the observed value and the population mean. For the sake of illustration, suppose that the following estimates were obtained for the data set shown in Table V: mean (± standard deviation) of ICTP, PINP, and PIIINP was 1.75 ± 0.35, 4.50 ± 0.55, and 1.61 ± 0.31 µg/L, respectively. The estimated within-subject variance for ICTP, PINP, and PIIINP was 0.66, 255, and 0.58 µg/L2, respectively. The coefficient of reliability for ICTP, PINP, and PIIINP was estimated to be 0.92, 0.96, and 0.88, respectively. Thus, for an individual with an ICTP of 2.45 µg/L (which is two standard deviations above the population mean), the individual’s estimated true value, using (3), is: 2.45 × 0.92 + (1 − 0.92) × 1.75 = 2.39 µg/L. The estimated variance of this true value, by (4), is 0.66 × 0.92 = 0.61 µg/L2, and the 95% credible interval for the individual’s true ICTP value ranged between 2.39 − 1.96 0.61 = 0.86 µg/L and 2.39 + 1.96 0.61 = 3.92 µg/L.

Table V. Percent Change in PINP and ICTP in a Group of 909 Men and Women without any Intervention during a 2-week Follow-up Quartile: Baseline PINP (µg/L) 1: < 4.1 2: 4.1–4.4 3: 4.5–4.8 4: >4.8 All quartiles

Mean percent change from baseline 2.2 ± 6.4 0.9 ± 4.3 0.3 ± 3.5 −0.7 ± 3.5 0.5 ± 4.5

Quartile: Baseline ICTP (µg/L) 1: <1.49 2: 1.49–1.72 3: 1.73–1.94 4: >1.94 All quartiles

Mean percent change from baseline 3.6 ± 16.3 0.7 ± 9.2 0.1 ± 7.2 −0.4 ± 8.9 0.9 ± 10.7

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D. Estimation of “True Change” for an Individual: A Frequentist Approach A common method of assessing whether an observed change in a marker is “significant” (e.g., beyond the normal within-subject variability) is the so-called “least significant change” (LSC) or “reference chance value” (RCV, see Chapter 21). The LSC or RCV is defined as 1.96 2CVw . The construction of this cut-off value is based on the assumption for any two consecutive measurements within an individual, each with a within-subject standard deviation of Sw (and hence CVw), then the variance of the differ2 ence between the two measurements is 2Sw . With a 95% 2 confidence interval, any change of 1.96 2 Sw or 1.96 2CVw is considered to be “clinically meaingful”. However, the above formulation is valid only if the following assumptions are met: (i) the variances are normally distributed; (ii) the analytic variance and withinsubject variance are independent of the underlying measurement; and (iii) any two consecutive measurements are independent (unrelated). While the first and second assumptions can be reasonably expected to be met, the third assumption is unlikely to be met because for an individual any two measurements are expected to be correlated. Furthermore, the correlation is expected to decrease with longer duration of follow-up. In fact, it can be shown that the variance of difference between two consecutive measurements within an individual is 2( Si2 + Sa2 )[(1 − r ) + rSa2 /( Si2 + Sa2 )], where r is the correlation coefficient between two measurements. Since r is expected to be positive and is less than or equal to 1, the LSC cut-off almost always underestimates the biologic effect and overestimates the effect of withinsubject variability. As noted in Chapter 21 and by the above demonstration, the LSC formulation tends to underestimate the biologic effect and overestimate the effect of within-subject random variability. Moreover, a disadvantage of the LSC formulation is that it can only indicate whether or not an observed change is due to random variability; it doesn’t indicate whether or not the change is clinically significant. In the following section, a Bayesian approach is proposed as an alternative method to quantify the clinical importance of an observed change.

E. Estimation of “True Change” for an Individual: A Bayesian Approach A better and more logical method to assess the change within an individual is by adjusting for the effect due to RTM within the Bayesian formulation. This method has

a sound empirical justification. In an analysis of a clinical trial, Chapurlat et al. [123] noted that patients who had experienced the largest decrease in bone turnover markers after 6 months of antiresorptive therapy subsequently experienced a significant increase in the following 6 months of treatment. Similar trends were also observed for bone mineral density [124]. Indeed, empirical data have revealed an inverse association between bone loss and initial level of bone resorption, in which individuals with higher baseline value tended to experience higher rate of bone loss than those with lower baseline value. Therefore, when consecutive measurements of bone turnover in an individual are obtained, the question arises as to what extent any observed change reflects a “true” change. The concept of RTM is also useful here [123, 127–129]. As in Section C, the Bayesian process of combining information from an individual and the population can be applied to estimate the true change in an individual. Let x1i and x2i be measured values at baseline and follow-up, respectively. The observed change is di = x2i − x1i. However, the adjusted observed change is: di = x2i − t1i = x2i − Rx1i − (1 − R)m. The variance of di is the sum of the variance of its two components, viz: sδ2 = Rsw2 + sw2 = sw2 (1 + R). Assuming that, from previous studies, the real changes are known to be normally distributed with mean ∆ and 2 variance of σ ∆ , then the estimate of a “true” change for an individual is: ∆ / s∆2 + δ i / sδ2 dˆi = 1/ s∆2 + 1/ sδ2

(5)

The estimated variance is:

( )

s dˆi 2

=

s∆2 sδ2

(6)

s∆2 + sδ2

and the two-sided 95% credible interval for the “true” 2 2 change is estimated as between dˆi ± 1.96 × s∆ sd ' / s∆ + sδ . As an illustration, consider the data obtained from a noninterventional study of 101 elderly men aged 60+, whose ICTPs have been measured at baseline and 2 years later. The overall mean ± SD of ICTP in the sample was m = 4.41 ± 1.29 µg/L. The mean (± SD) of ICTP at baseline and follow-up was 4.25 ± 1.13 and 4.57 ± 1.68 µg/L, respectively. The average of observed change was 0.32 µg/L with variance of 1.64 µg/L2, and these values are taken as estimates of ∆ and s∆2 , respectively. Furthermore, the within-subject variance was estimated at 0.90 µg/L2, which yielded an estimated coefficient of reliability of 0.57. The estimated variance of adjusted change for an individual was therefore: sδ2 = 0.9 (1 + 0.57 = 1.41 µg/L2. Suppose that the measured ICTP of a man in the study at baseline and follow-up was 2.0 and 5.0 mg/L, respectively,

)

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Chapter 34 Variability in the Measurement of Biochemical Markers of Bone Turnover

which represented an observed change of di = 3.0 µg/L. However, the adjusted change was di = 5.0 – 0.57(2.0) – (0.43)(4.41) = 1.97 µg/L. Therefore, the estimated “true change” for the man, by (5), was only dˆi = (0.32 /1.64 + 1.96 /1.41)/(1/1.64 + 1/1.41) = 1..2 µg/L, and the vari2 ance of the change was: s (dˆ ) = (1.64 × 1.41)/(1.64 + 1.41) = 0.76. The 95% credible interval of the change ranged between 1.2 − 1.96 0.76 = −0.51 and 0.38 + 1.96 0.76 = 2.91 µg/L. The flexibility of the Bayesian approach also allows one to answer a pertinent question such as “if a true change of a certain magnitude has occurred, what is the probability that the change can be detected with an observed change?” To address this question, the probability of a true increase in ICTP of 1 µg/L and 2 µg/L for a given range of observed changes could be constructed (Fig. 9). For example, for the above hypothetical man with an observed change of 3 µg/L, the probability that a change of 2 µg/L had occurred was 0.87. On the other hand, there needs to be an observed increase of 2 µg/L before one can be 87% certain that the true increase of 1 µg/L has occurred. Furthermore, to be 95% certain that the true increase of 1 µg/L has occurred in an individual, there needs to be an observed increase of 2.5 µg/L in the individual.

This can be straightforwardly done by noticing the fact that the expected change for a given baseline value x is: E(X2 – X1 | X1 = x) = ∆ + (x – m)(R – 1), where X1 and X2 denotes the baseline and follow-up measurements of a bone turnover marker, respectively. Thus, an estimated “true change” for a group, denoted by ∆ˆ , can be obtained as: ∆ˆ = ( x 2 − x1 ) − ( x1 − µ )( R − 1) . The second part of this result is the amount of effect due to RTM. Another way to estimate the RTM effect is first to compute the individual adjusted changes da = x2 − x1 − (x1 − m)(R − 1), and then average the da to obtain ∆ˆ . To illustrate this method, a simulation of 10 000 pseudo subjects was carried out with the following parameters: (a) mean ± SD of ICTP at baseline were 4.0 ± 1.2 µg/L; (b) the treatment effect was –1.0 ± 0.5 µg/L; and (c) the within-subject correlation was 0.7. Suppose that only subjects with the highest and lowest quartiles of ICTP are studied, then the following outcomes are expected (Table VI). As can be seen from this simulation, those subjects with extreme baseline values (either in the lowest or highest quartile of ICTP) exhibit differential changes in ICTP, with those in the highest baseline quartile being associated with the greatest decrease (−1.41 µg/L), although the adjusted change is 0.98 (which is, as expected, close to 1 µg/L as defined by the simulation) and the rest was due to RTM artefactual effect.

F. Estimation of “True Change” for a Group of Patients

IV. SUMMARY

For a group of individuals, one is often interested in estimating the effect of RTM on the observed change.

Numerous factors influence bone turnover, but there are even more sources of variability that need to be taken into account when measuring bone turnover with biochemical markers. To minimize some of the limitations linked to pre-analytical and analytical variability, standardized sampling and sample handling are mandatory to obtain reliable results. Controllable factors such as the mode of sample collection, samples handling and storage, diurnal and menstrual rhythms, pre-sampling exercise, and pre-sampling diet should be taken care of wherever possible. Laboratories are encouraged to establish their own reference ranges and to use genderand age-specific reference intervals. In order to further Table VI. Effect of RTM when Subjects are Selected Based on Extreme Values

Probability of a true change in ICTP by 1 µg/L and 2 µg/L for an observed change (x-axis). For a “true change” of 2 µg/L, an observed change of approximately 2.7 µg/L must be reached to obtain a probability of 80% of such change has indeed occurred. For a “true change” of 1 mg/L, an observed change of approximately 1.6 µg/L must be reached to obtain a probability of 80%.

Figure 9

Quartile

Baseline

Follow-up

Observed change

Adjusted change

1 4

2.50 ± 0.67 5.41 ± 0.66

2.00 ± 1.07 4.00 ± 1.06

–0.50 ± 1.00 –1.41 ± 1.00

–0.97 –0.98

578 reduce variability, standardization of bone marker assays and routine proficiency testing programs are strongly recommended. Changes in marker measurements should always be interpreted against the background of the respective marker’s within-subject variability. As a rule, markers showing large changes in response to disease processes or interventions also show substantial degrees of variability. This is so because the phenomenon of regression to the mean is ubiquitous in all clinical studies. For example, in randomized clinical trials, participants are usually selected because they have extreme values of an exposure variable (i.e., low bone mineral density, and hence high values of bone resorption marker), the RTM effect tends to yield unexpectedly pronounced “effect”, even though part of the result may have nothing to do with the intervention considered. In randomized clinical trials designed to evaluate the efficacy of a therapy the use of control group, if ethically possible, is usually sensible because it allows a direct adjustment of treatment effect. However, in the absence of a control group, the estimation of treatment efficacy should be adjusted for the RTM effect. In the presence of within-subject variability and RTM effect, it is often interesting to know whether an individual measurement represents the individual’s true value, or to what extent an observed change in the individual represents a biologic, but not stochastic, change. The Bayesian framework as presented in this chapter is helpful to address the two questions. In order to have an idea about the individual’s true value, one has to weight the data from a group of subjects where the individual was sampled from (e.g. known information) and combine with the observed data to provide a posterior distribution of the individual’s “true” value. As for the assessment of an individual’s true change, the LSC formulation tends to underestimate biologic effects and consequently overemphasize stochastic effects. Because the within-subject random variability may be constant over time for one individual, but likely varies between individuals, therefore a single criterion can never be applied to all individuals. To address this deficiency of the LSC approach, the Bayesian approach provides a much more logical answer. Instead of working based on the null and negative hypothesis (e.g., the change is within the effect of within-subject fluctuation), the Bayesian approach works on a forward-looking and more meaningful hypothesis (e.g., given an observed change, what is the probability that a 5% change has taken place). The Bayesian model presented here can help clinicians make informed decisions about treatment and information that may be conveyed to the patient.

TUAN V. NGUYEN ET AL.

References 1. Armstrong, B. G. (1990). The effects of measurement errors on relative risk regressions. Am. J. Epidemiol. 132, 1176–1184. 2. Seibel, M. J. (2000). Molecular markers of bone turnover: biochemical, technical and analytical aspects. Osteoporos. Int. 11, S18–29. 3. Blumsohn, A., Hannon, R. A., and Eastell, R. (1995). Apparent instability of osteocalcin in serum as measured with different commercially available immunoassays. Clin. Chem. 41, 318–319. 4. Lang, M., Seibel, M. J., Zipf, A., and Ziegler, R. (1996). Influence of a new protease inhibitor on the stability of osteocalcin in serum. Clin. Lab. 42, 5–10. 5. Li, Y., Woitge, H., Kissling, C., Lang, M., Oberwittler, H., Karmatschek, M., Armbruster, F. P., von Schickfus, A. R., Ziegler, R., and Seibel, M. J. (1998). Biological variability of serum immunoreactive bone sialoprotein. Clin. Lab. 44, 553–555. 6. Colwell, A., Russell, R. G., and Eastell, R. (1993). Factors affecting the assay of urinary 3-hydroxy pyridinium crosslinks of collagen as markers of bone resorption. Eur. J. Clin. Invest. 23, 341–349. 7. Gerrits, M. I., Thijssen, J. H., and van Rijn, H. J. (1995). Determination of pyridinoline and deoxypyridinoline in urine, with special attention to retaining their stability. Clin. Chem. 41, 571–574. 8. Beardsworth, L. J., Eyre, D. R., and Dickson, I. R. (1990). Changes with age in the urinary excretion of lysyl- and hydroxylysylpyridinoline, two new markers of bone collagen turnover. J. Bone Miner. Res. 5, 671–676. 9. Seibel, M. J., Gartenberg, F., Silverberg, S. J., Ratcliffe, A., Robins, S. P., and Bilezikian, J. P. (1992). Urinary hydroxypyridinium cross-links of collagen in primary hyperparathyroidism. J. Clin. Endocrinol. Metab. 74, 481–486. 10. Ju, H. S., Leung, S., Brown, B., Stringer, M. A., Leigh, S., Scherrer, C., Shepard, K., Jenkins, D., Knudsen, J., and Cannon, R. (1997). Comparison of analytical performance and biological variability of three bone resorption assays. Clin. Chem. 43, 1570–1576. 11. Bonde, M., Qvist, P., Fledelius, C., Riis, B. J., and Christiansen, C. (1994). Immunoassay for quantifying type I collagen degradation products in urine evaluated. Clin. Chem. 40, 2022–2025. 12. Risteli, J., Elomaa, I., Niemi, S., Novamo, A., and Risteli, L. (1993). Radioimmunoassay for the pyridinoline cross-linked carboxy-terminal telopeptide of type I collagen: a new serum marker of bone collagen degradation. Clin. Chem. 39, 635–640. 13. Lau, K. H., Onishi, T., Wergedal, J. E., Singer, F. R., and Baylink, D. J. (1987). Characterization and assay of tartrate-resistant acid phosphatase activity in serum: potential use to assess bone resorption. Clin. Chem. 33, 458–462. 14. Schmeller, N. T., and Bauer, H. W. (1983). Circadian variation of different fractions of serum acid phosphatase. Prostate 4, 391–395. 15. Colwell, A., Hamer, A., Blumsohn, A., and Eastell, R. (1996). To determine the effects of ultraviolet light, natural light and ionizing radiation on pyridinium cross-links in bone and urine using high-performance liquid chromatography. Eur. J. Clin. Invest. 26, 1107–1114. 16. Walne, A. J., James, I. T., and Perrett, D. (1995). The stability of pyridinium crosslinks in urine and serum. Clin. Chim. Acta. 240, 95–97. 17. Blumsohn, A., Colwell, A., Naylor, K., and Eastell, R. (1995). Effect of light and gamma-irradiation on pyridinolines and telopeptides of type I collagen in urine. Clin. Chem. 41, 1195–1197. 18. Heymsfield, S. B., Arteaga, C., McManus, C., Smith, J., and Moffitt, S. (1983). Measurement of muscle mass in humans: validity of the 24-hour urinary creatinine method. Am. J. Clin. Nutr. 37, 478–494. 19. Smith, S. M., Dillon, E. L., DeKerlegand, D. E., and Davis-Street, J. E. (2004). Variability of collagen crosslinks: impact of sample collection period. Calcif. Tissue Int. 74, 336–341. 20. Devgun, M. S., Paterson, C. R., and Martin, B. T. (1981). Seasonal changes in the activity of serum alkaline phosphatase. Enzyme 26, 301–305.

Chapter 34 Variability in the Measurement of Biochemical Markers of Bone Turnover 21. Qvist, P., Christgau, S., Pedersen, B. J., Schlemmer, A., and Christiansen, C. (2002). Circadian variation in the serum concentration of C-terminal telopeptide of type I collagen (serum CTx): effects of gender, age, menopausal status, posture, daylight, serum cortisol, and fasting. Bone 31, 57–61. 22. Ashizawa, N., Ouchi, G., Fujimura, R., Yoshida, Y., Tokuyama, K., and Suzuki, M. (1998). Effects of a single bout of resistance exercise on calcium and bone metabolism in untrained young males. Calcif. Tissue Int. 62, 104–108. 23. Wallace, J. D., Cuneo, R. C., Lundberg, P. A., Rosen, T., Jorgensen, J. O., Longobardi, S., Keay, N., Sacca, L., Christiansen, J. S., Bengtsson, B. A., and Sonksen, P. H. (2000). Responses of markers of bone and collagen turnover to exercise, growth hormone (GH) administration, and GH withdrawal in trained adult males. J. Clin. Endocrinol. Metab. 85, 124–133. 24. Welsh, L., Rutherford, O. M., James, I., Crowley, C., Comer, M., and Wolman, R. (1997). The acute effects of exercise on bone turnover. Int. J. Sports Med. 18, 247–251. 25. Woitge, H. W., Friedmann, B., Suttner, S., Farahmand, I., Muller, M., Schmidt-Gayk, H., Baertsch, P., Ziegler, R., and Seibel, M. J. (1998). Changes in bone turnover induced by aerobic and anaerobic exercise in young males. J. Bone Miner. Res. 13, 1797–1804. 26. Seibel, M. J., Lang, M., and Geilenkeuser, W. J. (2001). Interlaboratory variation of biochemical markers of bone turnover. Clin. Chem. 47, 1443–1450. 27. Mautalen, C. A. (1970). Circadian rhythm of urinary total and free hydroxyproline excretion and its relation to creatinine excretion. J. Lab. Clin. Med. 75, 11–18. 28. Schlemmer, A., and Hassager, C. (1999). Acute fasting diminishes the circadian rhythm of biochemical markers of bone resorption. Eur. J. Endocrinol. 140, 332–337. 29. Jensen, J. E., Kollerup, G., Sorensen, H. A., and Sorensen, O. H. (1997). Intraindividual variability in bone markers in the urine. Scand. J. Clin. Lab. Invest. Suppl. 227, 29–34. 30. Eastell, R., Calvo, M. S., Burritt, M. F., Offord, K. P., Russell, R. G., and Riggs, B. L. (1992). Abnormalities in circadian patterns of bone resorption and renal calcium conservation in type I osteoporosis. J. Clin. Endocrinol. Metab. 74, 487–494. 31. Schlemmer, A., Hassager, C., Jensen, S. B., and Christiansen, C. (1992). Marked diurnal variation in urinary excretion of pyridinium crosslinks in premenopausal women. J. Clin. Endocrinol. Metab. 74, 476–480. 32. Pagani, F., and Panteghini, M. (1994). Diurnal rhythm in urinary excretion of pyridinium crosslinks. Clin. Chem. 40, 952–953. 33. Schlemmer, A., Hassager, C., Pedersen, B. J., and Christiansen, C. (1994). Posture, age, menopause, and osteopenia do not influence the circadian variation in the urinary excretion of pyridinium crosslinks. J. Bone Miner. Res. 9, 1883–1888. 34. Bollen, A. M., Martin, M. D., Leroux, B. G., and Eyre, D. R. (1995). Circadian variation in urinary excretion of bone collagen cross-links. J. Bone Miner. Res. 10, 1885–1890. 35. Eastell, R., Simmons, P. S., Colwell, A., Assiri, A. M., Burritt, M. F., Russell, R. G., and Riggs, B. L. (1992). Nyctohemeral changes in bone turnover assessed by serum bone Gla-protein concentration and urinary deoxypyridinoline excretion: effects of growth and ageing. Clin. Sci. (Lond) 83, 375–382. 36. Wichers, M., Schmidt, E., Bidlingmaier, F., and Klingmuller, D. (1999). Diurnal rhythm of CrossLaps in human serum. Clin. Chem. 45, 1858–1860. 37. Sarno, M., Powell, H., Tjersland, G., Schoendorfer, D., Harris, H., Adams, K., Ogata, P., and Warnick, G. R. (1999). A collection method and high-sensitivity enzyme immunoassay for sweat pyridinoline and deoxypyridinoline cross-links. Clin. Chem. 45, 1501–1509. 38. Greenspan, S. L., Dresner-Pollak, R., Parker, R. A., London, D., and Ferguson, L. (1997). Diurnal variation of bone mineral turnover in elderly men and women. Calcif. Tissue Int. 60, 419–423.

579

39. Gertz, B. J., Clemens, J. D., Holland, S. D., Yuan, W., and Greenspan, S. (1998). Application of a new serum assay for type I collagen crosslinked N-telopeptides: assessment of diurnal changes in bone turnover with and without alendronate treatment. Calcif. Tissue Int. 63, 102–106. 40. Rosen, H. N., Moses, A. C., Garber, J., Iloputaife, I. D., Ross, D. S., Lee, S. L., and Greenspan, S. L. (2000). Serum CTX: a new marker of bone resorption that shows treatment effect more often than other markers because of low coefficient of variability and large changes with bisphosphonate therapy. Calcif. Tissue Int. 66, 100–103. 41. McLaren, A. M., Isdale, A. H., Whiting, P. H., Bird, H. A., and Robins, S. P. (1993). Physiological variations in the urinary excretion of pyridinium crosslinks of collagen. Br. J. Rheumatol. 32, 307–312. 42. Hassager, C., Risteli, J., Risteli, L., Jensen, S. B., and Christiansen, C. (1992). Diurnal variation in serum markers of type I collagen synthesis and degradation in healthy premenopausal women. J. Bone Miner. Res. 7, 1307–1311. 43. Hannon, R. A., Clowes, J. A., Eagleton, A. C., Al Hadari, A., Eastell, R., and Blumsohn, A. (2004). Clinical performance of immunoreactive tartrate-resistant acid phosphatase isoform 5b as a marker of bone resorption. Bone 34, 187–194. 44. Nielsen, H. K., Brixen, K., and Mosekilde, L. (1990). Diurnal rhythm and 24-hour integrated concentrations of serum osteocalcin in normals: influence of age, sex, season, and smoking habits. Calcif. Tissue Int. 47, 284–290. 45. Caillot-Augusseau, A., Vico, L., Heer, M., Voroviev, D., Souberbielle, J. C., Zitterman, A., Alexandre, C., and Lafage-Proust, M. H. (2000). Space flight is associated with rapid decreases of undercarboxylated osteocalcin and increases of markers of bone resorption without changes in their circadian variation: observations in two cosmonauts. Clin. Chem. 46, 1136–1143. 46. Bjarnason, N. H., Henriksen, E. E., Alexandersen, P., Christgau, S., Henriksen, D. B., and Christiansen, C. (2002). Mechanism of circadian variation in bone resorption. Bone 30, 307–313. 47. Wolthers, O. D., Heuck, C., and Heickendorff, L. (2001). Diurnal variations in serum and urine markers of type I and type III collagen turnover in children. Clin. Chem. 47, 1721–1722. 48. Christgau, S. (2000). Circadian variation in serum CrossLaps concentration is reduced in fasting individuals. Clin. Chem. 46, 431. 49. Markowitz, M. E., Arnaud, S., Rosen, J. F., Thorpy, M., and Laximinarayan, S. (1988). Temporal interrelationships between the circadian rhythms of serum parathyroid hormone and calcium concentrations. J. Clin. Endocrinol. Metab. 67, 1068–1073. 50. Calvo, M. S., Eastell, R., Offord, K. P., Bergstralh, E. J., and Burritt, M. F. (1991). Circadian variation in ionized calcium and intact parathyroid hormone: evidence for sex differences in calcium homeostasis. J. Clin. Endocrinol. Metab. 72, 69–76. 51. Nielsen, H. K., Laurberg, P., Brixen, K., and Mosekilde, L. (1991). Relations between diurnal variations in serum osteocalcin, cortisol, parathyroid hormone, and ionized calcium in normal individuals. Acta. Endocrinol. (Copenh) 124, 391–398. 52. Ebeling, P. R., Butler, P. C., Eastell, R., Rizza, R. A., and Riggs, B. L. (1991). The nocturnal increase in growth hormone is not the cause of the nocturnal increase in serum osteocalcin. J. Clin. Endocrinol. Metab. 73, 368–372. 53. Takahashi, M., Kawana, K., and Nagano, A. (2002). Biological variability of biochemical markers of bone turnover in healthy women. Endocr. Res. 28, 257–264. 54. Vesper, H. W., Demers, L. M., Eastell, R., Garnero, P., Kleerekoper, M., Robins, S. P., Srivastava, A. K., Warnick, G. R., Watts, N. B., and Myers, G. L. (2002). Assessment and recommendations on factors contributing to preanalytical variability of urinary pyridinoline and deoxypyridinoline. Clin. Chem. 48, 220–235. 55. Popp-Snijders, C., Lips, P., and Netelenbos, J. C. (1996). Intra-individual variation in bone resorption markers in urine. Ann. Clin. Biochem. 33 (Pt 4), 347–348.

580 56. Panteghini, M., and Pagani, F. (1995). Biological variation in bonederived biochemical markers in serum. Scand. J. Clin. Lab. Invest. 55, 609–616. 57. Garnero, P., Mulleman, D., Munoz, F., Sornay-Rendu, E., and Delmas, P. D. (2003). Long-term variability of markers of bone turnover in postmenopausal women and implications for their clinical use: the OFELY study. J. Bone Miner. Res. 18, 1789–1794. 58. Christgau, S., Bitsch-Jensen, O., Hanover Bjarnason, N., Gamwell Henriksen, E., Qvist, P., Alexandersen, P., and Bang Henriksen, D. (2000). Serum CrossLaps for monitoring the response in individuals undergoing antiresorptive therapy. Bone 26, 505–511. 59. Nielsen, H. K., Brixen, K., Bouillon, R., and Mosekilde, L. (1990). Changes in biochemical markers of osteoblastic activity during the menstrual cycle. J. Clin. Endocrinol. Metab. 70, 1431–1437. 60. Schlemmer, A., Hassager, C., Risteli, J., Risteli, L., Jensen, S. B., and Christiansen, C. (1993). Possible variation in bone resorption during the normal menstrual cycle. Acta. Endocrinol. (Copenh) 129, 388–392. 61. Gorai, I., Chaki, O., Nakayama, M., and Minaguchi, H. (1995). Urinary biochemical markers for bone resorption during the menstrual cycle. Calcif. Tissue Int. 57, 100–104. 62. Zittermann, A., Schwarz, I., Scheld, K., Sudhop, T., Berthold, H. K., von Bergmann, K., van der Ven, H., and Stehle, P. (2000). Physiologic fluctuations of serum estradiol levels influence biochemical markers of bone resorption in young women. J. Clin. Endocrinol. Metab. 85, 95–101. 63. Johansen, J. S., Jensen, S. B., Riis, B. J., and Christiansen, C. (1990). Time-dependent variations in bone turnover parameters during 2 months’ cyclic treatment with different doses of combined estrogen and progestogen in postmenopausal women. Metabolism 39, 1122–1126. 64. Christiansen, C., Riis, B. J., Nilas, L., Rodbro, P., and Deftos, L. (1985). Uncoupling of bone formation and resorption by combined oestrogen and progestagen therapy in postmenopausal osteoporosis. Lancet 2, 800–801. 65. Juttmann, J. R., Visser, T. J., Buurman, C., de Kam, E., and Birkenhager, J. C. (1981). Seasonal fluctuations in serum concentrations of vitamin D metabolites in normal subjects. Br. Med. J. (Clin Res Ed) 282, 1349–1352. 66. Overgaard, K., Nilas, L., Johansen, J. S., and Christiansen, C. (1988). Lack of seasonal variation in bone mass and biochemical estimates of bone turnover. Bone 9, 285–288. 67. Scharla, S. H., Scheidt-Nave, C., Leidig, G., Woitge, H., Wuster, C., Seibel, M. J., and Ziegler, R. (1996). Lower serum 25-hydroxyvitamin D is associated with increased bone resorption markers and lower bone density at the proximal femur in normal females: a population-based study. Exp. Clin. Endocrinol. Diabetes 104, 289–292. 68. Morgan, D. B., Rivlin, R. S., and Davis, R. H. (1972). Seasonal changes in the urinary excretion of calcium. Am. J. Clin. Nutr. 25, 652–654. 69. Lips, P., Hackeng, W. H., Jongen, M. J., van Ginkel, F. C., and Netelenbos, J. C. (1983). Seasonal variation in serum concentrations of parathyroid hormone in elderly people. J. Clin. Endocrinol. Metab. 57, 204–206. 70. Krall, E. A., Sahyoun, N., Tannenbaum, S., Dallal, G. E., and DawsonHughes, B. (1989). Effect of vitamin D intake on seasonal variations in parathyroid hormone secretion in postmenopausal women. N. Engl. J. Med. 321, 1777–1783. 71. Chapuy, M. C., Schott, A. M., Garnero, P., Hans, D., Delmas, P. D., and Meunier, P. J. (1996). Healthy elderly French women living at home have secondary hyperparathyroidism and high bone turnover in winter. EPIDOS Study Group. J. Clin. Endocrinol. Metab. 81, 1129–1133. 72. Woitge, H. W., Scheidt-Nave, C., Kissling, C., Leidig-Bruckner, G., Meyer, K., Grauer, A., Scharla, S. H., Ziegler, R., and Seibel, M. J. (1998). Seasonal variation of biochemical indexes of bone turnover: results of a population-based study. J. Clin. Endocrinol. Metab. 83, 68–75.

TUAN V. NGUYEN ET AL. 73. Hyldstrup, L., McNair, P., Jensen, G. F., and Transbol, I. (1986). Seasonal variations in indices of bone formation precede appropriate bone mineral changes in normal men. Bone 7, 167–170. 74. Vanderschueren, D., Gevers, G., Dequeker, J., Geusens, P., Nijs, J., Devos, P., De Roo, M., and Bouillon, R. (1991). Seasonal variation in bone metabolism in young healthy subjects. Calcif. Tissue Int. 49, 84–89. 75. Thomsen, K., Eriksen, E. F., Jorgensen, J. C., Charles, P., and Mosekilde, L. (1989). Seasonal variation of serum bone GLA protein. Scand. J. Clin. Lab. Invest. 49, 605–611. 76. Douglas, A. S., Miller, M. H., Reid, D. M., Hutchison, J. D., Porter, R. W., and Robins, S. P. (1996). Seasonal differences in biochemical parameters of bone remodelling. J. Clin. Pathol. 49, 284–289. 77. Pasco, J. A., Henry, M. J., Kotowicz, M. A., Sanders, K. M., Seeman, E., Pasco, J. R., Schneider, H. G., and Nicholson, G. C. (2004). Seasonal periodicity of serum vitamin D and parathyroid hormone, bone resorption, and fractures: the Geelong Osteoporosis Study. J. Bone Miner. Res. 19, 752–758. 78. Woitge, H. W., Knothe, A., Witte, K., Schmidt-Gayk, H., Ziegler, R., Lemmer, B., and Seibel, M. J. (2000). Circaannual rhythms and interactions of vitamin D metabolites, parathyroid hormone, and biochemical markers of skeletal homeostasis: a prospective study. J. Bone Miner. Res. 15, 2443–2450. 79. Storm, D., Eslin, R., Porter, E. S., Musgrave, K., Vereault, D., Patton, C., Kessenich, C., Mohan, S., Chen, T., Holick, M. F., and Rosen, C. J. (1998). Calcium supplementation prevents seasonal bone loss and changes in biochemical markers of bone turnover in elderly New England women: a randomized placebo-controlled trial. J. Clin. Endocrinol. Metab. 83, 3817–3825. 80. Meier, C., Woitge, H. W., Witte, K., Lemmer, B., and Seibel, M. J. (2004). Supplementation with oral vitamin d3 and calcium during winter prevents seasonal bone loss: a randomized controlled open-label prospective trial. J. Bone Miner. Res. 19, 1221–1230. 81. Zittermann, A., Scheld, K., and Stehle, P. (1998). Seasonal variations in vitamin D status and calcium absorption do not influence bone turnover in young women. Eur. J. Clin. Nutr. 52, 501–506. 82. Patel, R., Collins, D., Bullock, S., Swaminathan, R., Blake, G. M., and Fogelman, I. (2001). The effect of season and vitamin D supplementation on bone mineral density in healthy women: a double-masked crossover study. Osteoporos. Int. 12, 319–325. 83. Blumsohn, A., Naylor, K. E., Timm, W., Eagleton, A. C., Hannon, R. A., and Eastell, R. (2003). Absence of marked seasonal change in bone turnover: a longitudinal and multicenter cross-sectional study. J. Bone Miner. Res. 18, 1274–1281. 84. Hodgkinson, A., and Thompson, T. (1982). Measurement of the fasting urinary hydroxyproline: creatinine ratio in normal adults and its variation with age and sex. J. Clin. Pathol. 35, 807–811. 85. Stepan, J. J., Tesarova, A., Havranek, T., Jodl, J., Formankova, J., and Pacovsky, V. (1985). Age and sex dependency of the biochemical indices of bone remodelling. Clin. Chim. Acta. 151, 273–283. 86. Krabbe, S., Hummer, L., and Christiansen, C. (1984). Longitudinal study of calcium metabolism in male puberty. II. Relationship between mineralization and serum testosterone. Acta. Paediatr. Scand. 73, 750–755. 87. Hyldstrup, L., McNair, P., Jensen, G. F., Nielsen, H. R., and Transbol, I. (1984). Bone mass as referent for urinary hydroxyproline excretion: age and sex-related changes in 125 normals and in primary hyperparathyroidism. Calcif. Tissue Int. 36, 639–644. 88. Szulc, P., Seeman, E., and Delmas, P. D. (2000). Biochemical measurements of bone turnover in children and adolescents. Osteoporos. Int. 11, 281–294. 89. Mora, S., Pitukcheewanont, P., Kaufman, F. R., Nelson, J. C., and Gilsanz, V. (1999). Biochemical markers of bone turnover and the volume and the density of bone in children at different stages of sexual development. J. Bone Miner. Res. 14, 1664–1671.

Chapter 34 Variability in the Measurement of Biochemical Markers of Bone Turnover 90. Mora, S., Prinster, C., Proverbio, M. C., Bellini, A., de Poli, S. C., Weber, G., Abbiati, G., and Chiumello, G. (1998). Urinary markers of bone turnover in healthy children and adolescents: age-related changes and effect of puberty. Calcif. Tissue Int. 63, 369–374. 91. van der Sluis, I. M., Hop, W. C., van Leeuwen, J. P., Pols, H. A., and de Muinck Keizer-Schrama, S. M. (2002). A cross-sectional study on biochemical parameters of bone turnover and vitamin d metabolites in healthy Dutch children and young adults. Horm. Res. 57, 170–179. 92. van Coeverden, S. C., Netelenbos, J. C., de Ridder, C. M., Roos, J. C., Popp-Snijders, C., and Delemarre-van de Waal, H. A. (2002). Bone metabolism markers and bone mass in healthy pubertal boys and girls. Clin. Endocrinol. (Oxf) 57, 107–116. 93. Levine, M. A. (2003). Biochemical markers of bone metabolism: application to understanding bone remodeling in children and adolescents. J. Pediatr. Endocrinol. Metab. 16 (Suppl 3), 661–672. 94. Marowska, J., Kobylinska, M., Lukaszkiewicz, J., Talajko, A., Rymkiewicz-Kluczynska, B., and Lorenc, R. S. (1996). Pyridinium crosslinks of collagen as a marker of bone resorption rates in children and adolescents: normal values and clinical application. Bone 19, 669–677. 95. Tobiume, H., Kanzaki, S., Hida, S., Ono, T., Moriwake, T., Yamauchi, S., Tanaka, H., and Seino, Y. (1997). Serum bone alkaline phosphatase isoenzyme levels in normal children and children with growth hormone (GH) deficiency: a potential marker for bone formation and response to GH therapy. J. Clin. Endocrinol. Metab. 82, 2056–2061. 96. Seibel, M. J., Woitge, H. W., Pecherstorfer, M., Karmatschek, M., Horn, E., Ludwig, H., Armbruster, F. P., and Ziegler, R. (1996). Serum immunoreactive bone sialoprotein as a new marker of bone turnover in metabolic and malignant bone disease. J. Clin. Endocrinol. Metab. 81, 3289–3294. 97. Eastell, R., Delmas, P. D., Hodgson, S. F., Eriksen, E. F., Mann, K. G., and Riggs, B. L. (1988). Bone formation rate in older normal women: concurrent assessment with bone histomorphometry, calcium kinetics, and biochemical markers. J. Clin. Endocrinol. Metab. 67, 741–748. 98. Kelly, P. J., Pocock, N. A., Sambrook, P. N., and Eisman, J. A. (1989). Age and menopause-related changes in indices of bone turnover. J. Clin. Endocrinol. Metab. 69, 1160–1165. 99. Hassager, C., Colwell, A., Assiri, A. M., Eastell, R., Russell, R. G., and Christiansen, C. (1992). Effect of menopause and hormone replacement therapy on urinary excretion of pyridinium cross-links: a longitudinal and cross-sectional study. Clin. Endocrinol. (Oxf) 37, 45–50. 100. Seibel, M. J., Woitge, H., Scheidt-Nave, C., Leidig-Bruckner, G., Duncan, A., Nicol, P., Ziegler, R., and Robins, S. P. (1994). Urinary hydroxypyridinium crosslinks of collagen in population-based screening for overt vertebral osteoporosis: results of a pilot study. J. Bone Miner. Res. 9, 1433–1440. 101. Gundberg, C. M., Hanning, R. M., Liu, Y. A., Zlotkin, S. H., Balfe, J. W., and Cole, D. E. (1987). Clearance of osteocalcin by peritoneal dialysis in children with end-stage renal disease. Pediatr. Res. 21, 296–300. 102. Gundberg, C. M., and Weinstein, R. S. (1986). Multiple immunoreactive forms of osteocalcin in uremic serum. J. Clin. Invest. 77, 1762–1767. 103. Woitge, H. W., Pecherstorfer, M., Li, Y., Keck, A. V., Horn, E., Ziegler, R., and Seibel, M. J. (1999). Novel serum markers of bone resorption: clinical assessment and comparison with established urinary indices. J. Bone Miner. Res. 14, 792–801. 104. Robins, S. (1990). Collagen markers in urine in human arthritis. In: (A. Maroudas, ed.) “Methods in Cartilage Research”, pp. 348–352. Academic Press, London. 105. Seibel, M. J. (1997). Clinical use of pyridinium crosslinks. In: (H. Schmidt-Gayk, ed.) “Calcium Regulating Hormones and Markers of Bone Metabolism: Measurement and Interpretation”, pp. 157–169. Clin. Lab. Publications, Heidelberg.

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106. Stepan, J. J., Lachmanova, J., Strakova, M., and Pacovsky, V. (1987). Serum osteocalcin, bone alkaline phosphatase isoenzyme and plasma tartrate resistant acid phosphatase in patients on chronic maintenance hemodialysis. Bone Miner. 3, 177–183. 107. Halleen, J. M., Alatalo, S. L., Janckila, A. J., Woitge, H. W., Seibel, M. J., and Vaananen, H. K. (2001). Serum tartrate-resistant acid phosphatase 5b is a specific and sensitive marker of bone resorption. Clin. Chem. 47, 597–600. 108. McLaren, A. M., Hordon, L. D., Bird, H. A., and Robins, S. P. (1992). Urinary excretion of pyridinium crosslinks of collagen in patients with osteoporosis and the effects of bone fracture. Ann. Rheum. Dis. 51, 648–651. 109. Ohishi, T., Takahashi, M., Kushida, K., Hoshino, H., Tsuchikawa, T., Naitoh, K., and Inoue, T. (1998). Changes of biochemical markers during fracture healing. Arch. Orthop. Trauma Surg. 118, 126–130. 110. Ingle, B. M., Hay, S. M., Bottjer, H. M., and Eastell, R. (1999). Changes in bone mass and bone turnover following distal forearm fracture. Osteoporos. Int. 10, 399–407. 111. Ingle, B. M., Hay, S. M., Bottjer, H. M., and Eastell, R. (1999). Changes in bone mass and bone turnover following ankle fracture. Osteoporos. Int. 10, 408–415. 112. Hannon, R., and Eastell, R. (2000). Preanalytical variability of biochemical markers of bone turnover. Osteoporos. Int. 11 (Suppl 6), S30–44. 113. Fleiss, J. L. (1986). The design and analysis of clinical experiments, Chapter 2. Wiley, New York. 114. Littell, R. C., Milliken, G. A., Stroup, W. W., and Wolfinger, R. D. (1996). SAS System for Mixed Models (N. C. Cary, ed.) SAS Institute Inc. 115. Bland, J. M., and Altman, D. G. (1996). Measurement error and correlation coefficients. BMJ 313, 41–42. 116. Gertz, B. J., Shao, P., Hanson, D. A., Quan, H., Harris, S. T., Genant, H. K., Chesnut, C. H., 3rd, and Eyre, D. R. (1994). Monitoring bone resorption in early postmenopausal women by an immunoassay for cross-linked collagen peptides in urine. J. Bone Miner. Res. 9, 135–142. 117. Rosen, C. J., Chesnut, C. H., 3rd, and Mallinak, N. J. (1997). The predictive value of biochemical markers of bone turnover for bone mineral density in early postmenopausal women treated with hormone replacement or calcium supplementation. J. Clin. Endocrinol. Metab. 82, 1904–1910. 118. Allison, D. B. (1993). Limitations of coefficient of variation as index of measurement reliability. Nutrition 9, 559–561. 119. Yao, L., and Sayre, J. W. (1994). Statistical concepts in the interpretation of serial bone densitometry. Invest. Radiol. 29, 928–932. 120. Nguyen, T. V., Meier, C., Center, J. R., Eisman, J., and Seibel, M. J. Bone turnover markers in elderly men: effects of within-subject variability (submitted). 121. Galton, F. (1886). Regression toward mediocrity in hereditary stature. J. Antropological Inst. Great Britain and Ireland 15, 246–263. 122. Nguyen, T. V., Pocock, N., and Eisman, J. A. (2000). Interpretation of bone mineral density measurement and its change. J. Clin. Densitom 3, 107–119. 123. Chapurlat, R. D., Blackwell, T., Bauer, D. C., and Cummings, S. R. (2001). Changes in biochemical markers of bone turnover in women treated with raloxifene: influence of regression to the mean. Osteoporos. Int. 12, 1006–1014. 124. Cummings, S. R., Palermo, L., Browner, W., Marcus, R., Wallace, R., Pearson, J., Blackwell, T., Eckert, S., and Black, D. (2000). Monitoring osteoporosis therapy with bone densitometry: misleading changes and regression to the mean. Fracture Intervention Trial Research Group. JAMA 283, 1318–1321. 125. Congdon, P. (2002). Bayesian statistical modelling. Wiley, New York. 126. Harris, E. K., and Shakarki, G. (1979). Use of the population distribution to improve estimation of individual means in epidemiological studies. J. Chronic. Dis. 32, 233–243.

582 127. Davis, C. E. (1976). The effect of regression to the mean in epidemiologic and clinical studies. Am. J. Epidemiol. 104, 493–498. 128. Healy, M. J., and Goldstein, H. (1978). Regression to the mean. Ann. Hum. Biol. 5, 277–280. 129. Yudkin, P. L., and Stratton, I. M. (1996). How to deal with regression to the mean in intervention studies. Lancet 347, 241–243. 130. Jensen, J. E., Sorensen, H. A., Kollerup, G., Jensen, L. B., and Sorensen, O. H. (1994). Biological variation of biochemical bone markers. Scand. J. Clin. Lab. Invest. Suppl. 219, 36–39. 131. Hannon, R., Blumsohn, A., Naylor, K., and Eastell, R. (1998). Response of biochemical markers of bone turnover to hormone replacement therapy: impact of biological variability. J. Bone Miner. Res. 13, 1124–1133. 132. Stevenson, H. P., Leslie, H., and Sheridan, B. (1997). Intra-individual variation in serum type I procollagen carboxy-terminal propeptide and type I collagen carboxy-terminal cross-linked telopeptide concentrations. Ann. Clin. Biochem. 34 ( Pt 3), 317–318. 133. Scariano, J. K., Garry, P. J., Montoya, G. D., Wilson, J. M., and Baumgartner, R. N. (2001). Critical differences in the serial

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measurement of three biochemical markers of bone turnover in the sera of pre- and postmenopausal women. Clin. Biochem. 34, 639–644. Alvarez, L., Ricos, C., Peris, P., GuaNabens, N., Monegal, A., Pons, F., and Ballesta, A. M. (2000). Components of biological variation of biochemical markers of bone turnover in Paget’s bone disease. Bone 26, 571–576. Ginty, F., Flynn, A., and Cashman, K. (1998). Inter and intra-individual variations in urinary excretion of pyridinium crosslinks of collagen in healthy young adults. Eur. J. Clin. Nutr. 52, 71–73. Eastell, R., Mallinak, N., Weiss, S., Ettinger, M., Pettinger, M., Cain, D., Fressland, K., and Chesnut, C., 3rd (2000). Biological variability of serum and urinary N-telopeptides of type I collagen in postmenopausal women. J. Bone Miner. Res. 15, 594–598. Borderie, D., Roux, C., Toussaint, B., Dougados, M., Ekindjian, O. G., and Cherruau, B. (2001). Variability in urinary excretion of bone resorption markers: limitations of a single determination in clinical practice. Clin. Biochem. 34, 571–577.

Chapter 35

Validation of Biochemical Markers of Bone Turnover Kim Brixen Erik Fink Eriksen

Department of Endocrinology, Odense University Hospital, DK-5000 Odense C, Denmark Novartis Pharma, Basel, Switzerland

IV. Conclusions References

I. Introduction II. Validation of Biochemical Markers by Calcium Kinetics III. Validation of Biochemical Markers by Bone Histomorphometry

I. INTRODUCTION

(4) and reflect remodeling activity in the entire skeleton. However, they also have disadvantages: (1) they do not provide information about the work of individual cells; (2) they do not clearly distinguish between bone matrix formation and mineralization; (3) they do not differentiate between cortical and trabecular bone turnover, and (4) their serum and urine levels may be affected by their rate of clearance and ambient hormonal milieu. In addition to technical demands on precision, accuracy, and specificity, biochemical markers should be validated by comparison with independent means of bone formation and resorption in terms of amount of work or rate of work and not merely by clinical experience. The combined calcium kinetics and balance studies are considered the golden standard since these methods measure bone turnover at the organ level (i.e. the whole skeleton). These methods, however, have some inherent limitations. First, they are very cumbersome and require admission of the patients to a metabolic ward for weeks. Second, bone formation and resorption rates are not mathematically independent. Third, the measurement error of the resorption rate is rather large [1]. Histomorphometric analysis of bone biopsies provides another possibility for validation;

Assessment of bone turnover (i.e. the amount of bone renewed during the bone remodeling process) remains essential in understanding bone physiology, pathophysiology, and skeletal response to treatment. Unfortunately, direct measurement of bone turnover in the whole skeleton is only possible by combined calcium kinetics and balance. Direct assessment of bone turnover and bone cell activity is also possible using histomorphometric measurements on bone biopsies, but only provides information on a small compartment of the skeleton. Unfortunately, both bone biopsies and the combined calcium kinetics and balance studies are impractical for routine clinical use. A long and growing list of biochemical markers of bone formation and resorption, that can be measured in the blood and urine, have been used or proposed to provide easy, albeit indirect, means of assessing bone turnover (Table I; see also chapters 34 and 35). Moreover, bone tissue concentrations of growth factors produced by bone cells have been proposed to reflect bone turnover. Biochemical markers of bone turnover have many advantages: (1) they are noninvasive; (2) inexpensive; (3) they can be repeated numerous times; Dynamics of Bone and Cartilage Metabolism

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KIM BRIXEN AND ERIK FINK ERIKSEN Table I.

Biochemical Markers of Bone Turnover

Markers of bone formation Serum osteocalcin Serum total alkaline phosphatase (TALP) Serum bone specific alkaline phosphatase (BALP) Serum C-terminal pro-peptide of type-I collagen (PICP) Serum N-terminal pro-peptide of type-I collagen (PINP) Markers of bone resorption Urinary hydroxyproline (OHP) Urinary N-terminal telopeptide of type-I collagen (NTX-I) Urinary pyridinoline (PYD) Urinary deoxypyridinoline (DPD) Serum C-terminal cross-linked telopeptide of type-I collagen (CTX-I) Serum N-terminal telopeptide of type-I collagen (NTX-I) Urinary helical peptide (HELP) Serum tartrate-resistant acid phosphatase (TRAcP)

however, it has another set of limitations: (1) the analysis only reflects turnover in the bone tissue actually sampled (usually the iliac crest); (2) turnover is much greater in cancellous than cortical bone, which comprises approximately 80% of the skeleton; (3) the invasive nature of the bone biopsy procedure usually precludes more than two repeated measurements in each patient; (4) bone resorption rates can only be assessed indirectly from dynamic bone formation data; (5) most traditional histomorphometric analyses report indices pertaining only to cancellous bone, and thus correlation analyses will exclude remodeling activity taking place in cortical bone. When using either calcium kinetics or bone histomorphometry for the validation of biochemical markers of bone turnover, the results should be interpreted in light of the tight biological coupling between bone formation and resorption in most clinical situations. Finally, the significant day-to-day variation for all markers in serum or urine studied so far represents a problem [2, 3]. This variability amounts to 15–30% and affects the ability to demonstrate significant correlations and limits detection of changes over time in individual patients. Some of this variation can be overcome by standardized and repeated sampling, but this has only been performed in few studies reviewed here.

II. VALIDATION OF BIOCHEMICAL MARKERS BY CALCIUM KINETICS A. Biochemical Markers of Bone Formation The ideal marker of bone formation should be a structural protein released into the blood in a rate proportional to its incorporation into bone and the fraction released from osteoblasts should be unaffected by disease, hormonal or other metabolic pertubations. It should have a wellcharacterized function and should not be released during

bone resorption. Also, its metabolic pathway and serum half-life should be known. Osteoblasts produce collagen, a number of noncollagenous matrix proteins, enzymes, and growth factors. Although none of these constituents meet all the ideal demands, several can be measured in serum as fairly reliable indicators of osteoblastic activity (Table I; see also part I of this volume, and chapters 34 and 35). 1. OSTEOCALCIN

In one study, serum levels of osteocalcin were compared with bone formation rates as measured by calcium kinetics in normal volunteers [4] (Fig. 1). A close, positive correlation was found between formation rate (Vo+) and serum calcium (R = 0.82) (Table II) [4]. A lesser correlation (R = 0.55) was found in a study on 15 girls with cystic fibrosis [5] and in healthy pubertal girls (R = 0.49) [6]. In the latter study, no significant correlation could be demonstrated before puberty [6]. No correlation was found in another study on 19 normal volunteers [7], however, these results may be due to the small number of subjects and a small range of turnover. A study comprising 12 young and 11 older women did not find any significant correlation between serum osteocalcin and mineralization rate when both groups were pooled. The correlation was significant, however, when the two groups were analyzed separately [8]. Finally, in patients with high and low turnover bone diseases (myxedema, thyrotoxicosis, and primary hyperparathyroidism) [7] as well as osteoporosis [9], significant correlations between mineralization rate and serum osteocalcin have been demonstrated. 2. ALKALINE PHOSPHATASE

A number of studies have demonstrated significant correlations between serum levels of bone-specific alkaline phosphatase (BALP) and bone formation as assessed by calcium kinetics in normal subjects [4], healthy pubertal girls [6], patients with high and low turnover states [10], renal failure complicated with adynamic bone disease, aluminum-related osteomalacia, secondary hyperparathyroidism [11], and renal failure treated with continuous ambulatory peritoneal dialysis [12, 13]. Some [10], but not all [4, 7], studies have found that serum BALP levels correlate more closely with mineralization rate than total alkaline phosphatase; however, this may be due to a small number of subjects and a small range of bone turnover in some studies [7]. 3. AMINO- AND CARBOXY-TERMINAL PROPEPTIDES OF TYPE-I PROCOLLAGEN

Serum carboxy-terminal propeptide of type-I procollagen (PICP) has been validated as a marker of bone formation by studies demonstrating significant correlations with mineralization rate, as measured by calcium kinetics in

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Figure 1 Relationship between bone formation rate (Vo+) determined by calcium kinetics and biochemical markers of bone formation in young healthy females; (A) serum osteocalcin (OC), (B) serum total alkaline phosphatase (AlkP), (C) serum bone alkaline phosphatase (BAP). Reproduced from J. Bone Miner. Res. 1997, 12: 1714–20, with permission from the American Society for Bone and Mineral Research.

normal volunteers [7], patients with osteoporosis [7, 14], and patients with high turnover bone diseases [7]. PICP did not, however, correlate with mineralization rate in patients with renal failure [15] or on peritoneal dialysis [12]. The serum levels of amino-terminal propeptide of type-I procollagen (PINP) has not been correlated to calcium kinetics. Pons et al. [16], however, reported a close correlation (R = 0.69, P = 0.001) between serum PINP and a quantitative evaluation of scintigraphic activity in patients with Paget’s disease. Serum PINP and PICP have emerged as excellent markers of new bone formation during therapy with parathyroid hormone (1-34). Of the two collagen markers, PINP is superior to PICP. Several observations support this notion: (1) PINP peak concentrations coincide with maximal boneforming acitivity in bone biopsies at 6 months, while PICP reaches its maximum after 1 month; (2) PINP has a much

wider dynamic range than PICP. Median increases in PINP after 6 months of PTH therapy are 250% of basal vs. 40% for PICP at 1 month [17]. 4. ASSESSMENT OF OSTEOBLASTIC SECRETION RATES FOR MARKERS OF BONE FORMATION

The serum level of a given bone marker depends on several factors: (1) the osteoblastic formation rate, (2) the number of osteoblasts currently involved in bone formation, and (3) the metabolic clearance rate of the marker (in most cases, this depends on filtration and tubular treatment in the kidneys). In order to distinguish the effects of osteoblastic secretion of formative markers from bone turnover, Charles et al. [7] calculated the “apparent” osteoblastic secretion rate of the markers osteocalcin, alkaline phosphatase, and PICP dividing their serum levels by mineralization rate

Table II. Correlation between Mineralization Rate (m) and Resorption Rate (r) as Determined by Calcium Kinetics and Biochemical Markers of Bone Formation and Resorption in Young Healthy Women

m r

S-OC

S-TALP

S-BALP

S-TRAcP

U-OHP/Cr

U-NTX/Cr

0.82 0.66

0.92 0.81

0.90 0.72

0.77 0.71

0.94 0.79

0.88 0.70

Reproduced from, J. Bone Miner. Res. 1997, 12:1714–20, with permission from the American Society for Bone and Mineral Research. Data are shown as Pearson correlation coefficients. S-OC, serum osteocalcin; S-TAP, serum total alkaline phosphatase; S-BALP, serum bone specific alkaline phosphatase; S-TRAP, serum tartrate resistant acid phosphatase; U-OHP/Cr, urinary hydroxyproline/creatinine ratio; U-NTX/Cr, urinary N-telopeptide of type I collagen/creatinine ratio.

586 as determined by calcium kinetics (Fig 2). Apparent osteoblastic secretion rate of PICP was significantly reduced in primary hyperparathyroidism, thyrotoxicosis, and osteomalacia but similar to that of normal controls in myxedema and osteoporosis. The apparent secretion of osteocalcin remained similar to that of normal controls in all diseases except osteomalacia, highlighting the crucial role of vitamin D in the regulation of osteocalcin secretion. Apparent secretion of alkaline phosphatase was similar to that of normals in primary hyperparathyroidism and thyrotoxicosis but significant elevation was seen in myxedema,

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osteoporosis, and osteomalacia. Apart from osteomalacia, osteocalcin revealed the least perturbations across the disease spectrum. This analysis depends on the assumption that the metabolic clearance rate for the markers is similar in all the studied diseases. Furthermore, it is assumed that the matrix production and the 45Ca incorporation into the matrix are unaltered by disease, which is hardly the case in osteomalacia. Even with these limitations, the study indicates that the serum levels of biochemical bone markers depend not only on bone turnover, but also on ambient concentrations of calciotropic hormones and possible disease-specific alterations in osteoblastic function or clearance of the marker.

B. Biochemical Markers of Bone Resorption The ideal bone resorption marker should be a degradation product of a matrix component not found in any other tissue outside bone. Its serum or urine levels should not be under separate endocrine control and it should not be reutilized in new bone formation. Moreover, its clearance rates should stay constant over a wide range of metabolic bone diseases. During the degradation of collagen, several metabolites are released into circulation and excreted in the urine (see Chapter 35). Some of these are specific for bone or predominantly originate from bone. Furthermore, osteoclasts produce several enzymes among which tartrateresistant acid phosphatase may be used as a marker of bone resorption, albeit with some reservations. Recently, new immunoassays specific for bone tartrate-resistant acid phosphatase type 5b have been developed [18] (see also chapters 10 and 35). The close metabolic control of calcium absorption, excretion, and non-resorptive release from bone, as well as reutilization during bone formation precludes the use of serum levels or urinary excretion rates of calcium as a measure of bone resorption. 1. HYDROXYPROLINE

Figure 2 “Apparent” osteoblastic secretion rate of osteocalcin PICP, osteocalcin (OC), and alkaline phosphatase (AP) in normal controls and patients with primary hyperparathyroidism (PHP), thyrotoxicosis (TT), myxedema (Myx), osteoporosis (OP), and osteomalacia (Osmal). From Charles et al. Assessment of Bone Formation by Biochemical Markers in Metabolic Bone Disease (c) 1992 Springer. Calcif. Tissues Int. 1992, 51: 406–11, with permission.

The urinary hydroxyproline/creatinine ratio correlates with bone resorption, as measured by calcium kinetics in normal healthy volunteers (Fig. 3, Table II) [4] and in patients with high and low turnover bone diseases [19]. In one study on 37 women with osteoporosis the urinary hydroxyproline/creatinine ratio correlated with bone resorption as measured by 85Sr-kinetics [20]. In contrast, no significant correlation could be demonstrated in a similar but smaller study comprising 19 patients [21]. 2. COLLAGEN CROSS-LINK PEPTIDES

In recent years, a host of new, very specific collagen crosslink-related markers of bone resorption has emerged.

Chapter 35 Validation of Biochemical Markers of Bone Turnover

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Figure 3 Relationship between bone resorption rate (Vo-) determined by calcium kinetics and biochemical markers of bone resorption in young healthy females; (A) serum tartrate resistant acid phosphatase (TRAP), (B) urinary hydroxyproline/creatinine ratio (OHP/Cr), and (C) urinary telopeptide of type-I collagen/creatinine ratio (NTx/Cr). Reproduced from J. Bone Miner. Res. 1997, 12: 1714–20, with permission from the American Society for Bone and Mineral Research.

All are based on urine and more lately serum assays for type I collagen crosslink peptides (see also Chapter 35). The urinary excretion of the amino-terminal telopeptide of type I collagen (NTX-I) correlates with bone resorption as assessed by calcium kinetics in healthy volunteers [4] (Fig. 3, Table II). Similarly, urinary excretion of pyridinoline (PYD) and deoxy-pyridinoline (DPD) correlate with bone resorption in patients with osteoporosis [21] and perform better than urinary hydroxyproline. In this study, the strongest correlation was observed between DPD and calcium kinetic resorption rate with complete correction for long-term exchange (R = 0.71, P < 0.001). Also, during hormone replacement therapy (HRT) or treatment with parathyroid hormone peptide (1-38) combined with HRT, similar percent changes in biochemical markers and estimates of bone resorption were demonstrable. The newer crosslink peptides (serum NTX-I, CTX-I, and helical peptide) have not been compared to calcium kinetics. However, they have been correlated to disease activity in Paget’s disease and multiple myeloma. Urinary NTX-I correlated to disease activity in Paget’s disease (R = 0.63, P < 0.005) as assessed by quantitative scintigraphy [16]. Moreover, serum CTX-I has recently been reported to be an excellent marker of bone resorption and overall disease activity in multiple myeloma based on radiological features [22].

Resorption markers seem to underestimate the degree of cancellous bone turnover suppression elicited by antiresorptive therapy. Hannon et al. [18] analyzed histomorphometric indices pertaining to bone turnover and compared them to markers of bone resorption in 30 women with osteoporosis treated with the bisphosphonate, risedronate, for 3 years. The resorptive markers U-NTX-I, U-CTX-I exhibited 10–20% less suppression than histomorphometric indices like mineralization surface (MS/BS), volume referent resorption rate (BRs/BV), volume referent formation rate (BFR/BV), and activation frequency (Ac.f). However, only the decrease in MS/BS was statistically significantly different from the marker levels (P < 0.02–0.07). The decrease in free DPD was significantly less than that seen in Ac.f, BFR/BV, and BRs/BV (P < 0.0002–0.05). Another telopeptide, carboxy-terminal cross-linked telo-peptide of type I collagen (ICTP) can be measured in serum. Serum ICTP has been shown to correlate with bone resorption rate as assessed by calcium kinetics in patients with high and low turnover bone disease and osteoporosis, but not in normals [14]. In contrast, however, serum ICTP levels are poorly correlated to disease activity in Paget’s disease. Recent studies have shed some light on this discrepancy. These studies indicate that the release of ICTP versus crosslink peptides like CTX-I depend on different

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collagenolytic pathways. Release of CTX-I from bone collagen during bone resorption depends on cathepsin K activity, which generates the CTX-I epitope but destroys the epitope specific for ICTP. Conversely, ICTP release depends mainly on metalloproteinase (type 2, 9, 13, and 15) activity [23]. 3. TARTRATE-RESISTANT ACID PHOSPHATASE

Serum tartrate-resistant acid phosphatase (TRAcP) correlates with bone resorption as assessed by calcium kinetics in normal healthy volunteers (Fig. 3, Table II) [4]. Ballanti et al. [24] investigated 24 patients with various metabolic bone disease and found that S-TRACP correlated to osteoclast surface, osteoclast number, and number of TRAcPpositive osteoclasts in bone sections. Newer immunoassays are able to specifically measure osteoclastic TRAcP, also called TRAcP 5b. Serum levels of TRAcP 5b in serum are less variable, but the response is not as dynamic as that seen for cross-link peptides [18] (see also Chapter 34). No studies on correspondence with histomorphometry or calcium kinetics have been published for TRAcP 5b. 4. ASSESSMENT OF OSTEOCLASTIC SECRETION RATES FOR MARKERS OF BONE RESORPTION

In patients with various metabolic bone diseases (thyreotoxicosis, myxedema, primary hyperparathyroidism, and osteoporosis) we tried to normalize resorption markers (serum ICTP, urinary PYD, and urinary DPD) to whole body calcium kinetic resorption rates (r) (Eriksen et al. unpublished data). The normalization was performed by creating the fractions ICTP/r, PYD/r and DPD/r (Fig. 4). When compared to normal controls, patients with primary hyperparathyroidism generally displayed reduced fractions (i.e. lower rates of release of crosslinks per unit bone resorbed), while patients with myxedema and osteoporosis generally exhibited increased fractions (i.e. increased release of cross-links per unit bone resorbed). Patients with thyrotoxicosis had fractions no different from those of normal controls. The differences from normal controls, however, were only significant in patients with primary hyperparathyroidism and osteoporosis (serum ICTP/r) and for myxedema and osteoporosis (urinary PYD/r). Thus, just like markers of bone formation, resorption markers also show variations in secretion rates in different disease states.

III. VALIDATION OF BIOCHEMICAL MARKERS BY BONE HISTOMORPHOMETRY Bone histomorphometry offers the most detailed analysis of bone remodeling, but the interpretation of

Figure 4 Variation between different metabolic bone diseases for three different resorption markers corrected for whole skeletal resorption rate (r) as assessed by calcium kinetics: serum pyridinoline crosslinked telopeptide domain of type I collagen (ICTP/r), urinary pyridinoline crosslink peptide (PYD/r) and urinary doxypyridinoline (DPYR/r). Parentheses above bars denote 95% confidence intervals; *P<0.05; ***P<0.001.

histomorphometric findings in relation to biochemical markers is hampered by the fact that the biopsy core only reflects bone remodeling in a small area of the total skeleton. Moreover, the relative contribution of cortical, endocortical, and cancellous envelopes to total remodeling activity is not known in detail. Despite these limitations, histomorphometric analysis is the only way to assess the relationship between bone turnover at tissue level, single cell activity, and mineralization defects on the one hand and biochemical bone markers on the other hand. The production rates of most bone markers mainly depend on the number of currently active resorptive or

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formative cells. This number depends on two factors: (1) activation frequency (i.e. initiation rate of new remodeling cycles) and (2) single-cell activity at individual remodeling sites. Bone histomorphometry is the only technique that allows separation of these two variables. Most studies comparing histomorphometry and biochemical markers have mainly used surface estimates and indices reflecting osteoblastic and osteoclastic activity. Osteoblastic, osteoid, mineralization, or resorption surfaces, however, do not separate between single-cell activity and activation frequency. Very low turnover, for example, with low cellular activity and prolongation of all phases of remodeling (and low production rates) may in theory lead to increases in surface extension similar to those seen in high turnover states with increased cellular activity (and production rates) [25]. It is, therefore, preferable to compare volume referent, dynamic histomorphometric indices (e.g., by combining tetracycline-derived indices and surface estimates) to avoid these confounders. Standard formulae permit the calculation of bone resorption and formation based on the use of the surface to volume ratio.

A. Assessment of Bone Turnover in Normal Individuals Eastell et al. [26] assessed bone formation rates using multiple methods in 12 younger (aged 30–41 years) and 11 older (aged 55–73 years) healthy women. Bone formation was higher in the older than in the younger based on measurement of osteocalcin (1.67 ± 0.07 [SE] versus 1.14 ± 0.10 nmol/L, P < 0.01), serum bone-specific alkaline phosphatase activity (388 ± 42 versus 223 ± 22 U/l, p < 0.01) and bone formation rate by histomorphometry on iliac crest biopsies (31.1 ± 4.9% versus 15.1 ± 2.7%/year, P < 0.01). Bone formation, however, was similar in the two groups when accretion rates were assessed by calcium kinetics (5.9 ± 1.0 versus 7.5 ± 1.2, NS). This discrepancy was ascribed to several age-related factors; reduced mineralization of completed osteons and lack of correction for the decrease in total skeletal calcium in the older group. Younger women exhibited significant correlations between osteocalcin and volume referent formation rates. No such correlation was demonstrable for bone-specific alkaline phosphatase [26]. In older women, neither osteocalcin nor bone-specific alkaline phosphates correlated to histomorphometric formation rates. In a study comprising 43 healthy men aged 20–80 years, serum osteocalcin correlated significantly with volume referent bone formation rate (R = 0.43, P = 0.004) [27].

B. Assessment of Bone Turnover in Patients with Various Metabolic Bone Diseases 1. OSTEOPOROSIS

Delmas et al. [28] compared the urinary excretion of pyridinoline crosslinks (PYD and DPD) along with serum osteocalcin and urinary hydroxyproline in 36 elderly women with vertebral osteoporosis. The same women also had an iliac crest biopsy performed. The authors found that urinary pyridinoline crosslinks, but not hydroxyproline, correlated significantly with histological resorption, assessed by osteoclast surface (R = 0.35, P < 0.05 for PYD; R = 0.46, P < 0.01 for DPD). In addition, PYD and DPD were correlated with bone formation rate as well as serum osteocalcin, with correlation coefficients ranging from 0.69 to 0.80 (P < 0.0001). These data indicate that PYD and DPD are sensitive markers of bone turnover in elderly women with vertebral osteoporosis. The authors concluded that the poor correlation between markers of bone resorption and histological assessment of bone resorption indicated the low sensitivity of iliac crest histomorphometry in the measurement of resorption rate of the skeleton. As mentioned above a surface estimate is a poor index for comparison with volume-dependent variables like biochemical markers of bone remodeling because it depends on both activation frequency and cellular activity at individual resorption foci. Indices containing more dynamic aspects of bone resorption might yield better estimates. Brown et al. [29] investigated the correlation between osteocalcin levels and bone remodeling in osteoporosis. Mean serum osteocalcin was normal (7.0 ± 3.3 ng/ml [mean ± SD]) in 26 patients with untreated postmenopausal osteoporosis. Nine patients, however, exhibited osteocalcin values either above (n = 4) or below (n = 5) the normal values obtained in 35 age-matched control women (6.9 ± 1.3 ng/ml). Serum osteocalcin correlated positively with osteoid volume, osteoid surfaces, mineralizing surfaces, and bone formation rate but not with erosion surfaces, linking this marker primarily to matrix mineralization. Based on normal values for osteoid volume, the authors classified patients as having either high (HF, nine patients), normal (NF, 12 patients) and low osteoid formation (LF, five patients). Serum osteocalcin (mean ± SEM) was significantly lower in LF group (2.7 ± 0.9 ng/ml) and significantly higher in HF group (9.7 ± 0.8 ng/ml) than in the NF group (7.0 ± 0.6 ng/ml). Somewhat surprisingly, serum alkaline phosphatase and urinary hydroxyproline did not discriminate between these three groups and did not correlate significantly with any of the histomorphometric indices. The authors concluded that serum osteocalcin could predict the histological profile in postmenopausal

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osteoporosis and suggested that the marker might be useful both in the diagnosis and control of treatment effects in patients with osteoporosis. The use of osteocalcin in this area is, however, hampered by its sensitivity to ambient vitamin D levels and the resulting degree of secondary hyperparathyroidism in individual patients. 2. RENAL OSTEODYSTROPHY

Couttenye et al. [30] investigated the correlation between bone alkaline phosphatase and adynamic bone disease in 103 hemodialysis patients. They performed a bone biopsy after double tetracycline labeling and correlated the histomorphometric indices of bone turnover to serum levels of PTH, osteocalcin, and the bone isoenzyme of alkaline phosphatase. In 38 (37%) of the patients the diagnosis of adynamic bone disease was established by bone histomorphometry. The authors established cut off values being ≤27 U/L for the bone iso-enzyme of alkaline phosphatase, ≤14 µg/L for osteocalcin and ≤150 pg/ml for PTH. Analysis of receiver operator curves revealed that bone alkaline phosphatase and PTH to be the best indices for the detection of adynamic bone disease, as indicated by a sensitivity of 78.1 and 80.6% and a specificity of 86.4 and 76.2% respectively. Furthermore, the positive predictive values were calculated for the proposed cut-off values at 75% for bone alkaline phosphatase, 65% for intact PTH, and 55% for osteocalcin. The authors, therefore, concluded that bone alkaline phosphatase was the best marker of adynamic bone disease. Urena et al. [31] investigated 42 hemodialysis patients with bone histomorphometry study and assessment of bone alkaline phosphatase, total alkaline phosphatase, and PTH. They found that patients with high turnover bone disease exhibited significantly higher plasma bone alkaline phosphatase (BALP) levels than patients with normal or low bone turnover (66.9 ± 63.5 ng/ml versus 10.8 ± 4.2 ng/ml). BALP levels were positively correlated with osteoclast surface (R = 0.39, P < 0.0001), osteoclast number/mm2 (R = 0.36, P < 0.001), osteoblast surface (R = 0.50, P < 0.005), and bone formation rate (R = 0.91, P < 0.0001). The bone formation rate revealed a better correlation with plasma BALP levels than total alkaline phosphatase, or PTH. Plasma BALP levels >20 ng/mL showed the highest sensitivity and specificity for the diagnosis of high turnover bone disease, and formally excluded patients with normal or low turnover bone disease. The authors consequently concluded that both serum PTH and BALP were useful in the separation of high and low turnover renal osteodystrophy. Joffe et al. [32] tried to assess whether bone histology could be predicted by clinical assessment and noninvasive

techniques in patients on peritoneal dialysis. No correlations were found between clinical symptoms, biochemical markers, radiographic bone surveys, BMC measurements, and bone histology. The authors, therefore, concluded that bone histological changes in renal osteodystrophy are not reliably predicted by bone markers. Charhon et al. [33] correlated serum osteocalcin, serum alkaline phosphatase, and PTH to dynamic histomorphometry in 42 patients undergoing chronic hemodialysis. Serum osteocalcin was markedly increased compared with normal subjects (64.0 ± 74.8 [mean ± SD] vs. 6.2 ± 2.2 ng/ml). Furthermore, osteocalcin significantly correlated with serum levels of alkaline phosphatase (R = 0.53) and PTH (r = 0.55). Serum osteocalcin was significantly higher in the 14 patients with high turnover renal osteodystrophy (138.5 ± 90.8 ng/ml) than in the 28 patients with low turnover (26.8 ± 14.8 ng/ml). Serum osteocalcin was significantly correlated with the cellular parameters of bone resorption and formation (R = 0.57–0.69) and with the dynamic parameters of bone formation (R = 0.62–0.82). The extent of stainable bone aluminum was significantly negatively correlated with serum osteocalcin (R = −0.51) and PTH (R = −0.33), but not with serum alkaline phosphatase. These authors concluded that measurement of serum osteocalcin allowed better discrimination between low and high turnover renal osteodystrophy than serum alkaline phosphatase. In patients with low turnover disease, however, serum osteocalcin did not permit discrimination between patients with or without osteomalacia. Mazzafero et al. [15] investigated whether markers of bone collagen formation (serum PICP) and degradation (serum ICTP) could provide indirect estimates of the rate of bone turnover. They also measured serum intact and C-terminal PTH, osteocalcin, and alkaline phosphatase. A transiliac bone biopsy for histomorphometry was also performed. As shown in other studies, serum PTH, osteocalcin, and alkaline phosphatase were correlated to static histomorphometric parameters of resorption and formation. Seurm PICP showed a slightly positive correlation with osteocalcin (R = 0.365, P < 0.01), but not with PTH, AP, or histomorphometric indices of bone resorption and formation. 3. OTHER METABOLIC BONE DISEASES

Lauffenburger et al. [34] investigated two groups of patients: (1) low turnover bone disease (osteoporosis) before and after fluoride treatment; (2) high turnover bone disease (Paget’s disease) before and after bisphosphonate treatment with etidronate. The authors assessed the following histomorphometric indices: osteoblast surface (ObS/BS), osteoid surface (OS/BS), osteoclast number (N.ocl/BS),

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Chapter 35 Validation of Biochemical Markers of Bone Turnover

and erosion surface (ES/BS). The histomorphometric parameters were correlated to serum alkaline phosphatase, urinary hydroxyproline, calcium kinetic accretion rate (m), and calcium kinetic resorption rate (r). In both patient groups, bone formation indices were significantly correlated: ObS/BS/m, R = 0.85; OS/BS/m, R = 0.83; and alkaline phosphatase/m, R = 0.97. Bone resorption indices correlated less well than bone formation indices: Oc.N/BS/r, R = 0.68 and ES/BS/r, R = 0.63. Urinary hydroxyproline correlated highly significantly to the sum of r + m (R = 0.97). Eriksen et al. [35] measured serum levels of ICTP as a marker of bone resorption and serum PICP as a marker of bone formation. Serum levels of the two markers were correlated to histomorphometric indices of bone resorption and bone formation, calculated from iliac crest bone biopsies in a group of 18 individuals with high and low turnover bone disease (myxedema, primary hyperparathyroidism, and thyrotoxicosis). Mean serum ICTP and PICP values differed widely between the three groups. Mean serum PICP values in thyrotoxicosis and primary hyperparathyroidism were three and two times higher, respectively, than values obtained in patients with myxedema. Serum ICTP values revealed even more pronounced differences between the three groups. Matrix degradation assessed from serum ICTP was three times higher than the degradation rate seen in primary hyperparathyroidism and four times that seen in myxedema. The differences in bone formation rates as assessed by bone histomorphometry (BFR/BV) revealed relative differences similar to those seen for PICP. This was not the case, however, for serum ICTP. Patients with myxedema revealed the lowest values for volume referent resorption rate (BRs.R/BV), but cancellous bone resorption rates in thyrotoxicosis was lower than that seen in primary hyperparathyroidism, despite the pronounced differences in ICTP between the two groups. Linear regression of serum ICTP on BRs.R/BV was significant (R = 0.51, P < 0.03). Serum ICTP also correlated significantly to cancellous bone balance (R = 0.45, P < 0.03). No significant regressions on other histomorphometric resorption indices were demonstrable. Serum PICP was significantly correlated to the mineral appositional rate (R = 0.53, P < 0.05) and volume-referent bone formation rate (R = 0.61, P < 0.01). The correlation to bone turnover as expressed in the activation frequency was also highly significant (R = 0.61, P < 0.01). No significant correlations with wall thickness or bone balance were demonstrable. When investigating the relative contribution of PICP and ICTP to cancellous bone balance as assessed by bone histomorphometry revealed that the variation in ICTP was the only significant determinant.

4. HYPERCALCEMIC STATES

Delmas et al. [36] investigated the use of serum osteocalcin in the discrimination between hypercalcemia due to malignant disease and primary hyperparathyroidism. They measured serum osteocalcin in 25 patients with primary hyperparathyroidism and in 24 patients with bone metastases with or without hypercalcemia. Despite similar levels of hypercalcemia, serum osteocalcin was increased in primary hyperparathyroidism (14.2 ± 9.6 ng/ml, P < 0.001), while it was decreased in malignant hypercalcemia (3.1 ± 2.8 ng/ml, P < 0.001), and normal in patients with bone metastases without hypercalcemia (6.6 ± 2.7 ng/ml). In primary hyperparathyroidism, serum osteocalcin was correlated with serum PTH (R = 0.90), calcium (R = 0.73), and adenoma weight (R = 0.79). After parathyroidectomy, serum osteocalcin slowly returned to normal values within 2–6 months, suggesting that serum osteocalcin reflects increased bone turnover rather than a direct effect of PTH on osteocalcin synthesis. Iliac crest biopsies were performed in 11 patients with primary hyperparathyroidism and in nine cancer patients in nonaffected areas. Serum osteocalcin was significantly correlated with all parameters reflecting bone formation but not with bone resorption. Patients with bone metastases were analyzed according to the presence or the absence of hypercalcemia. In contrast to normocalcemic patients who had normal serum osteocalcin, hypercalcemic patients exhibited reduced osteocalcin levels and a lower bone formation at the cellular level (P < 0.05). 5. BONE MARKER PROFILES DURING ANTIRESORPTIVE AND ANABOLIC THERAPIES

The introduction of anabolic, bone-forming therapies like parathyroid hormone (1-34) (PTH(1-34), teriparatide) has given us new insights into how well bone turnover markers reflect osteoclastic and osteoblastic responses to different treatment modalities for osteoporosis (see also Chapter 38). After initiation of antiresorptive therapy resorption and formation markers decrease in parallel to levels around 50–90% below baseline (see also Chapter 39). The pattern is completely different during treatment with PTH(1-34). Formation markers undergo rapid and pronounced increases to 250% over baseline, while resorption markers show a delayed and less-pronounced increase to about 50% over baseline. These differences are in keeping with the demonstrable formation of new bone during teriparatide therapy [37, 38], and the preservation of bone demonstrated during bisphosphonate therapy [39]. After antiresorptive therapy bone markers seem to stabilize after 6 months. During PTH therapy, both resorptive and formative markers

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Table III. Correlation between 1 month Serum Levels of Bone Specific Alkaline Phosphatase and C Terminal Propeptide of Type I Procollagen (PICP) and Two-dimensional (2D) and Three-dimensional (3D) Indices Pertaining to Bone Structure Assessed in Bone Biopsies after 22 months of Treatment with Teriparatide (PTH(1-34)) BALP

2D-BV/TV 2D-W.Th 2D-Ma.St.V 3D BV/TV

PICP

r

p

N

r

p

N

0.58 0.73 −0.51 0.54

<0.05 <0.001 <0.05 <0.05

16 17 15 19

0.51 0.60 −0.37 0.43

<0.05 <0.01 NS NS

16 17 15 19

BV/TV: cancellous bone volume; W.Th: mean wall thickness; Ma.St.V: Marrow star volume. From Dobnig et al. 2005.

peak at 6 months, followed by a return to baseline over the next 2.5 years [40]. Recent studies have demonstrated that short-term (3–12 month) changes in resorption and formation markers (osteocalcin, CTX-I, and NTX-I) correlate to bone mass increases during antiresorptive therapy with r-coefficients ranging between –0.45 to –0.78 (P = 0.001) [41]. In this and another analysis by Rosen et al. [42] CTX-I emerged as the marker with the best sensitivity and specificity profile and lowest variability. The BMD responses of individuals with significant changes in CTX-I, however, were highly variable. The experience with bone markers for assessing effects of bone forming agents is more limited, but recent studies have demonstrated that the early changes in formation markers correlate to both increases in BMD and improvements in bone structure. Early (1–3 month) changes in bone turnover markers (BALP, osteocalcin, PICP, and PINP) correlate to BMD at endpoint. Moreover, the higher the baseline turnover, the better the BMD response [17]. Dobnig et al. [43] reported that 1 and 3 month increases in formative markers like PICP and BALP correlated to improvements in bone structure after teriparatide therapy as assessed by 2D and µCT based 3D analyses (Table III). Increases in BALP at 1 month were significantly positively correlated with changes in 2D trabecular bone volume (R = 0.58, P < 0.05), 2D mean wall thickness (R = 0.73, P = 0.001), 3D trabecular bone volume (R = 0.54, P < 0.05), and 3D trabecular thickness (R = 0.49, P < 0.05). The increase also exhibited a significant negative correlation to trabecular bone connectivity as assessed by marrow star volume.

IV. CONCLUSIONS Serum and urine markers of bone formation and resorption are valuable tools for the investigation of metabolic

bone diseases and evaluation of treatment effects in clinical trials. Numerous histomorphometric and calcium kinetic studies have demonstrated that biochemical bone markers reflect skeletal bone formation and resorption activity in groups of individuals. The studies have, however, also revealed that the serum and urine concentrations of a given bone marker has to be interpreted in the context of numerous other factors (e.g. disease state, hormone levels, metabolic perturbations) influencing the formation and clearance rate of that particular marker (see also Chapters 36, 41, 42).

References 1. Charles, P., Eriksen, E. F., Mosekilde, L., Melsen, F., and Jensen, F. T. (1987). Bone turnover and balance evaluated by a combined calcium balance and 47calcium kinetic study and dynamic histomorphometry. Metabolism 36, 1118–1124. 2. Blumsohn, A., and Eastell, R. (1992). Prediction of bone loss in postmenopausal women. Eur. J. Clin. Invest. 22, 764–766. 3. Takahashi, M., Kawana, K., and Nagano A. (2002). Biological variability of biochemical markers of bone turnover in healthy women. Endocr. Res. 28, 257–264. 4. Weaver, C. M., Peacock, M., Martin, B. R., McCabe, G. P., Zhao, J., Smith, D. L., and Wastney, M. E. (1997). Quantification of biochemical markers of bone turnover by kinetic measures of bone formation and resorption in young healthy females. J. Bone Miner. Res. 12, 1714–1720. 5. Schulze, K. J., O’Brien, K. O., Germain-Lee, E. L, Booth, S. L., Leonard, A., and Rosenstein, B. J. (2004). Calcium kinetics are altered in clinically stable girls with cystic fibrosis Endogenous fecal losses of calcium compromise calcium balance in pancreatic-insufficient girls with cystic fibrosis. J. Clin. Endocrinol. Metab 89, 3385–3391. 6. Abrams, S. A., Copeland, K. C., Gunn, S. K., Gundberg, C. M., Klein, K. O., and Ellis, K. J. (2000). Calcium absorption, bone mass accumulation, and kinetics increase during early pubertal development in girls. J. Clin. Endocrinol. Metab 85, 1805–1809. 7. Charles, P., Hasling, C., Risteli, L., Risteli, J., Mosekilde, L., and Eriksen, E. F. (1992). Assessment of bone formation by biochemical markers in metabolic bone disease: separation between osteoblastic activity at the cell and tissue level. Calcif. Tissue Int. 51, 406–411. 8. Eastell, R., Delmas, P. D., Hodgson, S. F., Eriksen, E. F., Mann, K. G., and Riggs, B. L. (1988). Bone formation rate in older normal

Chapter 35 Validation of Biochemical Markers of Bone Turnover

9.

10.

11.

12.

13.

14.

15.

16.

17.

18.

19.

20.

21.

22.

23.

women: concurrent assessment with bone histomorphometry, calcium kinetics, and biochemical markers. J. Clin. Endocrinol. Metab. 67, 741–748. Wand, J. S., Green, J. R., Hesp, R., Bradbeer, J. N., Sambrook, P. N., Smith, T., Hampton, L., Zanelli, J. M., and Reeve, J. (1992). Bone remodelling does not decline after menopause in vertebral fracture osteoporosis. Bone Miner. 17, 361–375. Brixen, K., Nielsen, H. K., Eriksen, E. F., Charles, P., and Mosekilde, L. (1989). Efficacy of wheat germ lectin-precipitated alkaline phosphatase in serum as an estimator of bone mineralization rate: comparison to serum total alkaline phosphatase and serum bone Gla-protein [see comments]. Calcif. Tissue Int. 44, 93–98. Cochran, M., Neville, A., and Marshall, E. A. (1994). Comparison of bone formation rates measured by radiocalcium kinetics and doubletetracycline labeling in maintenance dialysis patients. Calcif. Tissue Int. 54, 392–398. Joffe, P., Hyldstrup, L., Heaf, J. G., Podenphant, J., and Henriksen, J. H. (1992). Bisphosphonate kinetics in patients undergoing continuous ambulatory peritoneal dialysis: relations to dynamic bone histomorphometry, osteocalcin and parathyroid hormone. Am. J. Nephrol. 12, 419–424. Jarava, C., Armas, J. R., Salgueira, M., and Palma, A. (1996). Bone alkaline phosphatase isoenzyme in renal osteodystrophy. Nephrol. Dial. Transplant. 11 Suppl 3, 43–46. Charles, P., Mosekilde, L., Risteli, L., Risteli, J., and Eriksen, E. F. (1994). Assessment of bone remodeling using biochemical indicators of type I collagen synthesis and degradation: relation to calcium kinetics. Bone Miner. 24, 81–94. Mazzaferro, S., Pasquali, M., Ballanti, P., Bonucci, E., Costantini, S., Chicca, S., De, M. S., Perruzza, I., Sardella, D., and Taggi, F. (1995). Diagnostic value of serum peptides of collagen synthesis and degradation in dialysis renal osteodystrophy. Nephrol. Dial. Transplant. 10, 52–58. Pons, F., Alvarez, L., Peris, P., Guanabens, N., Vidal-Sicart, S., Monegal, A., Pavia, J., Ballesta, A. M., Munos-Gomez, J., and Herranz, R. (1999). Quantitative evaluation of bone scintigraphy in the assessment of Paget’s disease activity. Nucl. Med. Commun. 20, 525–528. McClung, M. R., Miller, P. D., Civitelli, R., Warren, M. L., Greenspan, S. L., Haddock, L., Tamayo, J. A., Donley, D. W., and San Martin, J. (2003). Differential effects of teriparatide and alendronate on markers of bone remodeling and areal and volumetric bone density in women with osteoporosis. J. Bone Miner. Res. 18, S40. Hannon, R. A., Clowes, J. A., Eagleton, A. C., Al Hadari, A., Eastell, R., and Blumsohn, A. (2004). Clinical performance of immunoreactive tartrate-resistant acid phosphatase isoform 5b as a marker of bone resorption. Bone 34, 187–194. Charles, P., Poser, J. W., Mosekilde, L., and Jensen, F. T. (1985). Estimation of bone turnover evaluated by 47Ca-kinetics. Efficiency of serum bone gamma-carboxyglutamic acid-containing protein, serum alkaline phosphatase, and urinary hydroxyproline excretion. J. Clin. Invest. 76, 2254–2258. Deacon, A. C., Hulme, P., Hesp, R., Green, J. R., Tellez, M., Reeve, J., Hesp, R., Deacon, A. C., Hulme, P., and Reeve, J. (1987). Estimation of whole body bone resorption rate: a comparison of urinary total hydroxyproline excretion with two radioisotopic tracer methods in osteoporosis: Trends in trabecular and cortical bone in the radius compared with whole body calcium balance in osteoporosis. Clin. Chim. Acta 166, 297–306. Eastell, R., Colwell, A., Hampton, L., and Reeve, J. (1997). Biochemical markers of bone resorption compared with estimates of bone resorption from radiotracer kinetic studies in osteoporosis. J. Bone Miner. Res. 12, 59–65. Houze, P., Bellik, B., Brouet, J. C., Bouro, F., and Bousquet, B. (2000). Comparison of urinary markers for bone resorption in multiple myeloma. Clin. Chim. Acta 300, 181–193. Garnero, P., Ferreras, M., Karsdal, M. A., Nicamhlaoibh, R., Risteli, J., Borel, O., Qvist, P., Delmas, P. D., Foged, N. T., Delaisse, J. M. (2003).

24.

25.

26.

27.

28.

29.

30.

31.

32.

33.

34.

35.

36.

37.

38.

593 The type I collagen fragments ICTP and CTX reveal distinct enzymatic pathways of bone collagen degradation. J. Bone Miner. Res. 18, 859–867. Ballanti, P., Minisola, S., Pacitti, M. T., Scarnecchia, L., Rosso, R., Mazzuoli, G. F., and Bonucci, E. (1997). Tartrate-resistant acid phosphate activity as osteoclastic marker: sensitivity of cytochemical assessment and serum assay in comparison with standardized osteoclast histomorphometry. Osteoporos. Int. 7, 39–43. Eriksen, E. F. (1986). Normal and pathological remodeling of human trabecular bone: three dimensional reconstruction of the remodeling sequence in normals and in metabolic bone disease. Endocr. Rev. 7, 379–408. Eastell, R., Delmas, P. D., Hodgson, S. F., Eriksen, E. F, Mann, K. G., and Riggs, B. L. (1988). Bone formation rate in older normal women: concurrent assessment with bone histomorphometry, calcium kinetics, and biochemical markers. J. Clin. Endocrinol. Metab. 67, 741–748. Clarke, B. L., Ebeling, P. R., Jones, J. D., Wahner, H. W., O’Fallon, W. M., Riggs, B. L., and Fitzpatrick, L. A. (1996). Changes in quantitative bone histomorphometry in aging healthy men. J. Clin. Endocrinol. Metab. 81, 2264–2270. Delmas, P. D., Schlemmer, A., Gineyts, E., Riis, B., and Christiansen, C. (1991). Urinary excretion of pyridinoline crosslinks correlates with bone turnover measured on iliac crest biopsy in patients with vertebral osteoporosis. J. Bone Miner. Res. 6, 639–644. Brown, J. P., Delmas, P. D., Malaval, L., Edouard, C., Chapuy, M. C., and Meunier, P. J. (1984). Serum bone Gla-protein: a specific marker for bone formation in postmenopausal osteoporosis. Lancet 1, 1091–1093. Couttenye, M. M., D’Haese, P. C., Van, H. V., Lemoniatou, E., Goodman, W., Verpooten, G. A., and De, B. M. (1996). Low serum levels of alkaline phosphatase of bone origin: a good marker of adynamic bone disease in haemodialysis patients. Nephrol. Dial. Transplant. 11, 1065–1072. Urena, P., Hruby, M., Ferreira, A., Ang, K. S., and De, V. M. (1996). Plasma total versus bone alkaline phosphatase as markers of bone turnover in hemodialysis patients. J. Am. Soc. Nephrol. 7, 506–512. Joffe, P., Heaf, J. G., and Jensen, L. T. (1995). Type I procollagen propeptide in patients on CAPD: its relationships with bone histology, osteocalcin, and parathyroid hormone. Nephrol. Dial. Transplant. 10, 1912–1917. Charhon, S. A., Delmas, P. D., Malaval, L., Chavassieux, P. M., Arlot, M., Chapuy, M. C., and Meunier, P. J. (1986). Serum bone Gla-protein in renal osteodystrophy: comparison with bone histomorphometry. J. Clin. Endocrinol. Metab. 63, 892–897. Lauffenburger, T., Olah, A. J., Dambacher, M. A., Guncaga, J., Lentner, C., and Haas, H. G. (1977). Bone remodeling and calcium metabolism: a correlated histomorphometric, calcium kinetic, and biochemical study in patients with osteoporosis and Paget’s Disease. Metabolism 26, 589–606. Eriksen, E. F., Charles, P., Melsen, F., Mosekilde, L., Risteli, L., Risteli, J. (1993). Serum markers of type I collagen formation and degradation in metabolic bone disease: correlation with bone histomorphometry. J. Bone Miner. Res. 8, 127–132. Delmas, P. D., Demiaux, B., Malaval, L., Chapuy, M. C., Edouard, C., and Meunier, P. J. (1986). Serum bone gamma carboxyglutamic acidcontaining protein in primary hyperparathyroidism and in malignant hypercalcemia. Comparison with bone histomorphometry. J. Clin. Invest. 77, 985–991. Neer, R. M., Arnaud, C. D., Zanchetta, J. R., Prince, R., Gaich, G. A., Reginster, J. Y., Hodsman, A. B., Eriksen, E. F., Ish-Shalom, S., Genant, H. K., Wang, O., and Mitlak, B. H. (2001). Effect of parathyroid hormone (1-34) on fractures and bone mineral density in postmenopausal women with osteoporosis. N. Engl. J. Med. 344, 1434–1441. Jiang, Y., Zhao, J. J., Mitlak, B. H., Wang, O., Genant, H. K., and Eriksen, E. F. (2003). Recombinant human parathyroid hormone (1-34) [teriparatide] improves both cortical and cancellous bone structure. J. Bone Miner. Res. 18, 1932–1941.

594 39. Borah, B., Dufresne, T. E., Chmielewski, P. A., Johnson, T. D., Chines, A., and Manhart, M. D. (2004). Risedronate preserves bone architecture in postmenopausal women with osteoporosis as measured by threedimensional microcomputed tomography. Bone 34, 736–746. 40. Lindsay, R., Nieves, J., Formica, C., Henneman, E., Woelfert, L., Shen, V., Dempster, D., and Cosman, F. (1997). Randomised controlled study of effect of parathyroid hormone on vertebral-bone mass and fracture incidence among postmenopausal women on oestrogen with osteoporosis. Lancet 350, 550–555. 41. Ravn, P., Bidstrup, M., Wasnich, R. D., Davis, J. W., McClung, M. R., Balske, A., Coupland, C., Sahota, O., Kaur, A., Daley, M., and Cizza, G. (1999). Alendronate and estrogen-progestin in the long-term prevention of bone loss: four-year results from the early postmenopausal intervention

KIM BRIXEN AND ERIK FINK ERIKSEN cohort study. A randomized, controlled trial. Ann. Intern. Med. 131, 935–942. 42. Rosen, H. N., Parker, R. A., Greenspan, S. L., Iloputaife, I. D., Bookman, L., Chapin, D., Perlmutter, I., Kessel, B., Qvist, P., Rosenblatt, M., and Ravn, P. (2004). Evaluation of ability of biochemical markers of bone turnover to predict a response to increased doses of HRT. Calcif. Tissue Int. 74, 415–423. 43. Dobnig, H., Sipos, A., Jiang, Y., Fahrleitner-Pammer, A., Ste-Marie, L. G., Gallagher, J. C., Pavo, I., Wang, J., and Eriksen, E. F. (2005). Early changes in biochemical markers of bone formation correlate with improvements in bone structure during teriparatide therapy. J. Clin. Endocrinol. Metab. 90, 3970–3977.

Chapter 36

Genetic Markers of Joint Disease Michel Neidhart

Research Scientist, INSERM research unit 403 and Vice-President Synarc Molecular Marker Division, Lyon, France

Renate E. Gay

Research Scientist, INSERM research unit 403 and Vice-President Synarc Molecular Marker Division, Lyon, France

Steffen Gay

Research Scientist, INSERM research unit 403 and Vice-President Synarc Molecular Marker Division, Lyon, France

V. Juvenile Rheumatoid Arthritis VI. Conclusion References

I. Introduction II. Ankylosing Spondylitis III. Reactive Arthritis IV. Rheumatoid Arthritis

I. INTRODUCTION

II. ANKYLOSING SPONDYLITIS

Genes associated with connective tissue diseases were recognized in two different ways: association studies and linkage analysis [1]. Association studies have confirmed a genetic influence in disease pathogenesis, especially in spondyloarthropathies (SpA), rheumatoid arthritis (RA), and juvenile rheumatoid arthritis (JRA). One of the strongest associations is with the human major histocompatibility (HLA) region. RA has an estimated genetic contribution of 30–50%, approximately one-third of which arises from the HLA complex on chromosome 6p21.3. Many studies have implicated alleles of HLA-DRB1 that encode a “shared epitope”. However, more recent studies have suggested that additional genetic influences exist. As techniques improve, genome scan studies confirmed that multiple genes are involved in each of the connective tissue diseases, with some genes in common that confer susceptibility to autoimmunity.

Ankylosing spondylitis (AS) is a highly heritable, common SpA, primarily affecting the axial skeleton. Genetic influences in this disease are reflected by the fact that children of AS patients exhibit an increased risk of developing the disease later in life [2, 3] and the obvious association with HLA-B27 [4]. It is present in 80–95% of patients with AS, in contrast to only 8% of the general population [5] (Fig. 1).

Dynamics of Bone and Cartilage Metabolism

A. HLA-B27 HLA-B27 is a serologic marker that encompasses 25 different alleles encoding 23 different products [5–7]. These alleles are also called subtypes of HLA-B27, and they may have evolved from the most widespread subtype, i.e. B*2705. These subtypes vary in frequency in different 595

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As a HLA class I protein, its primary function is to complex with β2-microglobulin forming a structure that presents short antigenic peptides for recognition by cytotoxic T lymphocytes [4]. It has been proposed that the role of HLA-B27 in SpA involves this process of antigen presentation, and of the numerous theories proposed to explain the association, the most popular have involved the binding and presentation of “arthritogenic” peptides [9].

B. Infectious Agents

Figure 1

Influence of genetic and environmental factors in the development of ankylosing spondylitis: HLA-B*2705 and polymorphisms of defined cytokine genes, but also the response to infectious agents (e.g., enterobacteria) are involved.

ethnic groups and seem to show different disease associations. HLA-B*2705 is strongly associated with AS and reactive arthritis. Occurrence of AS or related SpA has thus far been documented in subjects possessing any one of the first ten (B*2701 to B*2710) subtypes studied. However, B*2706 in Southeast Asian and B*2709 in the Italian island population of Sardinia appear not to be associated with AS. The B*2805 and B*2709 molecules differ by a single amino acid change, aspartic acid in B*2705, or histidine in B*2709 at position 116. Nevertheless, they have an identical B pocket in the base of the antigen-binding groove [8]. The high frequency of the HLA-B*2705 allele in patients with SpA, especially AS and reactive arthritis, has emerged as probably the best example of a disease association with a hereditary marker. The 13 most recent subtypes (HLA-B*2710 to B*2723), however, have not yet been studied for disease association. It is important to investigate which of them are and are not associated with AS and related SpA, and whether certain subtypes show any preferential association with some of the clinical features or forms of these diseases among the various ethnic populations and geographic regions of the world. This is expected to provide clues to the mechanism of disease association, and one of the strongest reasons to study the B27 subtypes is to learn the effects of the sequence variations on the peptide-binding specificity of the molecule.

The population frequency of SpA in most groups is proportional to that of HLA-B27. But the frequency of HLA-B27 in the population at large far exceeds that of SpA, suggesting other genetic and environmental factors also are operative. In fact, twin studies indicate that HLA-B27 contributes only 16–24% of the total genetic risk for disease [5, 10, 11]. The reason that only a small fraction of HLA-B27-positive individuals develop SpA is still unknown [10]. Much experimental evidence supports the fact that additional genetic factors influence the disease and that the environmental factors may be ubiquitous. Many HLA-B27-linked diseases begin after an infection with enterobacteria, suggesting a role for environmental antigens in addition to a HLA-B27 molecule [7]. To delineate the role of infection, studies have been carried out in animal models of reactive arthritides. It has been shown that HLA-B27 transgenic rats spontaneously develop a multisystemic inflammatory disease resembling human SpA [5, 12]. This disease is dependent on the presence of a normal bacterial flora and implicates the immune system. The presence of both T cells and antigen-presenting cells expressing high levels of HLA-B27 appears of critical importance in its pathogenesis. HLA-B27 transgenic mice also develop arthritis under the influence of the bacterial flora. In both types of model, CD8+ T cells are not necessary, arguing against the “arthritogenic” peptide hypothesis. In vitro models have been largely used to study the immune response against bacterial agents and the role of HLA-B27 in human SpA [12]. It appears that an impaired immune response against bacteria could be involved in triggering the disease. HLA-B27 could be implicated at the level of interaction between host cells and bacteria in driving a specific immune response against bacterial antigens or as a target of an autoimmune response. Thus, a predominant view is that AS is an autoimmune disease based on cross-reactivity between bacteria and HLA-B27. This hypothesis, however, has also become controversial [13]. Despite considerable efforts to define how HLA-B27 contributes to disease, its role remains enigmatic. Increasing evidence suggests it has effects that are unrelated to its

Chapter 36 Genetic Markers of Joint Disease

physiologic function. The basis for this is unknown but may be a consequence of the unusual tendency of this allele to misfold [14].

C. Other Genetic Markers Family studies have implicated other genes outside HLA that further enhance the susceptibility to AS [6]. Genomewide scans have implicated regions on chromosomes 2q, 6p, 6q, 10q, 11q, 16q, 17q, and 19q [6]. A recent study identified linkages with regions on chromosomes 2q, 11p, and 18p [15]. Regarding gender, clinical manifestations appear similar in men and women, whereas radiological features appear more frequent and severe in males [2]. In female patients with AS, spinal disease is unchanged while peripheral arthritis and uveitis are suppressed during childbearing. Furthermore, HLA-B27-positive subjects appear to have a low tumor necrosis factor-α (TNF-α) secretor status, possibly leading to diminished immune responses against certain microbes [16]. The frequencies of six polymorphic spanning in the region of genome close to the TNF gene have been investigated in a group of HLA-B27-positive patients with AS [17]. The TNF 308.1 allele is significantly increased in patients with AS, compared to the HLA-B27-positive healthy controls. These observations provide further evidence supporting the involvement of genes other than HLA-B27 in the susceptibility, pathogenesis, and features of this disease. Other potential genes include HLA-related genes, such as low-molecular-weight proteasome (LMP) and transporter associated with antigen processing (TAP), genes in the neighborhood of the HLA locus, such as HSP-70 and TNF-α, as well as non-HLA genes (IL-1RA, IL-6, IL-10, and CYP2D6) [5, 18, 19]. The search for nonHLA candidate genes, however, has been complicated by inadequate power, because of the small effect they exert on overall disease susceptibility [5, 11].

597 Yersinia, Shigella, Salmonella, and Klebsiella spp. [16, 20]. In HLA-B27-positive patients, Clostridium difficile [21] has also been associated with a form of reactive arthritis. However, other forms appeared unrelated to HLA-B27, including those triggered by group A streptococcus (rheumatoid fever), Borrelia burgdorferi (Lyme disease), Tropheryma whippelii (Whipple’s disease), Chlamydia spp., Giardia lamblia, hepatitis C virus, and Mycoplasma fermentans [16, 22]. HLA-B27-associated diseases may occur identical to a special form of reactive arthritis (previously referred to “Reiter’s syndrome”) with accompanying ureteritis and/or conjunctivitis, whereas in the HLA-B27nonassociated form this has not been clearly described [23] (Fig. 2). The link between HLA-B27 and infection is mirrored by the development of arthritis in HLA-B27-transgenic rats [7, 12]. As mentioned above, arthritis does not develop in animals maintained in a germ-free environment. However, infections of the gastrointestinal, genitourinary, and respiratory tract appear to provoke a reactive arthritis and a wide range of pathogens has now been implicated. Indeed, both antigens and DNA of several microorganisms have been detected in joint material from patients with reactive arthritis. However, the role of such disseminated microbial

III. REACTIVE ARTHRITIS The clinical features of reactive arthritis are those shared by other members of the SpA family, though it is distinguished by a clear relationship with a precipitating infection. Although several forms of disease could be described as “reactive”, particularly acute rheumatic fever, postmeningococcal septicemia arthritis and Lyme disease, in clinical practice the term is restricted to an acute spondyloarthritis, usually, but not exclusively, linked to acute genitourinary or gastrointestinal infection. Enterobacteria associated with HLA-B27 spondyloarthritis include

Figure 2 Influence of genetic and environmental factors in the development of reactive arthritis: HLA-B*27 and the response to certain bacteria, e.g. Yersinia, Shigella, Salmonella, and Klebsiella spp., are involved. However, other forms appeared unrelated to HLA-B27, e.g. those triggered by Group A streptococcus, Borrelia burgdorferi, Tropheryma whippelii, Chlamydia spp., Giardia lamblia, hepatitis C virus, and Mycoplasma fermentans.

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elements in the provocation or maintenance of arthritis remains unclear. HLA-B27-restricted T-cell responses to microbial antigens have been demonstrated and these may be important in the pathogenesis of the disease. HLA-B27-restricted CD8+ T cells specific for an ubiquitous self- or crossreactive microbial determinant may play an important role in the early phase of the disease [24]. Synovial lymphocyte proliferation in reactive arthritis to the 60-kDa heat-shock protein of Yersinia appears to play a major role [16]. On the other hand, enterobacterial infection can down-regulate expression of class I HLA molecules in vivo and that downregulation is predominant in patients with the HLA-B27 genotype [25]. Anti-Klebsiella antibodies are associated with gut lesions, but their significance for the pathogenesis of spondyloarthropathies remains uncertain [16]. Although no single class II allele is found to be increased among HLA-B27-positive patients with reactive arthritis, HLADRB1 *0408 (DR4) and DQB1 *0301 (DQ8) might represent markers conferring disease susceptibility [26]. The way in which a host accommodates invasive facultative intracellular bacteria must be the key to the development of reactive arthritis [27].

Figure 3

Influence of genetic and environmental factors in the development of rheumatoid arthritis: in this disease of unclear etiology, HLADRB1 shared epitopes are involved together with e.g. polymorphisms to cytokine and their receptor genes, as well as epigenetic alterations (ERV = endogenous retroviruses).

IV. RHEUMATOID ARTHRITIS Rheumatoid arthritis (RA) is characterized by persistent joint inflammation and concomitant destruction of the affected joint. The clinical course ranges from mild joint swelling to severe polyarthritis with progressive destruction of cartilage and bone. Although joint swelling is a major clinical feature, destruction of bone and cartilage might be dissociated from inflammation [28]. In recent years, research has focused on the identification of genes that influence the susceptibility as well as the severity of this disorder. On the one hand, the use of T cell-directed biological therapies for RA has been disappointing [29]. Nevertheless, the shared epitope, a common peptide sequence on the antigen-binding regions of some HLA-DR4 subtypes and more precisely on the HLA-DRB1 sequence, is associated with an increased prevalence and severity of RA [30]. On the other hand, processes underlying erosive RA accompanied by synovial tissue hyperplasia and activation of transformed-appearing synovial lining cells could represent more promising targets [28, 31]. For instance, promoter polymorphisms of TNF-α are associated with a more aggressive disease [32]. However, allelic polymorphisms of these genes can only partly explain the variance of the clinical course, because the genetic background of RA involves multiple genes [33] (Fig. 3).

A. Class II HLA-DRB1 “Shared Epitope” Investigations on the precise role of HLA genes in RA have confirmed the importance of this genetic risk factor and have identified a consensus sequence within the HLA-DR genes. The major risks for RA are conferred by female gender and associated with HLA-DRB1 genes that share a five amino acid sequence motif, QKRAA or QRRAA, from position 70 to 74 in the third hypervariable region (HV3) [34]. These so-called HLA-DRB1 “shared epitope” alleles are more common in IgM rheumatoid factor-positive RA patients [35]. The presence of shared epitope alleles is a particularly strong predictor of early erosions in seronegative RA patients. DRB1 alleles encoding the “shared epitope” have a dominant influence over “protective alleles” (e.g., the DERAA motif) [36]. A Spanish study reported that HLA-DR1 alleles are increased among male patients, whereas HLA-DR4 and DR10 alleles are preferentially associated with female RA [37]. The HLA polymorphism appears to play a dual role: (1) operating in disease induction in combination with gender, and (2) influencing disease severity and progression independently of gender. The establishment of functional transgenic mice expressing RA-associated HLA class II molecules has proven to

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be useful in the delineation of the role of these molecules in immune responses, possibly related to RA and in the development of humanized models for this disease [38]. Such humanized mice develop arthritis upon immunization with type II collagen (CII), which shows similarities with RA. Interestingly, the immunodominant T-cell determinant in CII is derived from positions 261–273, which overlap with a previously identified CII T-cell epitope restricted by the mouse Aq molecule, which is associated with collagen-induced arthritis. Studies in collagen transgenic mice have shown that recognition of this peptide may lead either to T-cell tolerance or to an arthritogenic response. It has been proposed that the T-cell recognition of the CII peptide bound by DR molecules is one of the molecular interactions of critical importance in the development of RA and accordingly also an important target for prevention and treatment of this disease. RA patients with autoantibodies to CII frequently showed the HLADRB1 *0101 allele (DR1) [39]. The observation that HLA polymorphisms are mostly associated with disease progression and severity and that a gene dose effect for HLA-DR genes is operational has challenged the simple model that HLA molecules select and present an arthritogenic antigen. These associations between RA and HLA-DR genes are, however, incomplete in that about 25% of patients do not carry RA-susceptibility HLA-DRB1 “shared epitopes” [40].

B. Class II HLA-DQ In early studies, the HLA-DRQ1 *0301 allele (DQ8) was not found to be associated with RA when linkage disequilibrium with HLA-DRB1 was considered [26, 41]. Subsequently, however, high risk for IgM rheumatoid factors and joint erosion have been reported in patients with both HLA-DRB1 “shared epitope” and HLA-DRQ1 *0301 [42, 43]. Various experimental animal models of autoimmune disease have been studied to address the role of immune response genes [44]. To study the interactions involved between class II molecules (DR and DQ) and define the immunologic mechanisms in various diseases, HLA-DR and DQ transgenic mice have been generated that lacked endogenous class II molecules. The HLA molecules in these mice are expressed on the cell surface and can positively select CD4+ T cells expressing various V beta T-cell receptors. A peripheral tolerance is maintained to transgenic HLA molecules thus indicating that these molecules act as self. Results obtained from HLA-DQ8 transgenic mice in the model of type II collagen-induced arthritis suggested that complementation of the HLA-DR

and HLA-DQ alleles may determine both disease susceptibility and severity in RA [45]. Thus, it is possible that certain HLA-DQ alleles, such as HLA-DQ8, predispose individuals to RA, while a self-peptide derived from the HV3 of HLA-DR alleles, such as DRB1 *0402, can protect disease if presented by the DQ molecule. To test this hypothesis, the immunomodulatory effects of the DRB1 *0402 derived peptide on collagen-induced arthritis in HLA-DQ8 transgenic mice have been examined [46]. Co-immunization of the DRB1 *0402 peptide significantly reduced the severity of arthritis, whereas multiple doses of the peptide reduced the incidence of disease. Subsequent analysis revealed that the DRB1 *0401 peptide-mediated protection may be due to the generation of a subset of regulatory cells, which down-regulate collagen-specific autoimmunity. Taken together, these recent reports emphasized the complex degree of genetic interaction between alleles at several HLA loci. At least one additional non-DRB1 susceptibility locus for RA exists in an interval that encompasses the junction of the class III and I regions [47]. This is a genomic segment of high linkage disequilibrium containing a large number of poorly characterized immunomodulatory genes.

C. Class II HLA-DM In RA, the first studies investigating the polymorphisms of the DQCAR region (i.e., the region between DQ and DR) and of the HLA-DM genes found no association with disease susceptibility [48–50]. HLA-DM is a nonclassical MHC class II molecule that has been shown to facilitate peptide loading with classical class II molecules. The DM heterodimer (A/B) is an antigen presentation regulator and may be linked to immune system alterations. A recent study reported that the DMA *0101 allele confers a predisposition to RA, while the DMA *0102 allele protects from the disease [51]. Ethnical differences exist and whether or not this association appears secondary to a linkage disequilibrium with DRB1 “shared epitope” is controversial [50, 52]. In French RA patients, an increased frequency of HLADMB *0101 and HLA-DMA *0103 [53] alleles has been found independently of a linkage disequilibrium with DRB1 alleles, but this appears not to be the case for US Caucasians [52] or Thais [54]. The transporter associated with antigen-processing (TAP) gene products is involved in the processing of endogenous peptides that bind to class I molecules. Polymorphism within these genes could alter the level of the immune response, a phenomenon relevant to the development of autoimmune diseases. The relation between

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RA and polymorphism of large-molecular-weight proteasome (LMP) and transporter associated with TAP genes has been investigated by different groups [55–57]. Higher frequency of a TAP2 gene polymorphism (TAP2A/A) was detected in HLA-DRB1*04-positive RA patients, compared with healthy controls [56].

highly conserved HSP70 [62]. This may cause abnormal trafficking of HLA-DRB1*0401 in B cells and/or abnormal T cell responses to bacterial and human HSP70 in people who express HLA-DRB1*0401. However, the polymorphism of HSP70 genes does not influence disease susceptibility [63]. 4. TUMOR NECROSIS FACTOR ALPHA

D. HLA Class III 1. I KAPPA BL

In a set of Japanese patients with RA and a control group, an RA-susceptibility gene within the telomeric HLA class III region has been described [58]. This locus has been identified as the T allele of SNP 96452 (T/A), in the promoter region (position −62) of the I kappa BL gene. This −62T/A SNP disrupts the putative binding motif for the transcriptional repressor, delta EF1, and hence may influence the transcription of I kappa BL, homologous to I kappa B alpha, the latter being a known inhibitor of NF kappa B, which is central to innate immunity. Thus, the HLA may harbor RA genetic determinants affecting the innate and adaptive arms of the immune system. 2. D6S273

The role of a CA repeat polymorphic microsatellite marker, D6S273, located between HSP70 and Bat2 genes in the class III region of HLA, has also been examined. Two D6S273 alleles (132 and 138) showed significant differences in their prevalence in RA patients as compared to healthy controls; allele 132 was higher in “shared epitope”positive, and allele 138 in “shared epitope”-negative patients [59]. The D6S273 138 allele shows a primary association with RA-susceptibility in “shared-epitope”-negative patients [59, 60]. These authors concluded that at least three separate regions in the HLA complex may contribute to susceptibility to RA, namely class II (DRB1) and class III (D6S273, HSP70, TNFa2) [40, 60, 61]. 3. HEAT SHOCK PROTEINS (HSPs)

Antibodies to heat shock protein 90 kDa (HSP90) are common in longstanding RA patients with articular erosions. In this context, it is important to note that the bacterial DnaJ protein, which shows homology to Drosophila Tid56 protein and vertebrate heat shock proteins, can be detected in RA synovial tissues. Thus, cross-reactivities between host heat shock proteins and infectious agents has been postulated. Furthermore, the induction of heat shock protein 70 kDa (HSP70) in rheumatoid synovial tissue is induced by pro-inflammatory cytokines. The unique properties of the QKRAA motif of HLA-DRB1*0401 constitutes a binding motif for the

The TNF locus is situated in the class III region of the HLA gene complex. TNF-α and interleukin-1 (IL-1) are considered pivotal cytokines in the process of human RA [31]. Studies in experimental models have revealed that TNF-α is a pivotal cytokine in acute joint swelling, yet IL-1β is the dominant cartilage destructive cytokine and its production may occur independently of TNF-α. This was found with anti-TNF/IL-1 neutralizing antibodies and the observations were recently supported by similar findings in arthritis models in TNF and IL-1 knockout mice. However, it needs to be stressed at this point that animal models have only limited value in reflecting the conditions in human RA. This is illustrated by the fact that several studies have revealed that anti-TNF-directed therapies (and not anti-IL-1-directed therapies) slow down or even arrest the destructive process in numerous RA patients. In humans, it is expected that functional heterogeneity in the TNF gene may have an influence on the inflammatory response [64] and probably also the destructive process in RA. Early studies [32, 65] investigated the polymorphism in the promoter region of TNF-α gene in adult and juvenile RA. The TNF –238G (A to G substitution) promoter polymorphism has been associated with a more destructive disease in RA, independently of the presence of HLADRB1 “shared epitope”. In Japanese RA patients, the −857T allele which is associated with systemic juvenile RA [65] has been found in linkage disequilibrium with the HLA-DRB1 *0405 allele [66]. The TNF locus is flanked by five microsatellite markers. Evidence from murine models has indicated that variation in this region could be associated with altered regulation of TNF biosynthesis. These TNF microsatellites have been examined in Caucasian RA patients for possible association with disease susceptibility [67, 68]. Differences in frequencies can be detected at the so-called TNFa and TNFe loci, but these are in linkage disequilibrium with HLA-DRB1 “shared epitope” alleles [67]. Thus, at this point, it has been concluded that there is little, if any, independent contribution of the TNF locus to the genetic background for RA susceptibility [67, 68]. Refining the analysis, other studies suggested that the association of the HLA-DRB1 “shared epitope” with disease severity in RA is influenced by an interaction with the TNFa6 [36, 66]

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and TNFa11 microsatellites [69]. The interaction with TNFa6 appears to be acting predominantly in female patients [36]. Interestingly, TNFa11 has been associated with the responder population to an anti-TNF therapy [70].

E. T Cell Receptor A dense infiltrate of activated T cells, macrophages, and B cells in the synovial membrane is one of the pathological features of RA. Frequently, tissue-infiltrating cells acquire a morphological organization reminiscent of secondary lymphoid tissue. The composition of the inflammatory lesions, the production of autoantibodies, and the association of disease risk with genes related to the HLA-DR and DQ regions have all been cited as evidence for a critical role of T cells in disease pathogenesis [71]. The selection of the primary T-cell repertoire appears to constitute a predisposing factor [72]. The synovial T-cell repertoire might be influenced by aberrations in the naive T-cell repertoire that indicate a defect in thymic education with the selection of high-affinity self-reactive T lymphocytes. RA has been postulated to result from a synovial immune response to an unidentified antigen(s), which should be mirrored by the T-cell response. However, studies analyzing the repertoire of tissue infiltrating T lymphocytes have not been able to identify a dominant and common disease-relevant T cell. CD4+ T cells are a major component of the inflammatory infiltrate in rheumatoid synovitis. In general, the clonal size of individual CD4+ T-cell clones is small and does not lead to a marked distortion of the synovial T-cell receptor (TCR) repertoire [73]. Thus, the associations between RA and specific TCRs are very weak [74] or the repertoire is highly heterogeneous even in early stages [75]. The comparison of TCR sequences derived from different patients has not provided evidence for common sequences. Either multiple antigens are recognized or the TCR repertoire is sufficiently plastic with a multitude of different TCR structures responding to the same antigen(s). However, within one individual, the repertoire of clonal T-cell populations is restricted. Identical T-cell clones can be identified in different joints and at different time points of the disease, emphasizing that the spectrum of antigens recognized might be conserved over time and that the T-cell response pattern is not subject to evolution [73]. Forty percent of accumulated T-cell clones found in one joint of a patient were found in multiple joints in the same patient [76]. Interestingly, T lymphoyctes derived from a subpopulation of RA patients are able to induce synovial hyperplasia in SCID mice [77]. These T cells preferentially use certain TCR V beta chains (V beta 8, 12, 13, and 14). In addition, the CD4+ T cell response

to the human cartilage antigen glycoprotein 39 (HCgp-39) has been analyzed in the context of RA-associated HLADRB1 *0401 and nonassociated HLA-DRB1 *0402 molecules. T-cell epitopes of HCgp-39 that were defined in HLA-DR4 transgenic mice stimulate T lymphocytes from human subjects carrying RA-associated HLA-DR4 alleles. HLA-DR4 molecules may influence the disease process in RA both by presentation of selected peptide epitopes and by promoting the local production of interferon-γ and tumor necrosis factor α [71]. Mouse co-stimulatory and accessory molecules can interact with the HLA-peptide– TCR complex leading to efficient T-cell activation [44]. Recent evidence indicates that RA patients carry expanded CD4 clonotypes which are characterized by deficient CD28 expression and autoreactivity. These autoreactive CD4 T cells are not restricted to the joint, raising the possibility that rheumatoid synovitis is a manifestation of a systemic autoimmune disease. Support for this model has come from studies in TCR transgenic animals which develop inflammation of the synovial tissue stimulated by a T cell response to ubiquitously expressed self-MHC molecules. Antigens driving the chronic persistent immune response in RA may not be restricted to the joint but rather may be widely distributed, providing an explanation for the difficulties in identifying arthritogenic antigens directly or indirectly through the selection of joint-infiltrating T cells [71].

F. Infectious Agents The most well-established association with both susceptibility and severity of disease is variations in the HLA genes. This fact constitutes evidence in favor of a contribution from specific HLA class II restricted adaptive immunity to the pathogenesis of RA. However, considerable difficulties have been encountered in identifying reactivities within the adaptive immune system that are responsible for the development of chronic arthritis in humans. Infectious agents have been discussed as possible etiological agents triggering or modulating certain pathways in this disease. For instance, synovial Epstein–Barr virus (EBV) infection has been suggested to function as an environmental risk factor for RA, particularly in patients with “shared-epitope” alleles [78]. HLA-DR polymorphism influences the frequency of T lymphocyte specific for EBV gp110, a protein of the late replicative cycle [79]. HLADRB1 07, an allele associated with reduced risk to develop RA, is associated with the highest frequencies of T cells specific for gp110 in peripheral blood. In contrast, HLADRB1 *0404, one of the susceptibility alleles, is associated with the lowest frequencies of gp110 specific T cells.

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The T cell repertoire in RA patients is prone to recognize certain microbial products and autoantigens. The selection of this response pattern can only partially be attributed to the disease-associated HLA-DRB1 alleles [73]. APCs are abundantly present in the synovial lining and produce large quantities of proinflammatory mediators through which they are involved in the breakdown of cartilage. In order to orchestrate their functions, APCs are well equipped with several receptors involved in antigen uptake and processing, including Toll-like receptors (TLRs). TLRs are phylogenetically conserved receptors that recognize pathogen-associated molecular patterns and play an extremely important role in both innate and adaptive immune responses. TLR-4 is the receptor most thoroughly investigated. Apart from being involved in the recognition of lipopolysaccharide (LPS), TLR-4 also interacts with endogenous ligands such as HSPs, fragments of hyaluronic acid, and fibronectin. These endogenous ligands are released by cells undergoing stress, damage, or necrotic death and are abundantly present in inflamed synovial joints of patients with RA. An association was reported between disease susceptibility and a functional TLR-4 variant in a large set of RA patients [80]. The TLR-4 polymorphism was also associated with the level of disease activity at baseline, but not with disease severity or outcome. Current research on the expression of TLRs in RA is in progress [81, 82].

G. Cytotoxic T-Lymphocyte Antigen-4 Cytotoxic T-lymphocyte antigen-4 (CTLA4) is considered to be one of the attractive candidates for the susceptibility genes to rheumatic diseases. CTLA4 polymorphisms located in the promotor region at positions −318 (substitution from C to T) and in exon 1 (position 49, substitution from A to G) were investigated in RA patients. When the CTLA4 49 A/G polymorphism was analyzed, a significant increase of A/G heterozygous individuals among female patients (48% versus 34% in healthy controls) was observed. This increase was absent among males (38%) [83]. Particularly, RA patients positive for HLADRB1 “shared epitope” showed more frequently the CTLA4 49 A/G polymorphism [84]. In Japanese populations, however, the CTLA-4 49 A/G polymorphism does not contribute significantly to the susceptibility to RA.

candidate RA locus. Indeed, in Chinese RA patients, a synergistic effect for susceptibility to RA was found between the tumor necrosis factor receptor 2 microsatellite allele 15 (TNFR2ms 15) and HLA-DR4 [85]. A case– control study in a United Kingdom Caucasian population has shown an association between a TNFRII genotype (196R/R in exon 6) and familial, but not sporadic, RA [86]. An additional TNFRII recessive factor, in linkage disequilibrium with the 196R allele, appears to play a major role in a subset of families with multiple cases of RA.

I. Interleukin-1 There are two members of the IL-1 gene family: IL-1α and IL-1β. They are considered to be important cytokines in the pathogenesis of many inflammatory diseases. Although anti-IL-1 treatment has been shown to cause amelioration of arthritis in animal models and in RA [31], clinical studies with IL-1 receptor antagonist (IL-1RA) have not supported these earlier animal studies claiming that IL-1 is a major player in the pathogenesis of RA. An early study [87] investigated whether cytokine gene polymorphisms could be used as markers for susceptibility or severity in RA. The combination of an RA-related HLA-DR allele expressing “shared epitope” and the presence of allele E2 in IL-1β exon 5 is found to expose patients to an increased risk of erosive disease. Similarly, another study [88] showed that carriage of the rare IL-1β (+3954) allele 2 is associated to destructive forms of the disease. On the other hand, a rare IL-1β –511 promotor polymorphism appears to be associated with a less severe course in RA of long duration [89]. IL-1α gene polymorphism has been analyzed for exon 5 (IL-1A1 and IL1A2) and promoter region in RA patients with destructive and nondestructive diseases [90]. The IL-1A2 allele was found in 45% of healthy controls, 54% in destructive and 27% in nondestructive RA; the difference between destructive and nondestructive diseases appeared significant. All parameters of disease activity and joint destruction were significantly higher in patients positive for IL-1A2, and lower in those positive for IL-1A2. Thus, the presence of the IL-1A2 allele could constitute a risk factor for the development of destructive RA.

J. Interleukin-1 Receptor Antagonist H. Tumor Necrosis Factor Receptor II TNF-α binds the receptors TNFRI and TNFRII. Results of genome scans have suggested that TNFRII is a

The gene encoding IL-1RA has a variable allelic polymorphism. The IL1RN*2 allele was described as a factor of severity in several autoimmune diseases and was paradoxically associated with increased production of IL-1RA

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by monocytes in vitro. In definite primary Sjögren’s syndrome, the IL1RN*2 allele was significantly more frequent [91]; in contrast, in RA, its frequency was not different from that of healthy controls. Another study [87] confirmed that IL-1RA gene polymorphism is not associated with the severity of RA.

K. Interleukin-4 IL-4 mRNA in peripheral blood mononuclear cells of active RA are produced at a low spontaneous level and the response to in vitro activation by mitogen is weak. The IL-4 gene shows polymorphisms and the RP1 allele was found with a significantly higher frequency in RA patients, compared with controls [87]. The implication of this polymorphism, however, is unclear so far. On the other hand, carriers of a rare IL-4 gene polymorphism (VNTR) could be protected against severe joint destruction [89, 92].

L. Interleukin-10 IL-10 has anti-inflammatory and immunoregulatory properties that suggest a potential therapeutic role in adult and juvenile RA [93]. IL-10 inhibits proinflammatory cytokine and chemokine production in addition to blocking T-cell responses to specific antigens. It acts primarily through inhibition of costimulatory properties of macrophages. IL-10 stimulates proliferation and differentiation of antibody-forming B cells. Preclinical studies in a variety of animal models, including collagen-induced arthritis, have shown that IL-10 is effective in preventing or inhibiting inflammation and autoreactivity. Although in RA, circulating and synovial levels of IL-10 are increased, accumulated evidence suggests that there may be a relative deficit of available IL-10. The delivery of IL-10 to patients with RA has, however, been disappointing. Various polymorphisms within the IL-10 gene promoter have been identified and appear also to influence its expression [94]. The −1082*A allele has been associated with low and the −1082*G allele with high in vitro IL-10 production. No significant differences in the allele frequencies of the three polymorphisms were found between RA patients and healthy controls [87, 94, 95]. However, the −1082*A allele appears be linked to the regulation of rheumatoid factor isotypes, favoring the production of IgA rheumatoid factors [94]. IgA rheumatoid factors have been associated with severe RA, and may thus be indirectly correlated with a genotype encoding low IL-10 production. In addition, the −2849 IL-10 promoter polymorphism was shown to be associated with

autoantibody production and subsequent joint damage in RA [96].

M. CC Chemokine Receptors The chemokine receptors CXCR3 and CCR5 are markers for T cells associated with certain inflammatory reactions [97]. A 32 base pair deletion allele in the CC chemokine receptor 5 (CCR5 delta 32 allele) affects both transmission of human immunodeficiency virus-1 and acquired immunodeficiency syndrome (AIDS)-free survival. Since chemokines are suggested to be critical for establishment of inflammation in autoimmune diseases, it has been hypothesized that a defective allele may modulate the inflammatory process in RA [98]. The gene frequency of CCR5-delta 32 allele does not deviate significantly from healthy controls (10%) [98, 99]. However, very few RA patients showed the homozygous CCR5delta 32 genotype, compared with healthy controls and systemic lupus erythematosus [99]. Patients carrying the CCR5-delta 32 allele were negative for IgM rheumatoid factors, had less swollen joints and experienced shorter morning stiffness [98]. Therefore, the CCR5-delta 32 polymorphism might be a genetic marker related to the severity of RA.

N. Genotoxic Stress and p53 Mutation In RA, the antioxidant system is impaired and peroxidation reactions are accelerated. It has been proposed that polymorphisms in the glutathione S-transferases (GST) gene may influence susceptibility and progression in RA and that this effect is presumed to be based on differences in the ability of various GST enzymes to utilize the cytotoxic products of oxidant stress [100]. Increased genomic DNA damage occurs in RA due to either defective repair mechanisms or increased production of reactive oxygen intermediates during inflammation. The development of point mutations in the p53 tumor suppressor gene and other key regulatory genes could contribute to convert inflammation into a chronic disease [101]. The process driving the activation of synovial fibroblasts is poorly understood. It is unclear how far the synovial hyperplasia of RA resembles tumor diseases. Along this line some contradictory results exist concerning the role of the tumor suppressor protein p53. Some investigations could show the expression of p53 in the synovial lining including p53 mutations in RA synovium and in RASF, while other research groups could not confirm these data. Indeed, DNA damage and somatic mutations in the

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p53 gene can occur because of genotoxic stress in many tissues, including the skin, colon, and synovial tissue [102].

been shown to be strongly associated with amyloidosis in Japanese and Caucasians [108].

O. Sex Hormones

Q. Apoptosis

RA of young-onset disease referring as a subset with age of onset < 55 years old, affecting more women than men, seems to differ from late-onset disease by more small joint complaints and extra-articular features. Therefore, the etiological role of sex hormones and their receptors in RA has been repeatedly discussed. The restriction fragment length polymorphisms (RFLPs) of the cytochrome P450c 17 alpha (CYP17) gene, denoted as the A1 and A2 alleles, are related to serum sex hormone production. RA patients with the A2 allele tended to develop the disease at a younger age than those without [103]. Furthermore, RFLPs of the estrogen receptor gene are denoted P and X alleles (for Pvu II and Xba I restriction enzymes), and homozygote PP female patients appeared to develop RA at a younger age [104]. In addition, analysis of CAG microsatellites of the androgen receptor gene revealed that shorter repeats, presenting high levels of transactivating activity, are related to younger-age-onset male RA [105]. Thus, it has been concluded that polymorphisms in the CYP17 and sex hormone receptor genes could be associated with the onset of the disease.

Fas (APO-1/CD95) is a cell-surface receptor involved in cell death signaling. Germline mutations in the Fas gene have been associated with autoimmune lymphoproliferative syndrome, and somatic Fas mutations have been found in multiple myeloma. Most patients with non-Hodgkin’s lymphoma and somatic Fas mutations developed autoimmune diseases [109]. A Fas promoter polymorphism (MvaI*2) might be more frequent in RA patients with extraarticular involvement, particularly females, and patients with early onset, compared to healthy controls [110]. Fas promoter polymorphism may also be involved in systemic lupus erythematosus [110], but not in Felty’s and large granular lymphocyte syndromes [111]. Interestingly, the clearest evidence of retrotransposon involvement in rodent arthritis is the insertion of the Etn retrotransposon in the fas gene of MRL-lpr/lpr mice [112]. This insertion causes massive lymphoproliferation and accelerated autoimmunity, but there is no evidence that such a mutation may play a role in the induction of autoimmune disease in man. More promising was the observation that autoimmunity develops in mice deficient in programmed death 1 (PD-1), suggesting that PD-1 is a candidate gene involved in the development of human autoimmune diseases. Interestingly, the human PD-1 gene was found to be significantly associated with disease development in RA patients [113].

P. Serum Amyloid A The serum amyloid A (SAA) family comprises a number of differentially expressed apolipoproteins, acute-phase SAAs (A-SAAs, SAA-1) and constitutive SAAs (C-SAAs, SAA-2). SAA-1 is a major acute-phase reactant, which increases by as much as 1000-fold during inflammation; it is the chief constituent of fibrillar deposits in reactive amyloidosis and has also been implicated in the pathogenesis of atheroscelerosis and RA [106]. In vitro, the dramatic induction of SAA-1 mRNA in response to proinflammatory stimuli is due largely to the synergistic effects of cytokine signaling pathways, particularly including interleukin-1 and interleukin-6 type cytokines. Studies of the SAA-1 promoter in several mammalian species have identified a range of transcription factors that are variously involved in defining both cytokine responsiveness and cell specificity. These include NF-κB, C/EBP, YY1, AP-2, SAF, and Sp1. The γ allele at the amyloid protein SAA-1 gene appears to render RA patients susceptible to amyloidosis [107]. A single nucleotide polymorphism at position −13 in the SAA-1 5′ flanking region has

R. Endogenous Retrovirus Some retroviruses appear to be harmless, others are associated with various diseases such as malignancy, immune deficiency, neurological and autoimmune diseases. The arthritis seen in ungulates infected by certain lentiviruses and the arthropathy in mice that carry the tax gene of human T-leukemia virus-1 (HTLV-I) allow investigation of the induction of arthritis by exogenous retroviruses. HTLV-1 causes a destructive arthropathy that has many similarities with RA [114]. However, attempts to isolate known infectious retroviruses from synovial tissues or cultured synovial fibroblasts of RA patients remained unsuccessful [115, 116]. Occasionally, during vertebrate evolution, retroviruses have integrated into the germline and have thus become Mendelian traits of their hosts. Such endogenous retroviruses (ERVs) usually become disabled by the insertion of stop codons or frame shifts

Chapter 36 Genetic Markers of Joint Disease

during the course of evolution. ERVs can maintain open reading frames which are expressed as protein. It has been suggested that such proteins have been retained throughout evolution, because they have a function of benefit to the host, e.g. as cellular reverse transcriptase (RT) [117]. Circulating antibodies to a human ERV, the HTLV-1-related endogenous sequence, are reported in Sjögren syndrome and systemic lupus erythematosus. It is possible that human ERV-related proteins act as autoantigens and thus may be responsible for autoantibodies crossreacting with exogenous retroviral core proteins [118]. A retroviral sequence related to human ERV-9 has been detected in multiple sclerosis [119], and an ERV related to human ERV-K10 might be involved in the pathogenesis of diabetes via a superantigen effect [120]. Type C retrovirus-like particles were found in ultracentrifuged synovial fluid pellets of patients with RA [121]. Based on this finding and on the demonstration that synovial hyperplasia observed in RA is associated with a transformed-appearing phenotype of the synovial cells that express ras and myc proto-oncogenes [28], retroviral RNAs have been searched in synovial fluid pellets. In RA, this approach allowed detection of retroviral-like RNAs, including sequences similar to human ERV-9 and ERV-K10, gibbon ape leukemia virus pol gene and HTLV-I tax gene [122]. A complex expression pattern of multiple ERV transcripts was found in both RA and normal synovial tissues, but specific ERV transcripts appear to be differentially expressed in RA [123]. In this regard, ERV-K long terminal repeats (LTRs) containing regulatory sequences adjacent to the HLA-DQB1 locus have been detected more frequently in RA patients than in healthy individuals [41]. In addition, a sequence highly similar to human retrotransposable L1 (LINE-1) elements has been isolated from an RA synovial fluid pellet [124]. Partial L1 sequences [124, 125] and a full-length L1 retrotransposon [126] have been also isolated from RA synovial tissues. Following this observation, a more comprehensive search for the role of these sequences was undertaken in our laboratory and more conclusive data could be obtained. L1 transcripts could be detected in the sublining layer and at sites of cartilage and bone destruction, but not in osteoarthritis or controls. In vitro, RA synovial fibroblasts expressed L1 mRNAs. The L1-related p40 protein showed a similar pattern of expression. Since genomic DNA hypomethylation occurs in inflammatory diseases, synovial fibroblasts were incubated with the methylation inhibitor 5-aza-2′-deoxycytidine and L1 mRNAs appeared in a time- and dose-dependent manner. Compared with osteoarthritis, RA synovial fibroblasts were more sensitive

605 to DNA hypomethylation. After transfection of RA synovial fibroblasts with a functional L1 element, p38δ, met proto-oncogene and galectin-3 binding protein genes are differentially expressed. Transcription of the p38δ gene was shown to be triggered by DNA hypomethylation and the L1-related p40 protein in vitro [126]. In situ, p38δ has been detected in RA synovial tissues at sites of joint destruction. In turn, p38δ favored the production of catabolic enzymes. Taken together, these results suggest that L1 elements and p38δ downstream pathways play an important role in the activation of RA synovial fibroblasts. These data might represent the basis for the hypothesis that, in addition to the cytokine-driven process in RA, a cytokine-independent pathway of synovial activation might be responsible for the chronic nature of the disease.

V. JUVENILE RHEUMATOID ARTHRITIS Juvenile rheumatoid arthritis (JRA) is the most frequently occurring inflammatory rheumatic condition in childhood. A possible cellular-mediated hypothesis versus a possible B cell hyperreactivity have been entertained [127]. Recent studies have evaluated the interrelationship of T-cell-dependent immune responses in the immunopathogenesis of JRA, and the specific cytokine release. Studies have indicated a pro-inflammatory response in systemic-onset JRA manifested by increased secretion of IL-6, whereas an anti-inflammatory response has been noted by increases of IL-4 mRNA and IL-10 mRNA in pauciarticular-onset JRA. An IL-6 promotor gene polymorphism (−174) has been associated with JRA [128] and a specific IL-10 5′-flanking region haplotype (ATA substitution for GCC) appears to be associated with severe polyarthritis, lower transcriptional activity and production of IL-10 [129]. In addition, it is interesting to note that children with deletion of the long arm of chromosome 18 develop chronic arthritis [130]. Thus, genetic loci on chromosome 18 may play a role in the expression of complex autoimmune diseases. The continued finding of elevated levels of tumor necrosis factor-α (TNF-α) and its receptors in association with inflammatory activity has been seen. The use of a TNF fusion protein to block TNF-α activity in JRA has further contributed to this finding [127]. Another study [65] investigated the polymorphism in the promoter region of tumor necrosis factor α gene in juvenile RA. The combined –1031C (T to C substitution) and −863A (C to A) genotype, and the –857T allele (C to T), both of which being related to high production of TNF-α, appear associated with systemic juvenile RA. The –857T allele

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may enhance the effect of the HLA-DRB1 “shared epitope” haplotype in predisposing to development of systemic JRA. In addition, the presence of HLA-B27 can be associated with sacroiliitis [131].

VI. CONCLUSION The polymorphisms of genes involved in the pathogenesis – including cytokines and proteins involved in antigen processing – and their association with disease susceptibility and/or severity demonstrated that these processes can be due to multiple genetic defects. The association of these polymorphisms with specific HLA alleles reflects the complexity of these associations. During inflammation, oxidant stress and DNA hypomethylation appear to be important factors in the complex interplay of mononuclear cells, synovial fibroblasts and joint destruction. Genotoxic stress appears to result in DNA damage and may be directly causing somatic mutations. While the investigation of T cell repertoire has been disappointing, the search for potential infectious agents – including retroviral sequences – has resulted in new hypotheses regarding the involvement of DNA hypomethylation and endogenous retroviral sequences in a cytokine-independent pathway of chronic and erosive arthritis. In addition, clarifying the role of TLRs and innate immunity will be a major focus of future research. Rheumatic disorders arise in certain individuals depending on the interaction of genetic and environmental factors, the contribution for each varying with the specific disease. A third variable, i.e., random or “stochastic” processes, may be important, but this has been poorly studied. In fact, it is important to note that only a small number of random events need to occur in a predisposed population to allow the emergence of a rheumatic disorder [132]. These random events might be environmental (e.g., infections or exposure to toxins) or due to acquired genetic changes (e.g., somatic mutations involving pivotal immune or growth/repair genes).

References 1. McCurdy, D. (1999). Genetic susceptibility to the connective tissue diseases. Curr. Opin. Rheumatol. 11, 399–407. 2. Gran, J. T., and Ostensen, M. (1998). Spondyloarthritides in females. Baillieres Clin. Rheumatol. 12, 695–715. 3. Brophy, S., Taylor, G., Blake, D., and Calin, A. (2003). The interrelationship between sex, susceptibility factors, and outcome in ankylosing spondylitis and its associated disorders including inflammatory bowel disease, psoriasis, and iritis. J. Rheumatol. 30, 2054–2058. 4. Allen, R. L., Bowness, P., and McMichael, A. (1999). The role of HLA-B27 in spondyloarthritis. Immunogenetics 50, 220–227.

5. Reveille, J. D. (2004). The genetic basis of spondyloarthritis. Curr. Rheumatol. Rep. 6, 117–125. 6. Al-Khonizy, W., and Reveille, J. D. (1998). The immunogenetics of the seronegative spondyloarthropathies. Baillieres Clin. Rheumatol. 12, 567–588. 7. Khare, S. D., Luthra, H. S., and David, C. S. (1998). Animal models of human leukocyte antigen B27-linked arthritides. Rheum. Dis. Clin. North Am. 24, 883–894. 8. Garcia-Peydro, M., Marti, M., and Lopez de Castro, J. A. (1999). High T cell epitope sharing between two HLA-B27 subtypes (B*2705 and B*2709) differentially associated to ankylosing spondylitis. J. Immunol. 163, 2299–2305. 9. Marti, M., Alvarez, I., Montserrat, V., and Lopez de Castro, J. A. (2001). Large sharing of T-cell epitopes and natural ligands between HLA-B27 subtypes (B*2702 and B*2705) associated with spondyloarthritis. Tissue Antigens 58, 351–362. 10. Gonzalez, S., Martinez-Borra, J., and Lopez-Larrea, C. (1999). Immunogenetics, HLA-B27 and spondyloarthropathies. Curr. Opin. Rheumatol. 11, 257–264. 11. Sims, A. M., Wordsworth, B. P., and Brown, M. A. (2004). Genetic susceptibility to ankylosing spondylitis. Curr. Mol. Med. 4, 13–20. 12. Breban, M. (1998). Animal models and in vitro models for the study of aetiopathogenesis of spondyloarthropathies. Baillieres Clin. Rheumatol. 12, 611–626. 13. Ringrose, J. H. (1999). HLA-B27 associated spondyloarthropathy, an autoimmune disease based on crossreactivity between bacteria and HLA-B27? Ann. Rheum. Dis. 58, 598–610. 14. Colbert, R. A., and Prahalad, S. (2001). Predisposing factors in the spondyloarthropathies: new insights into the role of HLA-B27. Curr. Rheumatol. Rep. 3, 404–411. 15. Brown, M. A., Brophy, S., Bradbury, L., Hamersma, J., Timms, A., Laval, S., Cardon, L., Calin, A., and Wordsworth, B. P. (2003). Identification of major loci controlling clinical manifestations of ankylosing spondylitis. Arthritis. Rheum. 48, 2234–2239. 16. Braun, J., and Sieper, J. (1999). Rheumatologic manifestations of gastrointestinal disorders. Curr. Opin. Rheumatol. 11, 68–74. 17. McGarry, F., Walker, R., Sturrock, R., and Field, M. (1999). The –308.1 polymorphism in the promoter region of the tumor necrosis factor gene is associated with ankylosing spondylitis independent of HLA-B27. J. Rheumatol. 26, 1110–1116. 18. McGarry, F., Neilly, J., Anderson, N., Sturrock, R., and Field, M. (2001). A polymorphism within the interleukin 1 receptor antagonist (IL-1Ra) gene is associated with ankylosing spondylitis. Rheumatology 40, 1359–1364. 19. Vargas-Alarcon, G., Londono, J. D., Hernandez-Pacheco, G., Gamboa, R., Castillo, E., Pacheco-Tena, C., Cardiel, M. H., Granados, J., and Burgos-Vargas, R. (2002). Heat shock protein 70 gene polymorphisms in Mexican patients with spondyloarthropathies. Ann. Rheum. Dis. 61(1), 48–51. 20. Simonet, M. L. (1999). Enterobacteria in reactive arthritis: Yersinia, Shigella, and Salmonella. Rev. Rhum. Engl. Ed. 66 (Suppl), 14S–18S. 21. Veillard, E., Guggenbuhl, P., Bello, S., Lamer, F., and Chales, G. (1998). Reactive oligoarthritis in a patient with Clostridium difficile pseudomembranous colitis. Review of the literature. Rev. Rhum. Engl. Ed. 65, 795–798. 22. Johnson, S., Sidebottom, D., Bruckner, F., and Collins, D. (2000). Identification of mycoplasma fermentans in synovial fluid samples from arthritis patients with inflammatory disease. J. Clin. Microbiol. 38, 90–93. 23. Toivanen, P., and Toivanen, A. (1999). Two forms of reactive arthritis? Ann. Rheum. Dis. 58, 737–741. 24. May, E., Dulphy, N., Frauendorf, E., Duchmann, R., Bowness, P., Lopez de Castro, J. A., Toubert, A., and Marker-Hermann, E. (2002). Conserved TCR beta chain usage in reactive arthritis; evidence for selection by a putative HLA-B27-associated autoantigen. Tissue Antigens 60, 299–308.

Chapter 36 Genetic Markers of Joint Disease 25. Kirveskari, J., He, Q., Leirisalo-Repo, M., Maki-Ikola, O., Wuorela, M., Putto-Laurila, A., and Granfors, K. (1999). Enterobacterial infection modulates major histocompatibility complex class I expression on mononuclear cells. Immunology 97, 420–428. 26. Tuokko, J., Reijonen, H., Ilonen, J., Anttila, K., Nikkari, S., Mottonen, T., Yli-Kerttula, U., and Toivanen, A. (1997). Increase of HLA-DRB1 *0408 and –DQB1 *0301 in HLA-B27 positive reactive arthritis. Ann. Rheum. Dis. 56, 37–40. 27. Ikeda, M., and Yu, D. T. (1998). The pathogenesis of HLA-B27 arthritis: role of HLA-B27 in bacterial defense. Am. J. Med. Sci. 316, 257–263. 28. Gay, S., Gay, R. E., and Koopman, W. J. (1993). Molecular and cellular mechanisms of joint destruction in rheumatoid arthritis: two cellular mechanisms explain joint destruction? Ann. Rheum. Dis. 52, S39–S47. 29. Yocum, D. E. (1999). T cells: pathogenic cells and therapeutic targets in rheumatoid arthritis. Semin. Arthritis. Rheum. 29, 27–35. 30. Gregersen, P. K., Silver, J., and Winchester, R. J. (1987). The shared epitope hypothesis. An approach to understanding the molecular genetics of susceptibility to rheumatoid arthritis. Arthritis Rheum. 30, 1205–1213. 31. Van den Berg, W. B., and Bresnihan, B. (1999). Pathogenesis of joint damage in rheumatoid arthritis: evidence of a dominant role for interleukin-I. Baillieres Best Pract. Res. Clin. Rheumatol. 13, 577–597. 32. Kaijzel, E. L., van Krugten, M. V., Brinkman, B. M., Huizinga, T. W., van der Straaten, T., Hazes, J. M., Zeigler-Heitbrock, H. W., Nedospasov, S. A., Breedveld, F. C., and Verweij, C. L. (1998). Functional analysis of a human tumor necrosis factor alpha (TNFalpha) promoter polymorphism related to joint damage in rheumatoid arthritis. Mol. Med. 4, 724–733. 33. Jawaheer, D., and Gregersen, P. K. (2002). Rheumatoid arthritis. The genetic components. Rheum. Dis. Clin. North Am. 28, 1–15. 34. Sherritt, M. A., Tait, B., Varney, M., Kanaan, C., Stockman, A., Mackay, I. R., Muirden, K., Bernard, C. C., and Rowley, M. J. (1996). Immunosusceptibility genes in rheumatoid arthritis. Hum. Immunol. 51, 32–40. 35. El-Gabalawy, H. S., Goldbach-Mansky, R., Smith, D. II., Arayssi, T., Bale, S., Gulko, P., Yarboro, C., Wilder, R. L., Klippel, J. H., and Schumacher, H. R. Jr. (1999). Association of HLA alleles and clinical features in patients with synovitis of recent onset. Arthritis Rheum. 42, 1696–1705. 36. Matthey, D. L., Hassell, A. B., Plant, M. J., Cheung, N. T., Dawes, P. T., Jones, P. W., Thomson, W., Poulton, K. V., Hajeer, A. H., and Ollier, W. E. (1999). The influence of HLA-DRB1 alleles encoding the DERAA amino acid motif on radiological outcome in rheumatoid arthritis. Rheumatology 38, 1221–1227. 37. Gonzalez-Escribano, M. F., Rodriguez, R., Valenzuela, A., Garcia, A., and Nunez-Roldan, A. (1999). Complex associations between HLADRB1 genes and female rheumatoid arthritis: results from a prospective study. Hum. Immunol. 60, 1259–1265. 38. Holmdahl, R., Andersson, E. C., Andersen, C. B., Svejgaard, A., and Fugger, L. (1999). Transgenic mouse models of rheumatoid arthritis. Immunol. Rev. 169, 161–173. 39. Cook, A. D., Stockman, A., Brand, C. A., Tait, B. D., Mackay, I. R., Muirden, K. D., Bernard, C. C., and Rowley, M. J. (1999). Antibodies to type II collagen and HLA disease susceptibility markers in rheumatoid arthritis. Arthritis Rheum. 42, 2569–2576. 40. Singal, D. P., Li, J., and Zhu, Y. (1999). Genetic basis for rheumatoid arthritis. Arch. Immunol. Ther. Exp. 47, 307–311. 41. Seidl, C., Koch, U., Brunnler, G., Buhleier, T., Frank, R., Moller, B., Markert, E., Koller-Wagner, G., Seifried, E., and Kaltwasser, J. P. (1997). HLA-DR/DQ/DP interactions in rheumatoid arthritis. Eur. J. Immunogenet. 24, 365–376. 42. Perdriger, A., Chales, G., Semana, G., Guggenbuhl, P., Meyer, O., Quillivic, F., and Pawlotsky, Y. (1997). Role of HLA-DR-DR and DR-DQ associations in the expression of extraarticular manifestations and rheumatoid factor in rheumatoid arthritis. J. Rheumatol. 24, 1272–1276.

607 43. Cranney, A., Goldstein, R., Pham, B., Newkirk, M. M., and Karsh, J. (1999). A measure of limited joint motion and deformity correlates with HLA-DRB1 and DQB1 alleles in patients with rheumatoid arthritis. Ann. Rheum. Dis. 58, 703–708. 44. Taneja, V., and David, C. S. (1999). HLA class II transgenic mice as models of human diseases. Immunol. Rev. 169, 67–79. 45. Nabozny, G. H., Baisch, J. M., Cheng, S., Cosgrove, D., Griffiths, M. M., Luthra, H. S., and David, C. S. (1996). HLA-DQ8 transgenic mice are highly susceptible to collagen-induced arthritis: a novel model for human polyarthritis. J. Exp. Med. 183, 27–37. 46. Das, P., Bradley, D. S., Geluk, A., Griffiths, M. M., Luthra, H. S., and David, C. S. (1999). An HLA-DRB1 *0402 derived peptide (HSV 65-79) prevents collagen-induced arthritis in HLA-DQ8 transgenic mice. Hum. Immunol. 60, 575–582. 47. Kilding, R., Iles, M. M., Timms, J. M., Worthington, J., and Wilson, A. G. (2004). Additional genetic susceptibility for rheumatoid arthritis telomeric of the DRB1 locus. Arthritis Rheum. 50, 763–769. 48. Singal, D. P., Li, J., and Ye, M. (1998). DQCAR microsatellite polymorphism and susceptibility to rheumatoid arthritis. Tissue Antigens 51, 613–617. 49. Singal, D. P., and Ye, M. (1998). HLA-DM polymorphisms in patients with rheumatoid arthritis. J. Rheumatol. 25, 1295–1298. 50. Reviron, D., Tezenas du Montcel, S., Foutrier, C., Guis, S., Benazet, J. F., Auquier, P., Busson, M., Roux, H., Mercier, P., and Roudier, J. (1999). HLA DRB1, DMA, and DMB gene polymorphisms in rheumatoid arthritis. Hum. Immunol. 60, 245–249. 51. Cucchi-Mouillot, P., Lai, S., Carcassi, C., Silicani-Amoros, P., Floris, L., Amoros, J. P., Genetet, B., Haras, D., and Contu, L. (1999). HLA-DMA alleles are possible new markers of rheumatoid arthritis: study of a corsican group. Exp. Clin. Immunogenet. 16, 192–198. 52. Moxley, G., and Han, J. (2001). HLA DMA and DMB show no association with rheumatoid arthritis in US Caucasians. Eur. J. Immunogenet. 28, 539–543. 53. Toussirot, E., Sauvageot, C., Chabod, J., Ferrand, C., Tiberghien, P., and Wendling, D. (2000). The association of HLA-DM genes with rheumatoid arthritis in Eastern France. Hum. Immunol. 61, 303–308. 54. Vejbaesya, S., Luangtrakool, P., Luangtrakool, K., Sermduangprateep, C., and Parivisutt, L. (2000). Analysis of TAP and HLA-DM polymorphism in Thai rheumatoid arthritis. Hum. Immunol. 61, 309–313. 55. Vinasco, J., Fraile, A., Nieto, A., Beraun, Y., Pareja, E., Mataran, L., and Martin, J. (1998). Analysis of LMP and TAP polymorphisms by polymerase chain reaction-restriction fragment length polymorphism in rheumatoid arthritis. Ann. Rheum. Dis. 57, 33–37. 56. Tuokko, J., Pushnova, E., Yli-Kerttula, U., Toivanen, A., and Ilonen, J. (1998). TAP2 alleles in inflammatory arthritis. Scand. J. Rheumatolm. 27, 225–229. 57. Zhang. S. L., Chabod, J., Penfornis, A., Reviron, D., Tiberghien, P., Wendling, D., and Toussirot, E. (2002). TAP1 and TAP2 gene polymorphism in rheumatoid arthritis in a population in eastern France. Eur. J. Immunogenet. 29, 241–249. 58. Okamoto, K., Makino, S., Yoshikawa, Y., Takaki, A., Nagatsuka, Y., Ota, M., Tamiya, G., Kimura, A., Bahram, S., and Inoko, H. (2003). Identification of I kappa BL as the second major histocompatibility complex-linked susceptibility locus for rheumatoid arthritis. Am. J. Hum. Genet. 72, 303–312. 59. Singal, D. P., Li, J., Ye, M., and Lei, K. (1998). D6S273 microsatellite polymorphism and susceptibility to rheumatoid arthritis. Tissue Antigen 52, 353–358. 60. Singal, D. P., Li, J., and Lei, K. (1999). Genetics of rheumatoid arthritis (RA): two separate regions in the major histocompatibility complex contribute to susceptibility to RA. Immunol. Lett. 69, 301–306. 61. Singal, D. P., Li, J., and Zhu, Y. (2000). HLA class III region and susceptibility to rheumatoid arthritis. Clin. Exp. Rheumatol. 18, 485–491. 62. Auger, I., Toussirot, E., and Roudier, J. (1998). HLA-DRB1 motifs and heat shock proteins in rheumatoid arthritis. Int. Rev. Immunol. 17, 263–271.

608 63. Vinasco, J., Beraun, Y., Nieto, A., Fraile, A., Pareja, E., Mataran, L., and Martin, J. (1997). Heat shock protein 70 gene polymorphisms in rheumatoid arthritis. Tissue Antigens 50, 71–73. 64. Verwij, C. L. (1999). Tumour necrosis factor gene polymorphisms as severity markers in rheumatoid arthritis. Ann. Rheum. Dis. 58 (Suppl 1), I20–I26. 65. Date, Y., Seki, N., Kamizono, S., Higuchi, T., Hirata, T., Miyata, K., Ohkuni, M., Tatsuzawa, O., Yokota, S., Joos, K., Ueda, K., Sasazuki, T., Kimura, A., Itoh, K., and Kato, H. (1999). Identification of a genetic factor for juvenile rheumatoid arthritis in the 5′-flanking region of the TNF alpha gene and HLA genes. Arthritis Rheum. 42, 2577–2582. 66. Seki, N., Kamizono, S., Yamada, A., Higuchi, T., Matsumoto, H., Niiya, F., Kimura, A., Tsuchiya, K., Suzuki, R., Date, Y., Tomita, T., Itoh, K., and Ochi, T. (1999). Polymorphisms in the 5′-flanking region of tumor necrosis factor alpha gene in patients with rheumatoid arthritis. Tissue Antigens 54, 194–197. 67. Field, M., Gallagher, G., Eskdale, J., McGarry, F., Richards, S. D., Munro, R., Oh, H. H., and Campbell, C. (1997). Tumor necrosis factor locus polymorphisms in rheumatoid arthritis. Tissue Antigens 50, 303–307. 68. Waldron-Lynch, F., Adams, C., Shanahan, F., Molloy, M. G., and O’Gara, F. (1999). Genetic analysis of the 3′ untranslated region of the tumour necrosis factor shows a highly conserved region in rheumatoid arthritis affected and unaffected subjects. J. Med. Genet. 36, 214–216. 69. Mu, H., Chen, J. J., Jiang, Y., King, M. C., Thomson, G., and Criswell, L. A. (1999). Tumor necrosis factor a microsatellite polymorphism is associated with rheumatoid arthritis severity through an interaction with the HLA-DRB1 shared epitope. Arthritis Rheum. 42, 438–442. 70. Martinez, A., Salido, M., Bonilla, G., Pascual-Salcedo, D., FernandezArquero, M., de Miguel, S., Balsa, A., de la Concha, E. G., and Fernandez-Gutierrez, B. (2004). Association of the major histocompatibility complex with response to infliximab therapy in rheumatoid arthritis patients. Arthritis Rheum. 50, 1077–1082. 71. Weyand, C. M., and Goronzy, J. J. (1999). HLA polymorphisms and T cells in rheumatoid arthritis. Int. Rev. Immunol. 18, 37–59. 72. Yang, H., Rittner, H., Weyand, C. M., and Goronzy, J. J. (1999). Aberations in the primary T-cell receptor repertoire as a predisposition for synovial inflammation in rheumatoid arthritis. J. Investig. Med. 47, 236–245. 73. Goronzy, J. J., Zettl, A., and Weyand, C. M. (1998). T cell receptor repertoire in rheumatoid arthritis. Int. Rev. Immunol. 17, 339–363. 74. Ibberson, M., Peclat, V., Guerne, P. A., Tiercy, J. M., Wordsworth, P., Lanchbury, J., Camilleri, J., and So, A. K. (1998). Analysis of T cell receptor V alpha polymorphism in rheumatoid arthritis. Ann. Rheum. Dis. 57, 49–51. 75. Fang, Q., Sun, Y. Y., El-Gabalawy, H., Cai, W., Ko, L., Chin, H., Arayssi, T., Schumacher, H. R., and Williams, W. V. (1999). Synovial T cell receptor heterogeneity in early arthritis. Pathobiology 67, 59–74. 76. Kurokawa, M., Kato, T., Masuko-Hongo, K., Ueda, S., Kobata, T., Okubo, M., Nishimaki, T., Akaza, T., Yoshino, S., Kasakawa, R., Nishioka, K., and Yamamoto, K. (1999). Characterization of T cell clonotypes that accumulated in multiple joints of patients with rheumatoid arthritis. Ann. Rheum. Dis. 58, 546–553. 77. Mima, T., Ohshima, S., Sasai, M., Nishioka, K., Shimizu, M., Murata, N., Yasunami, R., Matsuno, H., Suemura, M., Kishimoto, T., and Saeki, Y. (1999). Dominant and shared T cell receptor beta chain variable regions of T cells inducing synovial hyperplasia in rheumatoid arthritis. Biochem. Biophys. Res. Commun. 263, 172–180. 78. Saal, J. G., Krimmel, M., Steidle, M., Gerneth, F., Wagner, S., Fritz, P., Koch, S., Zacher, J., Sell, S., Einsele, H., and Muller, C. A. (1999). Synovial Epstein-Barr virus infection increases the risk of rheumatoid arthritis in individuals with the shared HLA-DR4 epitope. Arthritis Rheum. 42, 1485–1496.

Michel Neidhart ET AL. 79. Toussirot, E., Auger, I., Roudier, C., Luka, J., Wendling, D., Tiberghien, P., and Roudier, J. (1999). HLA-DR polymorphism influences T-cell precursor frequencies to Epstein-Barr virus (EBV) gp110: implications for the association of HLA-DR antigens with rheumatoid arthritis. Tissue Antigens 54, 146–152. 80. Radstake, T. R., Franke, B., Hanssen, S., Netea, M. G., Welsing, P., Barrera, P., Joosten, L. A., van Riel, P. L., and van den Berg, W. B. (2004). The Toll-like receptor 4 Asp299Gly functional variant is associated with decreased rheumatoid arthritis disease susceptibility but does not influence disease severity and/or outcome. Arthritis Rheum. 50, 999–1001. 81. Kyburz, D., and Gay, S. (2003). Toll-like receptors direct antimicrobial immune responses – and driving arthritis? Curr. Rheumatol. Rep. 5, 407–408. 82. Seibl, R., Kyburz, D., Lauener, R. P., and Gay, S. (2004). Pattern recognition receptors and their involvement in the pathogenesis of arthritis. Curr. Opin. Rheumatol. 16, 411–418. 83. Gonzalez-Escribano, M. F., Rodriguez, R., Valenzuela, A., Garcia, A., Garcia-Lozano, J. R., and Nunez-Roldan, A. (1999). CTLA4 polymorphisms in Spanish patients with rheumatoid arthritis. Tissue Antigens 53, 296–300. 84. Seidl, C., Donner, H., Fischer, B., Usadel, K. H., Seifried, E., Kaltwasser, J. P., and Badenhoop, K. (1998). CTLA4 codon 17 dimorphism in patients with rheumatoid arthritis. Tissue Antigens 51, 62–66. 85. Yen, J. H., Tsai, W. C., Chen, C. J., Ou, T. T., Lin, C. H., Hu, C. J., and Liu, H. W. (2003). Tumor necrosis factor receptor 2 microsatellite and exon 6 polymorphisms in rheumatoid arthritis in Taiwan. J. Rheumatol. 30, 438–442. 86. Dieude, P., Petit, E., Cailleau-Moindrault, S., Osorio, J., Pierlot, C., Martinez, M., Faure, S., Alibert, O., Lasbleiz, S., De Toma, C., Bardin, T., Prum, B., Cornelis, F., and European Consortium on Rheumatoid Arthritis Families (2002). Association between tumor necrosis factor receptor II and familial, but not sporadic, rheumatoid arthritis: evidence for genetic heterogeneity. Arthritis Rheum. 46, 2039–2044. 87. Cantagrel, A., Navaux, F., Loubet-Lescoulie, P., Nourhashemi, F., Enault, G., Abbal, M., Constantin, A., Laroche, M., and Mazieres, B. (1999). Interleukin-1 beta, interleukin-1 receptor antagonist, interleukin-4, and interleukin-10 gene polymorphisms: relationship to occurrence and severity of rheumatoid arthritis. Arthritis Rheum. 42, 1093–1100. 88. Buchs, N., di Giovine, F. S., Silvestri, T., Vannier, E., Duff, G. W., and Miossec, P. (2001). IL-1B and IL-1Ra gene polymorphisms and disease severity in rheumatoid arthritis: interaction with their plasma levels. Genes Immun. 2, 222–228. 89. Genevay, S., Di Giovine, F. S., Perneger, T. V., Silvestri, T., Stingelin, S., Duff, G., and Guerne, P. A. (2002). Association of interleukin-4 and interleukin-1B gene variants with Larsen score progression in rheumatoid arthritis. Arthritis Rheum. 47, 303–309. 90. Jouvenne, P., Chaudhary, A., Buchs, N., Giovine, F. S., Duff, G. W., and Miosse, P. (1999). Possible genetic association between interleukin-1 alpha gene polymorphism and the severity of chronic polyarthritis. Eur. Cytokine Netw. 10, 33–36. 91. Perrier, S., Coussediere, C., Dubost, J. J., Albuisson, E., and Sauvezie, B. (1998). IL-1 receptor antagonist (IL-1RA) gene polymorphism in Sjogren’s syndrome and rheumatoid arthritis. Clin. Immunol. Immunopathol. 87, 309–313. 92. Buchs, N., Silvestri, T., di Giovine, F. S., Chabaud, M., Vannier, E., Duff, G. W., and Miossec, P. (2000). IL-4 VNTR gene polymorphism in chronic polyarthritis. The rare allele is associated with protection against destruction. Rheumatology 39, 1126–1131. 93. Keystone, E., Wherry, J., and Grint, P. (1998). IL-10 as a therapeutic strategy in the treatment of rheumatoid arthritis. Rheum. Dis. Clin. North Am. 24, 629–639. 94. Hajeer, A. H., Lazarus, M., Turner, D., Mageed, R. A., Vencovsky, J., Sinnott, P., Hutchinson, I. V., and Ollier, W. E. (1998). IL-10 gene

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95.

96.

97.

98.

99.

100.

101.

102.

103.

104.

105.

106.

107

108.

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promoter polymorphisms in rheumatoid arthritis. Scand. J. Rheumatol. 27, 142–145. Coakley, G., Mok, C. C., Hajeer, A. H., Ollier, W. E., Turner, D., Sinnott, P. J., Hutchinson, I. V., Panayi, G. S., and Lanchbury, J. S. (1998). Interleukin-10 promoter polymorphisms in rheumatoid arthritis and Felty’s syndrome. Br. J. Rheumatol. 37, 988–991. Lard, L. R., van Gaalen, F. A., Schonkeren, J. J., Pieterman, E. J., Stoeken, G., Vos, K., Nelissen, R. G., Westendorp, R. G., Hoeben, R. C., Breedveld, F. C., Toes, R. E., and Huizinga, T. W. (2003). Association of the -2849 interleukin-10 promoter polymorphism with autoantibody production and joint destruction in rheumatoid arthritis. Arthritis Rheum. 48, 1841–1848. Qin, S., Rottman, J. B., Myers, P., Kassam, N., Weinblatt, M., Loetscher, M., Koch, A. E., Moser, B., and Mackay, C. R. (1998). The chemokine receptors CXCR3 and CCR5 mark subsets of T cells associated with certain inflammatory reactions. J. Clin. Invest. 101, 746–754. Garred, P., Madsen, H. O., Petersen, J., Marquart, H., Hansen, T. M., Freiesleben Sorensen, S., Volck, B., Svejgaard, A., and Andersen, V. (1998). CC chemokine receptor 5 polymorphism in rheumatoid arthritis. J. Rheumatol. 25, 1462–1465. Gomez-Reino, J. J., Pablos, J. L., Carreira, P. E., Santiago, B., Serrano, L., Vicario, J. L., Balsa, A., Figueroa, M., and de Juan, M. D. (1999). Association of rheumatoid arthritis with a functional chemokine receptor, CCR5. Arthritis Rheum. 42, 989–992. Matthey, D. L., Hassell, A. B., Plant, M. J., Dawes, P. T., Ollier, W. R., Jones, P. W., Fryer, A. A., Alldersea, J. E., and Strange, R. C. (1999c). Association of polymorphism in glutathione S-transferase loci with susceptibility and outcome in rheumatoid arthritis: comparison with the shared epitope. Ann. Rheum. Dis. 58, 164–168. Tak, P. P., Zvaifler, N. J., Green, D. R., and Firestein, G. S. (2000). Rheumatoid arthritis and p53: how oxidative stress might alter the course of inflammatory diseases. Immunol. Today 21, 78–82. Yamanishi, Y., Boyle, D. L., Rosengren, S., Green, D. R., Zvaifler, N. J., and Firestein, G. S. (2002). Regional analysis of p53 mutations in rheumatoid arthritis synovium. Proc. Natl Acad. Sci. USA 99, 10025–10030. Huang, J., Ushiyama, T., Inoue, K., Mori, K., and Hukuda, S. (1999). Possible association of CYP17 gene polymorphisms with the onset of rheumatoid arthritis. Clin. Exp. Rheumatol. 17, 721–724. Ushiyama, T., Mori, K., Inoue, K., Huang, J., Nishioka, J., and Hukuda, S. (1999). Association of estrogen receptor polymorphisms with age at onset of rheumatoid arthritis. Ann. Rheum. Dis. 58, 7–10. Kawasaki, T., Ushiyama, T., Ueyama, H., Inoue, K., Mori, K., Ohkubo, I., and Hukuda, S. (1999). Polymorphic CAG repeats of the androgen receptor gene and rheumatoid arthritis. Ann. Rheum. Dis. 58, 500–502. Uhlar, C. M., and Whitehead, A. S. (1999). Serum amyloid A, the major vertebrate acute-phase reactant. Eur. J. Biochem. 265, 501–523. Moriguchi, M., Terai, C., Koseki, Y., Uesato, M., Nakajima, A., Inada, S., Nishinarita, M., Uchida, S., Nakajima, A., Kim, S. Y., Chen, C. L., and Kamatani, N. (1999). Influence of genotypes at SAA1 and SAA2 loci on the development and the length of latent period of secondary AA-amyloidosis in patients with rheumatoid arthritis. Hum. Genet. 105, 360–366. Yamada, T., Okuda, Y., Takasugi, K., Wang, L., Marks, D., Benson, M. D., and Kluve-Beckerman, B. (2003). An allele of serum amyloid A1 associated with amyloidosis in both Japanese and Caucasians. Amyloid 10, 7–11. Gronback, K., Straten, P. T., Ralfkiaer, E., Ahrenkiel, V., Andersen, M. K., Hansen, N. E., Zeuthen, J., Hou-Jensen, K., and Guldberg, P. (1998). Somatic Fas mutations in non-Hodgkin’s lymphoma: association with extranodal disease and autoimmunity. Blood 92, 3018–3024.

609 110. Huang, Q. R., Danis, V., Lassere, M., Edmonds, J., and Manolios, N. (1999). Evaluation of a new Apo-1/Fas promoter polymorphism in rheumatoid arthritis and systemic lupus erythematosus patients. Rheumatology 38, 645–651. 111. Coakley, G., Manolios, N., Loughran, T. P. Jr., Panayi, G. S., and Lanchbury, J. S. (1999). A Fas promoter polymorphism at position –670 in the enhancer region does not confer susceptibility to Felty’s and large granular lymphocyte syndromes. Rheumatology 38, 883–886. 112. Wu, J., Zhou, T., He, J., and Mountz, J. D. (1993). Autoimmune disease in mice due to integration of an endogenous retrovirus in an apoptosis gene. J. Exp. Med. 1783, 461–468. 113. Lin, S. C., Yen, J. H., Tsai, J. J., Tsai, W. C., Ou, T. T., Liu, H. W., and Chen, C. J. (2004). Association of a programmed death 1 gene polymorphism with the development of rheumatoid arthritis, but not systemic lupus erythematosus. Arthritis Rheum. 50, 770–775. 114. Hermann, M., Neidhart, M., Kalden, J. R., and Gay, S. (1998). Retrovirus-associated rheumatic syndromes. Curr. Opin. Rheumatol. 10, 347–354. 115. Di Giovine, F. S., Bailly, S., Bootman, J., Almond, N., and Duff, G. W. (1994). Absence of lentiviral and human T cell leukemia viral sequences in patients with rheumatoid arthritis. Arthritis Rheum 37, 349–358. 116. Seemayer, C. A., Kolb, S. A., Neidhart, M., Ohshima, S., Gay, R. E., Michel, B. A., Gay, S., Böni, J., and Schüpbach, J. (2002). Absence of inducible retroviruses from synovial fibroblasts and synovial fluid cells of patients with rheumatoid arthritis. Arthritis Rheum 46, 2811–2813. 117. Becker, Y. (1995). Endogenous retroviruses in the human genome – a point of view. Virus Genes 9, 211–218. 118. Hermann, M., Hagenhofer, M., and Kalden, J. R. (1996). Retroviruses and systemic lupus erythematosus. Immunol. Rev. 152, 145–156. 119. Perron, H., Garson, J. A., Bedin, F., Beseme, F., Paranhos-Barccala, G., Komurian-Pradel, F., Mallet, F., Tuki, P. W., Voisset, C., Blond, J. L., Lalande, B., Seigneurin, J. M., Mandrand, B., and Collaborative Research Group on Multiple Sclerosis. (1997). Molecular identification of a novel retrovirus repeatedly isolated from patients with multiple sclerosis. Proc. Natl Acad. Sci. USA 94, 7583–7588. 120. Conrad, J. M., Weissmahr, R. N., Bohni, J., Arcari, R., Schupbach, J., and Mach, B. (1997). A human endogenous retroviral superantigen as candidate autoimmune gene in type I diabetes. Cell 90, 303–313. 121. Stransky, G., Vernon, J., Aicher, W. K., Moreland, L. W., Gay, R. E., and Gay, S. (1993). Virus-like particles in synovial fluids from patients with rheumatoid arthritis. Br. J. Rheumatol. 32, 1044–1048. 122. Müller-Ladner, U., Gay, R. E., and Gay, S. (1998). Retroviral sequences in rheumatoid arthritis synovium. Intern. Rev. Immunol. 17, 273–290. 123. Nakagawa, K., Brusic, V., McColl, G., and Harrison, L. C. (1997). Direct evidence for the expression of multiple endogenous retroviruses in the synovial compartment in rheumatoid arthritis. Arthritis Rheum. 40, 627–638. 124. Neidhart, M., Rethage, J., Gay, R. E., and Gay, S. (1999). L1 retrotransposons in rheumatoid arthritis are related to genomic DNA hypomethylation and affect gene expression. Arthritis Rheum. 42 (Suppl): S248. 125. Ali, M., Veale, D. J., Reece, R. J., Quinn, M., Henshaw, K., Zanders, E. D., Markham, A. F., Emery, P., and Isaacs, J. D. (2003). Overexpression of transcripts containing LINE-1 in the synovia of patients with rheumatoid arthritis. Ann. Rheum. Dis. 62, 663–666. 126. Kuchen, S., Seemayer, C. A., Rethage, J., von Knoch, R., Kuenzler, P., Michel, B. A., Gay, R. E., Gay, S., and Neidhart, M. (2004). The L1 retroelement-related p40 protein induces p38delta MAP kinase. Autoimmunity 37, 57–66. 127. Moore, T. L. (1999). Immunopathogenesis of juvenile rheumatoid arthritis. Curr. Opin. Rheumatol. 11, 377–383. 128. Ogilvie, E. M., Fife, M. S., Thompson, S. D., Twine, N., Tsoras, M., Moroldo, M., Fisher, S. A., Lewis, C. M., Prieur, A. M., Glass, D. N.,

610 and Woo, P. (2003). The -174G allele of the interleukin-6 gene confers susceptibility to systemic arthritis in children: a multicenter study using simplex and multiplex juvenile idiopathic arthritis families. Arthritis Rheum. 48, 3202–3206. 129. Crawley, E., Kay, R., Sillibourne, J., Patel, P., Hutchinson, I., and Woo, P. (1999). Polymorphic haplotypes of interleukin-10 5¢ flanking region determine variable interleukin-10 transcription and are associated with particular phenotypes of juvenile rheumatoid arthritis. Arthritis Rheum. 42, 1101–1108. 130. Rosen, P., Hopkin, R. J., Glass, D. N., and Graham, T. B. (2004). Another patient with chromosome 18 deletion syndrome and juvenile rheumatoid arthritis. J. Rheumatol. 31, 998–1000.

Michel Neidhart ET AL. 131. Flato, B., Smerdel, A., Johnston, V., Lien, G., Dale, K., Vinje, O., Egeland, T., Sorskaar, D., and Forre, O. (2002). The influence of patient characteristics, disease variables, and HLA alleles on the development of radiographically evident sacroiliitis in juvenile idiopathic arthritis. Arthritis Rheum. 46, 986–994. 132. Roberts-Thomson, P. J., Jones, M. E., Walker, J. G., Macfarlane, J. G., Smith, M. D., and Ahern, M. J. (2002). Stochastic processes in the causation of rheumatic disease. J. Rheumatol. 29, 2628–2634.

Chapter 37

Laboratory Assessment of Postmenopausal Osteoporosis Patrick Garnero, PhD

Research Scientist, INSERM Research Unit 403 and Vice-President Synarc Molecular Marker Division, Lyon, France

and Pierre D. Delmas, MD, PhD

Professor of Medicine, Université Claude Bernard, Lyon, France and Director INSERM Research Unit 403, Lyon, France

I. Introduction II. Postmenopausal Bone Loss III. Management of Postmenopausal Osteoporosis

IV. Conclusion References

I. INTRODUCTION

forearm fracture. The lifetime risk of any fracture of the hip, spine, and forearm is almost 40% in white women from age 50 years onward [2]. The definition of osteoporosis captures the notion that a low bone mass is by far the most important risk factor for osteoporosis fractures, although other factors must also be taken into account, including age per se, familial and personal history of fracture, some ill-defined structural parameters which influence the fragility of bone architecture, hip axis length, and some factors unrelated to bone such as the propensity to fall in the elderly. Nevertheless, it is only bone mass assessed by bone mineral density (BMD) that forms the current basis for the diagnosis of osteoporosis. BMD in late postmenopausal women, at the time when fractures occur, depends upon the peak bone mass achieved in adolescence and the subsequent rate of bone loss. Women show an accelerated phase of bone loss after the menopause which occurs for about 5–8 years followed by a sustained but somewhat smaller bone loss throughout life. Estrogen deficiency is the main determinant of early postmenopausal bone loss but there is good

The internationally agreed definition of osteoporosis is: “a progressive systemic skeletal disease characterized by low bone mass and microarchitectural deterioration of bone tissue, with a consequent increase in bone fragility and susceptibility to fracture” [1]. Osteoporosis is one of the most prevalent diseases associated with aging and, because of its cost, it is viewed as a major health problem. The most common fractures include vertebral compression fractures, fractures of the distal radius and of the proximal femur (hip fracture). In addition osteoporotic patients commonly sustain fractures at several other sites including the pelvis, proximal humerus, distal femur, tibia, and ribs. Osteoporotic fractures occurring at the spine and the forearm are associated with significant morbidity, but the most serious consequences arise in patients with hip fracture, which is associated with a significant increase in mortality (15–20%), particularly in elderly men and women. In white women, the lifetime risk of hip fracture is 19%, 15.6% for vertebral fracture, and 16% for distal Dynamics of Bone and Cartilage Metabolism

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evidence that it also plays a role in bone loss occurring in the elderly. Other age-related changes in hormonal status, such as vitamin D deficiency and secondary hyperparathyroidism, are likely to play a role in bone loss in the elderly, both in men and women [3, 4]. The prognostic assessment of fracture risk – which is important to optimize therapeutic strategies – is mainly based on the quantification of BMD and in some cases on the rate of bone loss that can be assessed indirectly by measurement of biochemical markers of bone turnover. Some laboratory tests, such as measurements of plasma PTH and 25 hydroxyvitamin D, may be useful in the differential diagnosis of osteoporosis, while other tests, such as assessment of hormones, cytokine and growth factors productions, may provide insights into the pathogenesis of the disease. BMD measurements and biochemical markers of bone turnover, which have greatly improved in recent years, appear to be key methods to diagnose osteoporosis, predict future fracture, and monitor therapeutic regimens. In this paper we will discuss the use of diagnostic techniques – including BMD and bone turnover markers – in postmenopausal osteoporosis.

II. POSTMENOPAUSAL BONE LOSS A. Pattern of Bone Loss Osteoporosis is much more prevalent in women than in men, because of the major role that estrogen deficiency plays in the pathogenesis of bone loss. Numerous studies have shown that among women there is an accelerated bone loss that coincides temporally with menopause [5–7]. Although osteoporosis is a systemic disease affecting the whole skeleton (with the exception of the bones of the skull and face), the pattern of bone loss differs slightly according to skeletal envelopes. Because of a much slower rate of bone remodeling within cortical compared to trabecular envelopes, cortical bone loss may start later than cancellous loss, but the total amount of cortical bone loss in women in their eighties is probably similar to that of cancellous bone, i.e. from 30–50% of the peak bone mass achieved at adulthood. The loss of bone after the menopause follows an exponential pattern [8–11] and varies markedly from one woman to another [12–14]. After the accelerated rate of bone loss lasting the 5–8 years following menopause, the rate of bone loss decreases linearly to an average of about 0.5–1%/year and may accelerate again after the age of 75 years, at least at the hip [15]. The proportion of the overall bone loss related to menopause and estrogenindependent aging processed is still debated, but clearly bone loss in the elderly is still under the influence of estrogen deficiency [16].

B. Sex Hormone and Postmenopausal Bone Loss Of the many factors which influence bone loss in postmenopausal women, sex hormone deficiency is by far the most important. This central role in the pathogenesis of postmenopausal bone loss is indeed strongly supported by the higher prevalence of osteoporosis in women than in men, the increase in bone resorption following blockage of estrogen synthesis by aromatase inhibitor in postmenopausal women [17], the increase in the rate of bone loss after artificial or natural menopause [18], the existence of a relationship between estrogen levels and rates of bone loss [19] and the protective effect of estrogen replacement with respect to bone loss and fracture incidence [20–27]. The mechanism by which estrogen prevents bone loss is mainly related to its ability to block resorption, although stimulation of bone formation may also play a contributory role [28, 29]. Estrogen-dependent inhibition of bone resorption is due to both decreased osteoclastogenesis and diminished resorptive activity of mature osteoclasts. In addition, estrogen may also promote osteoclast apoptosis. The mechanism of action of estrogen is still controversial. The discovery of estrogen receptors in osteoblasts, their stromal precursors and mature osteoclasts suggest that a direct effect on bone or bone marrow may be involved [30]. However, it is not clear which of these cells are the primary targets for estrogen in vivo. There is evidence that estrogen may mediate its effects indirectly through the stimulation of the release of growth factors such as TGF-β [31] – which has been shown in specific experimental conditions to decrease both osteoclastic resorptive activity and recruitment – and/or the inhibition of the secretion of various potent bone resorbing cytokines such as interleukins (II) 1 and 6, tumor necrosis factor (TNF), GM-CSF, MCSF, and PGE2 [32, 33] (Fig. 1). Several studies have shown that natural and surgical menopause are associated with increased production of IL-1 and TNF from peripheral blood and bone marrow monocyte, with a greater persistence of the increase in osteoporotic patients [33–36]. The culture media of monocytes from postmenopausal women have a higher in vitro bone resorption activity than that of either premenopausal women or estrogen-treated postmenopausal women, suggesting a direct link between increased monocytic production of cytokines and increased postmenopausal bone resorption [37]. Because of the identification of the pivotal role of the RANKL/RANK/ OPG regulatory system in the osteoclast formation and function [reviewed in references 38, 39], these earlier findings need to be re-evaluated. Recently, EghbaliFatourechi et al. [40] used an elegant model to investigate the role of the RANKL/RANK/OPG system in postmenopausal bone loss. They isolated bone marrow mononuclear cells, T lymphocytes, and B lymphocytes

Chapter 37 Laboratory Assessment of Postmenopausal Osteoporosis

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Figure 1

Regulation of osteoclast formation, function, and apoptosis by cytokines produced by bone marrow stromal cells, osteoblasts, monocytes, T cells, and B cells. Stimulatory factors are shown in orange and inhibitory factors in green. The effects of estrogen (E) to enhance (+) and inhibit (−) the factors are shown in red. From Bell, N.H. (2003). RANK ligand and the regulation of skeletal remodeling. J. Clin. Invest. 111: 1120–1222.

from premenopausal women, untreated postmenopausal women, and postmenopausal women treated with estrogen. They found that the levels of RANKL per cell were increased two- to threefold in untreated postmenopausal women compared to premenopausal women and correlated positively with serum and urine markers of bone resorption and negatively with circulating estradiol. These data strongly suggest that up-regulation of RANKL on bone marrow cells is an important determinant of increased bone resorption induced by estrogen deficiency in postmenopausal women. Importantly, however, in the same study there was no difference in circulating levels of RANKL and OPG between premenopausal women, untreated postmenopausal women, and estrogen-treated postmenopausal women, suggesting that the measurement of these cytokines from peripheral blood can not be used as a diagnostic test to quantify the effects of estrogen deficiency on bone turnover (see below). Although estrogen deficiency is one of major determinants of postmenopausal bone loss and skeletal fragility, circulating 17β-estradiol levels explain only a small proportion of interindividual variability of BMD and bone loss [41] and most of the cross-sectional studies have failed so

far to show different serum 17β-estradiol values in fracture cases and in controls. Cummings et al. [42] showed that women 65 years of age and older with undetectable serum estradiol (< 5 pg/ml) had a 2.5-fold increase in the risk of subsequent hip and vertebral fractures as compared with women with detectable serum estradiol, an association that remained significant after adjustment for age, weight, serum estrone, sex-hormone-binding globulin (SHBG), and calcaneal BMD. Such a finding was however not confirmed in another large prospective study (EPIDOS) in women 75 years of age and older in which the women with low estradiol levels were not at increased risk of hip fracture [43]. In contrast, we found that elderly women with serum levels in the highest quartile were moderately protected from fracture with a relative risk of 0.66 compared to the other women. However this protective effect disappeared after adjustment for body weight, suggesting that the association between increased estradiol levels and decreased hip fracture risk is mainly mediated by body weight. The reasons for the discrepancies between the SOF and EPIDOS studies on serum estradiol data are unclear and could be due to differences in body weight (higher in the SOF study), but more probably to difference in age which

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was on average 8 years lower in the SOF study. It seems indeed reasonable to believe that the role of residual secretion of steroids is more likely to play a role in younger postmenopausal women. This hypothesis is supported by results of the OFELY study comprising women with a mean age of 64 years. We found that women with serum estradiol levels in the lowest quartile (< 11 pg/ml) had a relative risk of all osteoporotic fractures of 2.2 compared to the other women, an association that remained significant after adjustment for age and body weight [44]. Melton et al. [45] reported similar findings in postmenopausal women of mean age 68 years and found that the free estradiol index (as determined by the ratio of total estradiol on SHBG), but not total estradiol was predictive of overall fracture risk 20 years later, independently of age and BMD. More recently, it was shown that women participating in the Rotterdam study (68 years old on average) with serum estradiol in the lowest tertile had a 2.1-fold higher risk of vertebral fractures – assessed over the 6.5 years following measurements – compared to the other women, and that this association was also independent of BMD [46]. In the OFELY study, we also found that decreased levels of dehydroepiandrosterone sulfate (DHEAs), which can be metabolized to both active androgens and 17β-estradiol, were also associated with increased fracture risk independently of body weight and estradiol levels [44]. Thus, all together, these data suggest that the influence of the residual secretion of estrogen on the risk of fracture may decrease with advancing age. The increased risk associated with low circulating estradiol levels in younger postmenopausal women does not seem to be mediated by increased bone resorption because in the OFELY study markers of bone resorption were predictive of fracture risk independently of serum estradiol levels. A potential mechanism could be that low estradiol levels are associated with increased osteocyte death [47], a common finding in elderly women with hip fracture [48]. SHBG binds 17β-estradiol and thereby decreases its bioavailability. In the above-described studies, a high serum SHBG was modestly and not consistently across all fracture types associated with an increased risk of osteoporotic fractures. In the SOF and the Rotterdam studies, women with both low serum 17β-estradiol and a high SHBG had a marked increase in hip and vertebral fractures [42, 46].

C. Parathyroid Hormone, Vitamin D and Postmenopausal Bone Loss It is well established that parathyroid hormone (PTH) levels increase with advancing age, mainly after 65 years of age [49]. Aging is characterized by a decrease in calcium

intake and intestinal absorption, by a decrease of vitamin D intake and synthesis and by declining production of 1,25 dihydroxyvitamin D, resulting in secondary hyperparathyroidism. This is likely to increase bone turnover and may play a role in cortical bone loss with age, thus increasing the risk of fracture. However, in a large cohort of healthy postmenopausal women, we found no significant correlation between serum intact PTH and markers of bone turnover in women within 20 years of menopause and only a modest contribution of serum PTH levels to bone turnover variance (r 2 < 0.10) in elderly women over 20 years of menopause [50]. Thus, secondary hyperparathyroidism is probably not a major determinant of increased bone turnover in postmenopausal women and osteoporosis. In a prospective study of the determinants of hip fracture, neither baseline serum 25 hydroxyvitamin D nor PTH levels were predictive of the subsequent risk of hip fracture [51]. These findings were confirmed in women over 65 years of age participating in SOF in which no predictive value of serum PTH and 25-OHD for either hip or vertebral fractures [42] was found. In the same study, women in the lowest quintile for serum 1,25-(OH)2 D (< 23 pg/ml) had a twofold increase in the risk of hip (but not of vertebral) fracture that was no longer significant after adjustment for calcaneal BMD. In the younger women of the OFELY study, we found no significant association between serum levels of 25-OH D and the risk fractures (40% vertebral, 60% nonspine fractures) whereas women in the highest quartile of PTH had a 1.8-fold increased risk [44]. The identification of vitamin D deficiency which is highly prevalent in postmenopausal women [52] and its correction by calcium and low doses of vitamin D in the elderly is however very important [4, 53, 54].

D. Local Cytokines and Growth Factors Histomorphometric studies of iliac crest biopsies suggest that postmenopausal bone loss is due to distinct abnormalities: an imbalance between bone resorption and bone formation within a remodeling unit (due to increased osteoclastic activity and/or decreased osteoblastic activity) and to an increase in the activation frequency, i.e. in the number of remodeling units initiated per unit of time and space. Thus, the small negative bone balance within each remodeling unit is amplified by the overall increase in bone turnover which is mainly due to estrogen deficiency. The imbalance results from abnormalities in the coupling mechanism that links resorption and formation. This coupling process is regulated by several local regulatory factors produced by cells of the microenvironment of the bone remodeling unit as described above. Changes in the

Chapter 37 Laboratory Assessment of Postmenopausal Osteoporosis

local production of these factors and/or abnormal responsiveness of cells to these factors are likely to be responsible for alterations in the coupling mechanism. In addition to the cytokines mentioned above, these local factors include the growth regulatory factors which are stored in bone matrix such as insulin-like growth factors (IGF) I and II, TGF-β, fibroblast growth factors, and bone morphogenetic proteins. It should be recognized, however, that most of the data supporting a role of these factors are derived from animal models and there is little information on humans. Of special interest is the potential role of IGF-I. It has been shown that with aging in humans, there is a decrease in the levels of IGF-I in serum and cortical bone [55, 56]. Serum IGF-I correlated with the spine, femoral neck, and radius BMD and decreased levels have been reported in postmenopausal women with vertebral fractures [57]. Studies of the role of IGF-I are obscured by the fact that its action may be modulated by binding proteins (IGFBP) – at least six different ones have been identified – all of them synthesized by osteoblasts. Some, such as IGFBP5, stimulate bone formation, whereas others are inhibitory, such as IGFBP-4. A recent study analyzed the age-related changes of IGFBP-3 and IGFBP-5 in the cortical bone of iliac crest biopsies from 60 postmenopausal women aged 47–74 years [58]. There was a significant age-related increase in bone content of both IGFBP-3 and IGFBP-5, and levels were negatively correlated with BMD of the lumbar spine and femoral neck. These data suggest that these IGFBPs may be involved in postmenopausal osteoporosis, possibly by regulating the bioavailability of IGFs and thereby their anabolic activity. The involvement of IGFBPs in regulation of bone metabolism is also supported by a recent study showing that transgenic mice over-expressing IGFBP-3 are characterized by increased osteoclast number and bone resorption, and impairment of osteoblast proliferation and reduced bone formation [59]. Another cytokine-like protein which has received attention in the recent years is leptin. Leptin is an adipocytederived protein, produced by the ob gene. Beyond its initial function in regulating appetite and energy expenditure, this hormone appears to play important roles in various biological functions including bone metabolism. In vitro, leptin inhibits differentiation of marrow stromal cells to adipocytes [60], inhibits osteoclast generation – an effect probably mediated by the RANKL/RANK/OPG system, as leptin increases OPG mRNA and protein expression in human peripheral blood mononuclear cells – and consequently may contribute to linkage of formation and resorption in bone [61]. In animals, some studies have shown that leptin stimulates bone growth in leptin deficient ob/ob mice [62], whereas Ducy et al. [63] showed that ob−/ob−

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mice have higher bone mass despite hypogonadism and hypercortisolism. Intracerebroventricular infusion of leptin causes bone loss in these mice, and leptin may regulate bone metabolism via the sympathetic nervous system [64]. In ovariectomized rats administration of leptin prevents the increase in bone turnover [65]. The use of these cytokines and growth factors as a diagnostic test in postmenopausal women is however unclear, because studies relating their circulating levels to bone metabolism have yielded conflicting results. Indeed, an important issue in blood measurements is that they do not always adequately reflect levels in the bone microenvironment. For example, Sheidt-nave et al. [66], reported that increased serum IL-6 levels were associated with accelerated bone loss in early – but not late – postmenopausal women, whereas the opposite, i.e. that high serum IL-6 was protective against postmenopausal bone loss, was reported in another larger study [67]. Whether or not circulating levels of IGF-I and IGFBPs reflect adequately local bone levels remains controversial. Serum IGF-I has been found to decrease with age in postmenopausal women, like bone content, and to correlate with BMD in most studies, but it is likely that circulating IGF-I mainly reflects liver synthesis, which is regulated by different systemic and local factors than IGF-I synthesized by osteoblasts. However, we found that low serum IGF-I was strongly associated with the risk fracture in postmenopausal women, independently of BMD [68]. Although the bone content of IGFBP-3 and IGFBP-5 increases with age [58], their serum levels decrease in postmenopausal women [68, 69]. Similar uncertainties about the relevance of circulating levels have been recently reported for leptin and RANKL/OPG. Significant positive association between serum leptin levels and BMD in postmenopausal women was reported in some studies [70–73] independently of body weight. However, others reported that the association between serum leptin and BMD disappeared after adjustment for body mass index or fat mass [74–77]. In a recent study performed in healthy early postmenopausal women, we found that although the association between serum leptin and BMD at the spine and total hip was explained by body mass index, leptin remained a significant independent predictor of bone resorption, women with levels in the highest quartile having on average urinary CTX levels 38% lower than women in the lowest quartile [76]. The association between circulating leptin and bone metabolism is also obscured by the complex interaction with other hormones such as insulin and estrogens, the sympathic nervous system, and body fat distribution. Yano et al. [78] were the first to assess serum OPG in relation with osteoporosis. They found that serum OPG levels were negatively correlated with BMD at the

616 lumbar spine, femoral neck, and total body and positively correlated with biochemical markers of bone turnover. These findings appeared counterintuitive due to the wellestablished protective effects of OPG on bone, but were interpreted as a compensatory mechanism to prevent bone loss. In contrast, other investigators have reported a weak positive association between serum OPG and BMD and a negative correlation with biochemical markers of bone turnover both in healthy postmenopausal women [79] and in cardiac transplant recipients [80]. Finally, more recent studies failed to demonstrate any significant relationships between serum OPG levels, BMD, bone turnover markers, and the risk of all fractures [81–83] in postmenopausal women. Discrepant findings have also been reported in men, Szulc et al. [84] reporting no association between serum OPG and BMD and a weak negative association with bone resorption markers, whereas Khosla et al. [82] found a negative association with radius BMD and a positive association with bone turnover markers. As discussed above, these conflicting findings may be related in part to the fact that circulating levels does not reflect adequately bone microenvironment concentrations [40]. Additional factors are likely to be involved including the methodology used to measure serum OPG. OPG circulates in monomeric, dimeric, and RANKL-bound forms and most of the assays currently available appear to measure all three forms. In mammalian cells, the dimeric form seems to be secreted in a larger amount than monomeric OPG and is biologically more active [85]. However, it is at present unclear what proportion of circulating OPG is monomeric, dimeric, or bound to RANKL and which is the appropriate fraction to measure. It has also been suggested that the ratio between OPG and RANK-L may be more relevant to measure than OPG alone. Schett et al. [83] have recently measured both OPG and RANK-L in a prospective population-based study of 906 men, pre- and postmenopausal women followed for 10 years. They found a strong relationship between decreased serum RANK-L and increased risk of nontraumatic fractures independently of bone resorption and of calcaneus ultrasound parameters (Table I). OPG was not predictive after adjustment for age. These findings need however to be interpreted with caution because of the low number of nontraumatic fractures (n = 32), the unexpected direction of the association – increased RANKL being associated with lower fracture risk – and the heterogeneity of the population. In summary, although measurements of cytokines and growth factors are very useful for the investigation of the pathophysiology of postmenopausal bone loss, clinical data suggest that their measurements in peripheral blood can not be used as a diagnostic test for the management of individual postmenopausal women.

PATRICK GARNERO AND PIERRE D. DELMAS Table I.

Biochemical Markers of Bone Turnover

Formation • Total and bone-specific alkaline phosphatase (bone ALP) • Osteocalcin (OC) • Type I collagen extension propeptides (PICP, PINP)

Resorption Serum/plasma • Tartrate-resistant acid phosphatase (TRAP, 5b isoenzyme) • N-terminal (S-NTX) and C-terminal (S-CTX) crosslinking telopeptide of type I collagen • C-terminal cross-linking telopeptide of type I collagen generated by MMPs (CTX-MMP) Urine • Pyridinoline (PYD) • Deoxypyridinoline (DPD) • U-NTX • U-CTX • Type I collagen helicoidal peptide 620–633 • Galactosyl-hydroxylysine • Hydroxyproline (Hyp) • Mid-region osteocalcin fragments

The markers with the best performance characteristics in osteoporosis are in bold type.

III. MANAGEMENT OF POSTMENOPAUSAL OSTEOPOROSIS As mentioned previously, the identification of postmenopausal women at risk for osteoporosis relies mainly on the quantification of the amount of bone and on the estimation of rate of bone loss. Both of these parameters can be noninvasively assessed by diagnostic techniques such as dual X-ray absorptiometry and biochemical markers of bone turnover. Bone fragility also depends on the morphology, the architecture as well as on the quality (properties) of the bone matrix that cannot be readily assessed.

A. Measurement of Bone Mineral Density Several cross-sectional studies have shown that BMD – measured by a variety of techniques including single and dual photon absorptiometry, single or dual X-ray absorptiometry, quantitative computed tomography (QCT), radiography absorptiometry and radiogrammetry of the metacarpal – is significantly lower in osteoporotic patients than in controls. The strength of the association has been confirmed by several prospective studies showing that each decrease of 1 standard deviation of BMD measured by DXA is associated with an age-adjusted 50–150% increase in the risk of osteoporotic fractures in

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postmenopausal women, mostly in short-term study [86, 87]. The risk prediction bas been found across various age ranges [88] (but is weaker after 80 years of age) in ambulatory and long-term care residents [89]. Melton et al. [45] analyzed the contribution of BMD to predict fracture risk over a median follow-up of 16 years (up to 21 years) in postmenopausal women. They found that the overall fracture risk including hip, vertebral, and forearm fractures was best predicted by baseline femoral neck BMD, and that the predictive value was similar within the first decade and second decade of follow-up. Measurements of central (spine and hip) and peripheral (forearm, heel, finger), and total BMD with an adequate technique have similar value in predicting the risk of all osteoporotic fractures [90], as recently confirmed in the large prospective National Osteoporosis Risk Assessment (NORA) study [88]. It has also been shown that BMD estimated by digital X-ray radiogrammetry predicts hip, vertebral, and wrist fracture in elderly women involved in the SOF study and follow-up for an average of 3.7 years, with similar predictive value as BMD of distal forearm, calcaneus, and spine [91]. If the main concern is to detect the risk of hip fracture, measurement of hip BMD is preferable. Prediction of intertrochanteric fractures is improved by measuring the trochanteric area; prediction of femoral neck fractures is increased by measuring the femoral neck and further enhanced when measurement is limited to the upper half of the femoral neck [92, 93]. Although highly precise, recent studies indicate that assessment of BMD by DXA in vivo is characterized by a significant inaccuracy that may exceed 20–50% at relevant clinical sites such as the lumbar spine, especially in postmenopausal women with osteoporosis and in elderly patient [94–96]. This inaccuracy, which cannot be avoidable, is related to the so called twocomponent limitation of DXA, while body is composed of three main tissues, i.e. bone, muscle, and fat. This limitation is actually of clinical relevance as DXA may yield to artifactual lower T-scores in subjects of lower stature. Some of the limitation can be overcome by new generation of peripheral QCT (pQCT), which provides with good precision, separate assessment of trabecular and cortical bone density, and geometry of bone. A recent study investigating the relationships between BMD assessment with DXA or pQCT of the distal radius and bone strength, as measured in vitro by mechanical tests, showed that the single best predictor of bone strength was pQCT [97]. In addition measures of cortical bone mineral content and bone geometry by pQCT improve significantly the prediction of bone strength. Thus pQCT may prove superior to DXA in the prediction of bone strength and fracture, although large prospective studies will be necessary to confirm this hypothesis.

B. Measurement of Bone Turnover with Biochemical Markers Postmenopausal bone loss results both from an imbalance between resorption and formation at the level of a bone remodeling unit and from an increase in the activation frequency, i.e. in the number of remodeling units initiated per unit of time and space. As reviewed previously the increase in activation frequency after the menopause is mainly dependent on estrogen deficiency, while the remodeling imbalance might be related to a combination of agerelated factors, such as a decrease in IGF-I production. Biochemical markers, whose technical aspects are reviewed in detail elsewhere, reflect the degree of increase in the overall skeletal turnover and cannot be used to give insights in the mechanism of imbalance at the level of a bone remodeling unit. At present, the most sensitive markers for bone formation are serum osteocalcin, bone alkaline phosphatase, and procollagen type I N-terminal propeptide (PINP) (Table I). For the evaluation of bone resorption, immunological assays of pyridinium cross-links of collagen have superseded total pyridinoline assay by high-performance liquid chromatography. Immunological assays are now available for pyridinoline and deoxypyridinoline (DPD) in urine and for C-terminal and N-terminal type I collagen peptides (CTX and NTX, respectively) in serum or urine. More recently new immunoassays for serum tartrate-resistant acid phosphate (TRACP) have been developed, including ones which preferentially detect the isoenzyme 5b, which is predominantly expressed by the osteoclast. In contrast to type I collagen-related markers, TRACP5b isoenzyme is likely to represent mainly the osteoclast number and activity and not directly the rate of bone matrix degradation, although in osteoporosis TRACP5b was found to be correlated with NTX and DPD.

C. Estimation of the Rate of Postmenopausal Bone Loss Several cross-sectional studies indicate that bone turnover increases rapidly after the menopause and this increase in both bone formation and bone resorption is sustained long after the menopause (up to 40 years) [98]. BMD measured at various skeletal sites correlated negatively with bone turnover assessed by various markers in postmenopausal women. We have shown that the correlation between bone markers and BMD becomes much stronger with advancing age, so that in women more than 30 years after the menopause bone turnover accounts for 40–50% of the variance of bone mineral density of the

618 whole skeleton [98]. These cross-sectional data suggest that a sustained increase of bone turnover in postmenopausal women induces a faster bone loss and therefore an increased risk of osteoporosis. Longitudinal studies, which are necessary to avoid potential confounding factors, suffer however from methodological issues: indeed, when bone loss is assessed by annual measurement of BMD at the spine, hip, or radius over 2–4 years, the amount of bone loss is of the same order of magnitude as the precision error of repeated measurements in a single individual, i.e. 3–4%. This technical limitation precludes a valid assessment of the relationship between the rate of bone turnover and the subsequent rate of bone loss in individual postmenopausal women, and probably explains some of the conflicting results that have been published and recently extensively reviewed [99]. Indeed, the association between baseline levels of bone markers and rate of bone loss is more consistent and stronger when bone loss is measured at a very precise site such as radius [100, 101] than when measured at the spine or hip [102]. When the precision error on the rate of bone loss is reduced by performing nine measurements over 24 months, the correlation coefficient bone markers tends to improve from about −0.3 to −0.8 [100]. In a cohort of 305 postmenopausal women aged 50–88 years (mean 64 years) who had annual radius BMD measurements over 4 years, we have investigated the predictive value of baseline values of bone turnover markers [101]. Serum and urinary CTX and urinary NTX for assessing bone resorption, and serum osteocalcin and PINP for assessing bone formation, were found to be highly correlated with the rate of bone loss. In addition, in women within 5 years of menopause who had the highest rate of bone loss, correction of the observed correlation coefficients by errors on bone loss and bone marker measurements resulted in a marked increase in the predictive value of bone markers [99]. Women with baseline values of bone turnover above the premenopausal range had a rate of bone loss four- to sixfold higher than women with a low turnover. Ultimately, the weight of evidence will come from long-term studies. A retrospective study [103] performed over 13 years in women (mean age at baseline: 62 years) in whom calcaneal BMD loss was assessed by eight measurements, showed that 1 standard deviation increase in new specific bone markers such as osteocalcin, bone-specific alkaline phosphatase, and free pyridinoline crosslinks was associated with a twofold increased risk of rapid bone loss defined as the upper tertile of rate of loss. There is currently only one prospective longitudinal long-term study in which the assessment of bone turnover with nonspecific markers (serum alkaline phosphatase, fasting urinary calcium, and hydroxyproline) at the time

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of menopause was correlated with the spontaneous rate of bone loss over the next 12 years. This study showed that early postmenopausal women classified as fast losers based on these markers had lost 50% more bone 12 years later than those diagnosed as slow losers [104]. In summary, with the current performance of bone marker and bone loss measurements, it appears that a single measurement of biochemical markers of bone turnover can not predict the absolute rate of bone loss in an individual woman. However, clearly increased levels of bone markers in postmenopausal women can be regarded as a risk factor for rapid bone loss in the subsequent years.

D. Biochemical Markers of Bone Turnover and Fracture Risk With the emergence of effective – but rather expensive – treatments, it is essential to detect those women at higher risk of fracture. Indeed, although several prospective studies have clearly demonstrated a strong association between BMD measurements and the risk of hip, spine and forearm fractures, recent prospective studies indicate that half of patients with incident fractures have baseline BMD assessed by DXA above the diagnostic threshold of osteoporosis defined as a T-score of −2.5 SD or more below the average value of young healthy women [88, 105, 106] (Fig. 2). Clearly there is a need for improvement in the identification of patients at risk for fracture. In addition to age, several other clinical risk factors have been shown to contribute to fracture probability independently of BMD in postmenopausal women, including a family history of hip fracture, prior fragility fractures, low body mass index in some studies, although these data have been generated mainly in elderly populations and for the prediction of hip fracture risk. Relating baseline bone turnover levels with the subsequent risk of osteoporotic fractures is the valid methodology to assess their clinical utility. The previously mentioned study [104] was extended to a 15-year follow-up of 182 women during which 23 women experienced a peripheral fracture and 25 had one or more spinal fractures [107]. The fracture group had a significantly lower BMD than the group without fracture and a higher initial 3 year rate of bone loss after the menopause. Interestingly, bone mass and the rate of bone loss predisposed to fracture to the same extent, with odds ratios of about 2. Women with both initial low bone mass and an increased rate of bone loss had a threefold increase in the risk of fracture compared with the whole population. This study suggests that initial peak bone mass and postmenopausal rate of bone loss are both important determinants of osteoporosis, although

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Figure 2 Percentage of nonvertebral and hip fractures that occurred in men and women from the Rotterdam study with osteoporosis, osteopenia, or normal bone mineral density using gender-specific NHANES reference population. The Rotterdam study included 7806 men and women aged 55 years and older followed for an average of 6.8 years. From Shuit et al. [106].

a predictive value of the rate of hip bone loss and overall fracture was not confirmed in a recently published 20 year follow-up study [45]. More recently, prospective studies have looked at the relationships between the measurement of biochemical markers of bone formation and bone resorption measured before the fracture has occurred and the subsequent risk fractures (reviewed in reference [108]). 1. Markers of Bone Formation

Prospective studies investigating the relationships between bone formation markers and fracture risk have yielded conflicting results. In the large multicentric cohort of elderly women in France (EPIDOS), no significant relationship was found between levels of serum osteocalcin and bone alkaline phosphatase and the risk of hip fracture occurring during a 2-year follow-up [109]. Similar findings were recently reported in the elderly women involved in the Malmö study [110]. In contrast, in two prospective studies performed in younger healthy postmenopausal women (OFELY and Hawai Osteoporosis Study (HOS)), a significant positive association between increased levels of bone alkaline phosphatase and the risk of vertebral and nonvertebral fracture was observed [44, 111]. The differences between the studies may be related to the type of fracture, the age of the population investigated or to the duration of follow-up which was of 22 months in the

EPIDOS study and of 5 years in the OFELY and HOS studies. More recently, we re-assessed the OFELY data [112] after a median of 10 years of follow-up in which we recorded 158 incident fractures in 115 women including 50 vertebral and 108 nonvertebral fractures. Over this long follow-up period we also found a significant association between increased baseline levels of serum osteocalcin, bone alkaline phosphatase, and PINP and the risk of fractures (unpublished data). Melton et al. [45], however, could not find any significant relationship between total alkaline phosphatase levels and serum osteocalcin and the risk of fracture which occurred within the following 20 years. However these markers which were measured 20 years ago were not the most specific ones and this may have resulted in decreased power to detect a significant association. 2. Markers of Bone Resorption

More consistent data have been obtained on the relationship between increased levels of bone resorption markers and fracture risk. Five prospective studies (EPIDOS, Rotterdam, OFELY, HOS, and Malmö) found that bone resorption assessed by urinary or serum CTX, urinary free deoxypyridinoline, or serum TRAP5b above the premenopausal range, were consistently associated with about a twofold higher risk of hip, vertebral, and nonhip

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and nonvertebral fractures over follow-up periods ranging from 1.8–5 years [44, 98, 109–113]. It remains, however, to be investigated whether bone resorption markers would predict fracture risk over periods which exceed 5 years. Because serum CTX levels are markedly affected by food intake [114, 115], an effect which is likely to be mediated mainly by gastrointestinal hormones [116], it should be measured on fasting morning samples to reduce the variability of the measurement. This technical limitation probably explains the lack of significant predictive value of serum CTX levels measured on nonfasting morning serum samples both in the EPIDOS [117] and SOF [118] studies. In positive studies, the odds ratios were not modified after adjusting for potential confounding factors such as mobility status and were only marginally decreased after adjusting for BMD measured by DXA. Thus, the combination of BMD and bone turnover measurement allows the identification of a subgroup of elderly women at much higher risk of hip fracture than those identified by each test alone (Fig. 3). Increased bone resorption was associated with increased fracture risk only for values which exceed a threshold, especially when defined as levels above the upper limit of the premenopausal range, indicating that bone resorption becomes deleterious for bone strength only when it exceeds a normal physiological threshold which appears to be the upper range of premenopausal women. These data also suggest that increased resorption could lead to increased skeletal fragility through two independent mechanisms. First, a prolonged increase of bone turnover will lead after several years to a lower BMD as discussed previously, which is a major determinant of

Figure 3

reduced bone strength. Second, increased bone resorption above the upper limit of the normal range may induce microarchitectural deterioration of bone tissue such as perforation of trabeculae, a major component of bone strength. 3. Post-translational Modifications of Bone Matrix Proteins

Type I collagen molecules, the main organic component of bone matrix, undergo enzymatic and nonenzymatic post-translational modifications and some of them may be involved in bone mechanical properties (Fig. 4). Among the enzymatic modifications, biochemical studies performed on human bone specimens have shown that an overhydroxylation of lysine residues, an over-glycosylation of hydroxylysine, and a reduction in the concentration of nonreducible crosslinks can be associated with reduced bone strength [119–122]. In human vertebral specimens, Banse et al. [123] recently analyzed the content of immature (hydroxylysononorleucine (HLNL) and dihydroxylysinorleucine (DHLNL)) and mature crosslinks (PYD, DPD, and pyrolle). They showed that the ratio of PYD/DPD was significantly associated with the compressive biomechanical properties of the vertebrae independently of BMD, suggesting that type I collagen crosslinks may be a determinant of bone strength. Nonenzymatic modifications of collagen could also play a role in the mechanical properties of bone tissue. For example, Wang et al. [124] showed that the pentosidine concentration of human femoral bone – an index of nonenzymatic advanced glycation end products – increases with age and higher levels are associated with

Combination of bone mineral density (BMD) at the hip assessed by dual X-ray absorptiometry (DXA) and of bone resorption to predict hip fracture risk in elderly women followed prospectively for 2 years: the EPIDOS study. Low BMD was defined according to the WHO guidelines, i.e. by a value lower than 2.5 SD below the young adult mean (T-score ≤2.5). High bone resorption was defined by urinary CTX or free DPD values higher than the upper limit (mean + 2 SD) of the premenopausal range. From Garnero et al. [109].

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Figure 4

Schematic representation of the extracellular post-translational modifications of type I collagen in bone. Type I collagen is constituted by the association in triple helix of two alpha 1 and one alpha 2 chains except of the two ends (N- and C-telopeptides). In bone matrix, type I collagen is subjected to different post-translational modifications: the mature trivalent crosslinks, including pyridinoline (PYD), deoxypyridinoline (DPD), and pyrrole which make bridges between two telopeptides and the helicoidal region of another collagen molecule. These molecules result from the maturation of divalent crosslinking molecules (dihydroxylysinornorleucine, DHLNL and hydroxylysinonorleucine, HLNL) whose synthesis requires an enzymatic process (Lysil oxydase); the advanced glycation end products (AGEs), which are formed by nonenzymatic glycation involving a sugar such as glucose and amino acids such as lysine. Some AGEs, such as pentosine, are crosslinking molecules although their precise location remains to be determined; the nonenzymatic isomerization of aspartic acid (D) occurring in the C-telopetides of alpha 1 chains.

decreased bone strength. This suggests that nonenzymatic glycation of collagen could lead to alterations in the biomechanical properties of bone that may ultimately result in increased skeletal fragility. β isomerization of the aspartate (D) residue of the 1209AHDGGR1214 sequence (CTX) of the C-telopeptide of type I collagen is another nonenzymatic post-translational modification more recently investigated [125]. Histological studies have shown a decreased degree of type I collagen isomerization within the woven pagetic bone, a tissue characterized by increased fragility [126]. Alterations in the degree of bone type I collagen isomerization can be detected in vivo by the differential measurement of native (α) and isomerized (β) CTX fragments in urine. In the OFELY prospective study, we found that the urinary ratio between native and β-isomerized CTX was significantly associated with increased fracture risk independently of both the level of hip BMD and of bone turnover rate measured by serum bone ALP [127, 128] (Table II). In an in vitro model where the extent of crosslinking can be modified, keeping constant the size and the mineral content of bone, we recently demonstrated that changes in the extent of post-translational modifications of type I collagen (e.g. intermolecular crosslinking such as pyridinoline and pentosidine and CTX isomerization) play a role in

Table II. Increased Urinary CTX Ratio as an Independent Predictor of the Risk of Osteoporotic Fractures Relative risk* (95% CI) of fracture for values in the upper quartile All fractures

Nonvertebral fractures only

αL/βL Unadjusted Adjusted for bone ALP Adjusted for femoral neck BMD Adjusted for bone ALP + femoral neck BMD

2.0 (1.2–3.5) 1.8 (1.1–3.2) 1.8 (1.03–3.1) 1.7 (0.95–2.9)

2.5 (1.3–4.6) 2.2 (1.1–4.2) 2.2 (1.2–4.0) 2.0 (1.04–3.8)

αL/αD Unadjusted Adjusted for bone ALP Adjusted for femoral neck BMD Adjusted for bone ALP + femoral neck BMD

1.8 (1.02–3.2) 1.6 (0.92–2.9) 1.7 (0.97–2.9) 1.6 (0.89–2.8)

1.9 (1.01–3.7) 1.7 (0.87–3.3) 1.8 (0.95–3.5) 1.7 (0.86–3.2)

Urinary CTX ratio at baseline

*Adjusted for age, presence of prevalent fracture, and physical activity. Four hundred and eight women participating in the OFELY study were followed prospectively during 6.8 years. 55 nonvertebral fractures and 16 incident vertebral fractures were recorded. The table shows the relative risks of fracture for women with baseline levels of αL/βL CTX and αL/αD CTX in the upper quartile. From Garnero et al. [127], with permission.

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determining the mechanical competence of cortical bone, and especially post-yield energy absorption independently of BMD [129]. These data indicate that the degree of posttranslational modifications of collagen matrix plays an independent role in determining the mechanical competence of cortical bone and the ratio of α/β CTX may provide an in vivo marker of bone collagen quality. The clinical relevance of bone collagen properties as an important independent determinant of skeletal strength has also recently been suggested by two studies investigating the relationships between circulating homocysteine levels and the risk of fracture in elderly women and men. Homocysteinuria is a rare autosomal recessive disease which is characterized by generalized osteoporosis. This finding has been attributed to an inhibition of the formation of collagen crosslinking by high homocysteine levels [130, 131]. McLean et al. [132] and van Meurs et al. [133] found that elderly women and men with plasma levels of homocysteine in the highest quartile had a risk of nonvertebral and hip fracture that was twice the risk in the other individuals (Fig. 5). Interestingly, in the study from the Netherlands [133], the risk was independent of BMD of the spine and total hip. Although this association may be indirectly mediated by estrogen levels or vitamin status as discussed by Dr Raiz [134], a possible explanation could be alteration of collagen crosslinking properties. Osteocalcin is a noncollagenous protein of bone matrix which contains three residues of γ-carboxyglutamic acid (GLA), a vitamin K-dependent amino acid. It was postulated that impaired γ-carboxylation of osteocalcin could

Figure 5

be an index of both vitamin D and vitamin K deficiency in elderly populations. Vitamin K status may be involved in the maintenance of skeletal integrity. Vitamin K2 treatment in postmenopausal women with osteoporosis decreases serum levels of undercarboxylated osteocalcin (ucOC), increases spine BMD, and reduces the risk of fragility fractures [135]. In two prospective studies performed in a cohort of elderly institutionalized women followed for 3 years [136] and in a population of healthy elderly women (EPIDOS study) [137], levels of ucOC above the premenopausal range were associated with a two- to threefold increase in the risk of hip fracture, although total osteocalcin was not predictive. As for markers of bone resorption, the prediction was still significant after adjusting for hip BMD. More recently, it was reported that a decreased ratio between carboxylated and total osteocalcin – which is an index of increased ucOC – was also associated with increased fracture risk in elderly women living at home [138]. The mechanisms relating increased undercarboxylation of osteocalcin and fracture risk is unclear. Serum ucOC [139], but not total osteocalcin, have been found to be associated more strongly with ultrasonic transmitted velocity (which has been suggested to reflect in part changes in bone structure) at the os calcis and tibia than with BMD [139], suggesting that poor quality may explain the fracture risk associated with undercarboxylated osteocalcin, possibly as a result of inadequate vitamin K status [140]. Clearly these findings open new perspectives for the clinical use of bone markers, not only to measure quantitative changes of bone turnover, but also to assess

Cumulative incidence of fracture among subjects with homocysteine levels in the highest age- and sex-specific quartile as compared to all other subjects. The Rotterdam study included 351 women and 211 men aged 55 years and older (mean 70 years). Eighty-three incident nonvertebral and clinical vertebral fractures occurred during an average of 8.1 years follow-up. The LASA study included 663 women and 628 men aged 65 years and older (mean 76 years). Fifty-eight incident nonvertebral and clinical vertebral fractures occurred during an average 2.7 years follow-up. From van Meurs et al. [133].

Chapter 37 Laboratory Assessment of Postmenopausal Osteoporosis

changes of bone quality, an important determinant of bone strength. 4. Combined Assessment of Fracture Risk

In the OFELY prospective cohort we have shown that combination of the strongest single clinical risk factor (history of fracture after the age of 45 years), with a low hip BMD and high levels of bone resorption assessed by urinary CTX, improve the predictive value of a single test with relative risk of fracture increasing from 1.8–2.8 to 5.8 [44]. Similarly, in a nested case-control analysis of the EPIDOS study, we have compared the ability of history of fracture after the age of 45 years, hip BMD, heel broadband ultrasound attenuation, and urinary CTX to predict the risk of hip fracture and we investigated whether a combination of these parameters could improve the predictive value [141]. In the EPIDOS study, the combination of urinary CTX with either hip BMD or heel BUA increases the specificity by 10% with sensitivity similar to hip BMD or heel BUA alone. Such a combined diagnostic approach might be more cost effective than BMD measurement alone, as it results in a lower number of patients to be treated to avoid one hip fracture. If DXA or ultrasound is not available, we found that the combination of a high bone resorption marker and a positive history of any type of fracture gave a predictive value similar to that obtained with

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BMD or heel BUA alone [141]. As discussed by Johnell et al. [142], the use of odds ratio is not ideal for clinical decision making, since the risk may decrease or remain stable with age whereas absolute risk increases. Thus calculating absolute risk such as 10-year probabilities – which depend on knowledge of the fracture and death hazards – is probably more appropriate. Based on the probability of hip fracture in the Swedish population and on the data from the EPIDOS and OFELY studies, it was found that combining urinary CTX with BMD or history of previous fracture results in a 10-year probability of hip fracture that was about 70–100% higher than that associated with low BMD alone, with a similar pattern for the prediction of all fractures in younger postmenopausal women [142] (Fig. 6). The use of bone markers in individual patients may be appropriate in some situations, especially in women who are not detected at risk by BMD measurements. In the OFELY study including 668 postmenopausal women followed prospectively over 10 years, we found that, among the 115 incident fractures, 54 (47%) actually occurred in osteopenic women. Among these women, the combination of bone markers and history of previous fracture was highly predictive of fracture and allowed the detection of 59% of women with incident fracture [112]. Women at high risk of fracture may benefit from therapeutic intervention, especially if risk factors are amenable to bone-specific agents.

Figure 6 Combination of clinical risk factors, bone mineral density, and bone turnover measurements to identify women with the highest risk of fracture. The figure shows the 10-year probability of hip fracture according to age and relative risk. The symbols show the effect of risk factors on fracture probability derived from women aged 65 years (OFELY study) and 80 years (EPIDOS study). The data from the OFELY study are derived from information of all fractures. Low hip BMD was defined as values at 2.5 SD or below the mean of young adults. High urinary CTX corresponds to values above the upper limit of premenopausal women (mean + 2 SD). Reprinted from Johnell et al. [142], with permission from Elsevier.

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Indeed, some BMD-independent risk factors such as falls are particularly important for hip fracture, but may not be modified by pharmacological interventions [143]. Conversely, some studies have shown a positive effect of bisphosphonates and raloxifene on fracture risk in osteopenic women [144, 145]. In addition, a greater reduction in fracture risk has been shown among osteopenic postmenopausal women with high pretreatment levels of bone turnover with alendronate [146]. Thus, bone markers may be used in the assessment of fracture risk in selected cases where BMD and clinical risk factors are not sufficient to make a treatment decision.

IV. CONCLUSION Low peak bone mass and fast rate of postmenopausal bone loss, the two main determinants of postmenopausal osteoporosis, are determined by complex interactions of genetic and environmental factors, which regulate bone cell activity and bone turnover. Peak bone mass is mainly under genetic influence, whereas estrogen deficiency plays the major role in determining postmenopausal bone loss. Decisions for therapeutic intervention are made in individual patients. The level of intervention will depend not only on the severity of bone loss measured by DXA, but also on age, the existence or not of prevalent fractures, associated risk factors, and the benefit/risk ratio of the proposed therapeutic regimen. For intermediate values (neither high nor low) of BMD, the subsequent rate of bone loss is likely to be important in the assessment of risk. A high bone resorption assessed by specific biochemical markers is associated with increased risk of subsequent fracture, both at the spine and hip, with a predictive value similar to that obtained from bone mass assessment. There is some evidence that an integrated strategy using major clinical risk factors (e.g. history of fractures, low body weight), measurement of BMD by DXA, and measurement of bone resorption by urinary or serum biochemical markers and potentially posttranslation modifications of type I collagen or other bone matrix proteins could be useful to detect high-risk patients.

References 1. World Health Organisation. (1994). Assessment of fracture risk and its application to screening for postmenopausal osteoporosis. Technical Report Series 843, WHO, Geneva. 2. Cooper, C., Atkinson, E. J., O’Falon, W. M., and Melton III, L. J. (1992). Incidence of clinically diagnosed vertebral fractures: a population-based study in Rochester Minnesota, 1985–1989. J. Bone Miner. Res., 7, 221–227. 3. Gallagher, J. C., Kinyamu, H. K., Fowler, S. E., Dawson-Hughes, B., Dalsky, G. P., and Sherman, S. S. (1998). Calciotropic hormones and bone markers in the elderly. J. Bone Miner. Res. 13, 475–482.

4. Dawson-Hughes, B., Harris, S. S., Krall, E. A., and Dallal, G. E. (1997). Effect of calcium and vitamin D supplementation on bone density in men and women 65 years of age or older. N. Engl. J. Med. 337, 670–676. 5. Riggs, B. L., Wahner, H. W., Melton, III, L. J., Richelson, L. S., Judd, H. L., and Offord, K. P. (1985). Rates of bone loss in the appendicular and axial skeletons of women: evidence of substantial bone loss prior to menopause. J. Clin. Invest. 77, 1487–1491. 6. Riggs, B. L., Wahner, H. W., Dunn, W. L., Seeman, Offord, K. P., Dunn, W. L., Mazen, R. B., Johnson, K. A., and Melton III, L. J. (1982). Changes in bone mineral density of the proximal femur and spine with aging. J. Clin. Invest. 70, 716–723. 7. Arlot, M. E., Sornay-Rendu, E., Garnero, P., Vey-Marty, B., and Delmas, P. D. (1997). Pre and postmenopausal bone loss evaluated by DXA at different skeletal sites in healthy women: The OFELY cohort. J. Bone Miner. Res. 12, 683–690. 8. Pouilles, J. M., Tremollieres, F., and Ribot, C. (1993). The effects of menopause on longitudinal bone loss from the spine. Calcif. Tissue Int. 52, 340–343. 9. Hui, S. L., Slemenda, W., Johnston, C. C., and Appledorn, C. R. (1987). Effects of age and menopause on vertebral bone density. Bone Miner. 2, 141–146. 10. Ribot, C., Tremollières, F., Pouilles, J. M., Louvet, J. P., and Guiraud, R. (1988). Influence of the menopause and aging on spinal density in French women. Bone Miner. 5, 89–97. 11. Heldlund, L. R., and Gallagher, J. C. (1989). The effect of age and menopause on bone mineral density of the proximal femur. J. Bone Miner. Res. 4, 639–641. 12. Nordin, B. E. C., Need, A. G., Chatterton, B. E., Horowitz, M., and Morris, H. A. (1989). The relative contributions of age and years since menopause to postmenopausal bone loss. J. Clin. Endocrinol. Metab. 70, 83–88. 13. Christiansen, C., Riis, B. J., and Rodbro, P. (1987). Prediction of rapid bone loss in postmenopausal women. Lancet, 1, 1105–1108. 14. Pouillès, J. M., Trémollière, F., and Ribot, C. (1996). Variability of vertebral and femoral postmenopausal bone loss. Osteoporosis Int. 6, 320–324. 15. Jones, G., Nguyen, T., Sambrook, P., Kelly, P. J., and Eisman, J. A. (1994). Progressive loss of bone in the femoral neck in elderly people, longitudinal findings from the Dubbo osteoporosis epidemiology study. Brit. Med. J. 309, 691–695. 16. Prestwood, K. M., Pilbeam, C. C., Burleson, J. A., Woodiel, F. N., Delmas, P. D., Deftos, L. J., and Raisz, L. G. (1994). The short-term effects of conjugated estrogen on bone turnover in older women. J. Clin. Endocrinol. Metab. 79, 366–371. 17. Heshmati, H. M., Khosla, S., Robins, S. P., O’Fallon, M., Melton III, L. J., and Riggs, B. L. (2002). Role of low levels of endogeneous estrogen in regulation of bone resorption in late postmenopausal women. J. Bone Miner. Res. 17, 172–178. 18. Genant, H. K., Cann, C. E., Ettinger, B., and Gordon, G. S. (1982). Quantitative computed tomography of vertebral spongiosa: a sensitive method for detecting early bone loss after oophorectomy. Ann. Intern. Med. 97, 699–705. 19. Slemenda, C., Hui, S. L., Longcope, C., and Johnston, C. C. (1987). Sex steroids and bone mass: a study of changes about the time of menopause. J. Clin. Invest. 80, 1261–1269. 20. Gordan, G. S., Picchi, J., and Roof, B. S. (1973). Antifracture efficacy of long term estrogens for osteoporosis. Trans. Assoc. Am. Physicians 86, 326–332. 21. Ettinger, B., Genant, H. K., and Cann, C. E. (1979). Long term estrogen replacement therapy. I. Metabolic effects. Am. J. Obstet. Gynecol. 133, 525–536. 22. Hutchinson, T. A., Polansky, S. M., and Feinstein, A. R. (1979). Postmenopausal estrogens protect against fractures of hip and distal radius. Lancet 706–710.

Chapter 37 Laboratory Assessment of Postmenopausal Osteoporosis 23. Weiss, N. S., Ure, C. L., Ballard, J. H., Williams, A. R., and Dailing, J. R. (1980). Decreased risk of fractures of the hip and lower forearm with postmenopausal use of estrogens. N. Engl. J. Med. 303, 1195–1198. 24. Johnson, R. E., and Specht, E. E. (1981). The risk of hip fracture in postmenopausal females with and without estrogen drug exposure. Am. J. Public Health 71, 138–144. 25. Kreiger, N., Keysley, J. L., Holford, T. R., and O’Connor, T. (1982). An epidemiologic study of hip fracture in postmenopausal women. Am. J. Epidemiol. 116, 141–148. 26. Williams, A. R., Weiss, N. S., Ure, C. L., Ballard, J., and Dailing, J. (1982). Effect of weight, smoking, and estrogen use on the risk of hip and forearm fractures in postmenopausal women. Obstet. Gynecol. 60, 695–699. 27. Kanis, J. A., Johnell, O., Gulberg, B., Allander, E., Dilsen, G., Gennari, C., Lopes Vaz, A. A., Lyritis, G. P., Mazzuoli, G., and Miravet, L. (1992). Evidence for efficacy of drugs affecting bone metabolism in preventing hip fracture. Brit. Med. J. 305, 1124–1128. 28. Chow, J., Tobias, J. H., Colston, K. W., and Chambers, T. J. (1992). Estrogen maintains trabecular bone volume in rats not only by suppression of bone resorption but also by stimulation of bone formation. J. Clin. Invest. 89, 74–78. 29. Bain, S. D., Bailey, S. C., Celino, D. L., Lantry, M. M., and Edwards, M. W. (1993). High-dose estrogen inhibits bone resorption and stimulates bone formation in the ovariectomized mouse. J. Bone Miner. Res. 8, 435–441. 30. Oursler, M. J., Kassem, M., Turner, R., Riggs, B. L., and Spelsberg, T. C. (1996). Regulation of bone cell function by gonadal steroids. In “Osteoporosis,” (Marcus, R., Feldman, D., Kelsey, J. eds), 237–260. Academic Press, San Diego, CA. 31. Oursler, M. J., Cortese, C., Keeting, P., Anderson, M. A., Bonde, S. K., Riggs, B. L., and Spelsberg, T. C. (1991). Modulation of transforming growth factor-beta production in normal osteoblast-like cells by 17 beta-estradiol and parathyroid hormone. Endocrinology 112, 3313–3320. 32. Riggs, B. L., Khosla, S., and Melton, L. J. (2002). Sex steroids and the construction and conservation of the adult skeltaon. Endocr. Rev. 23, 279–302. 33. Manolagas, S. C. (2000). Birth and death of bone cells: basic regulatory mechanisms and implications for the pathogenesis and treatment of osteoporosis. Endrocr. Rev. 21, 115–137. 34. Pacifici, R., Rifas, L., McCracken, R., Vered, I., McMurtry, C., Avioli, L. V., and Peck, W. A. (1989). Ovarian steroid treatment blocks a postmenopausal increase in blood monocyte interleukine 1 release. Proc. Natl. Acad. Sci. USA 86, 2398–2402. 35. Pacifici, R., Vannice, J. L., Rifas, L., and Kimble, R. B. (1993). Monocytic secretion of interleukine-1 receptor antagonist in normal and osteoporotic women: Effects of menopause and estrogen/progesterone therapy. J. Clin. Endocrinol. Metab. 77, 1135–1141. 36. McKane, W. R., Khosla, S., Peterson, J. M., Egan, K., Riggs, B. L. (1994). Circulating levels of cytokines that modulate bone resorption. Effects of age and menopause in women. J. Bone Miner. Res. 9, 1313–1318. 37. Cohen-Solal, M. E., Craulet, A. M., Denne, M. A., Gueris, D., Baylink, D., and de Vernejoul, M. D. (1993). Peripheral monocyte culture supernatants of menopausal women can induce bone resorption: Involvement of cytokines. J. Clin. Endocrinol. Metab. 77, 1648–1653. 38. Suda, T., Takahashi, N., Udagawa, N., Jimi, E., Gillespie, M. T., Martin, T. J. (1999). Modulation of osteoclast differentiation and function by the new members of the tumor necrosis factor receptor and ligand families. Endocr. Rev. 20, 345–357. 39. Hofbauer, L. C., Khosla, S., Dunstan, C. R., Lacey, D. L., Boyle, W. J., and Riggs, B. L. (2000). The roles of osteoprotegerin and osteoprotegerin ligand in the paracrine regulation of bone resorption. J. Bone Miner. Res. 15, 2–12. 40. Eghbali-Fatourechi, G., Khosla, S., Sanyal, A., Boyle, W. J., Lacey, D. L., and Riggs, B. L. (2003). Role of RANK-ligand in mediating increased bone resorption in early postmenopausal women. J. Clin. Invest. 111, 1221–1230.

625

41. Slemenda, C., Longcope, C., Peacock, M., Hui, S., and Johnston, C. C. (1996). Sex steroids, bone mass, and bone loss. A prospective study of pre,-peri and postmenopausal women. J. Clin. Invest. 97, 14–21. 42. Cummings, S. R., Browner, W. S., Bauer, D., Stone, K., Ensrud, K., Jamal, S., and Ettinger, B. (1998). Endogenous hormones and the risk of hip and vertebral fractures among older women. N. Engl. J. Med. 339, 733–738. 43. Chapurlat, R. D., Garnero, P., Bréart, G., Meunier, P. J., and Delmas, P. D. (2000). Serum estradiol and sex hormone-binding globulin and the risk of hip fracture in elderly women: The EPIDOS study. J. Bone Miner. Res. 15, 1835–1841. 44. Garnero, P., Sornay-Rendu, E., Claustrat, B., and Delmas, P. D. (2000). Biochemical markers of bone turnover, endogenous hormones and the risk of fractures in postmenopausal women: The OFELY study. J. Bone Miner. Res. 15, 1526–1536. 45. Melton, L. J., Crowson, C. S., O’Fallon, W. M., Wahner, H. W., and Riggs, B. L. (2003). Relative contributions of bone density, bone turnover and clinical risk factors to long-term fracture predictions. J. Bone Miner. Res. 18, 312–318. 46. Goderie-Pomp, H. W., van der Klift, M., de Ronde, W., Hofman, A., de Jong, F. H., and Plos, H. A. (2004). Endogeneous sex hormones, sexhormone-binding globulin, and the risk of incident vertebral fractures in elderly men and women: the Rotterdam study. J. Clin. Endocrinol. Metab. 898, 3261–3269. 47. Tomkinson, A., Reeve, J., Shaw, R. W., and Noble, B. S. (1997). The death of osteocytes via apoptosis accompanies estrogen withdrawal in human bone. J. Clin. Endocrinol. Metab. 82, 3128–3135. 48. Dunstan, C. R., Evans, R. A., Hills, E., Wong, S. Y. P., Higgs, R. J. E. D. (1990). Bone death in hip fracture in the elderly. Calcif. Tissue Int. 47, 270–275. 49. Epstein, S., Bryce, G., Hinnan, J. W., Miller, O. N., Riggs, B. L., Hui, S. L., and Johnson, C. C., Jr (1986). The influence of age on bone mineral regulating hormones. Bone 7, 421–425. 50. Garnero, P., Sornay-Rendu, E., Chapuy, M. C., and Delmas, P. D. (1996). Increased bone turnover in late postmenopausal women is a major determinant of osteoporosis. J. Bone Miner. Res. 11, 337–349. 51. Garnero, P., Hausherr, E., Chapuy, M. C., Marcelli, C., Grandjean, H., Muller, C., Cormier, C., Breart, G., Meunier, P. J. and Delmas, P. D. (1996). Markers of bone resorption predict hip fracture in elderly women: The EPIDOS Prospective study. J. Bone Miner. Res. 11, 1531–1538. 52. Mezquita-Raya, P., Munoz-Torres, M., De Dios Luna, J., Luna, V., Lopez-Rodriguez, F., Tores-Vela, E., and Escobar-Jimenez, F. (2001). Relation between vitamin D insufficiency, and bone metabolism in healthy postmenopausal women. J. Bone Miner. Res. 16, 1408–1415. 53. Chapuy, M. C., Arlot, M. E., Duboeuf, F., Brun, J., Crouzet, B., Arnaud, S., Delmas, P. D., and Meunier, P. J. (1992). Vitamin D3 and calcium to prevent hip fractures in elderly women. N. Engl. J. Med. 327, 1637–1642. 54. Passeri, G., Pini, G., Troiano, L., Vescovini, R., Sansoni, P., Passeri, M., Guerisi, P., Delsignore, R., Pedrazzoni, M., and Franceschi, C. (2003). Low vitamin D status, high bone turnover, and bone fractures in centenarians. J. Clin. Endocrinol. Metab. 88, 5109–5115. 55. Hammerman, M. R. (1987). Insuline-like growth factors and aging. Endocrinol. Metab. Clin. North Am. 16, 995–1008. 56. Nicolas, V., Prewett, V., Bettica, P., Mohan, P., Finfleman, R. D., Baylink, D. J., and Farley, J. T. (1994). Age-related decreases in IGF-I and transforming growth factor beta in femoral cortical bone from both men and women: implications for loss in aging. J. Clin. Endocrinol. Metab. 78, 1001–1016. 57. Sugimoto, T., Nishiyama, K., Kuribayashi, F., and Chihara, K. (1997). Serum levels of insuline-like growth factor (IGF) I and IGF-Biding protein (IGFBP)-2 and IGFBP-3 in osteoporotic patients with and without spinal fractures. J. Bone Miner. Res. 12, 1272–1279. 58. Ueland, T., Brixen, K., Mosekilde, L., Fyvbjerg, A., and Bollerslev, J. (2003). Age-related changes in cortical bone content of insulin-like growth factor binding (IGFBP)-3, IGFBP-5, osteoprotegerin and

626

59.

60.

61.

62.

63.

64.

65.

66.

67.

68.

69.

70.

71.

72.

73.

74.

calcium in postmenopausal osteoporosis: a crossectional study. J. Clin. Endocrinol. Metab. 88, 1014–1018. Silha, J. V., Mishra, S., Rosen, C. J., Beamer, W. G., Turner, R. T., Powell, D. R., and Murphy, L. J. (2003). Perturbations in bone formation and resorption in insulin-like growth factor binding protein-3 transgenic mice. J. Bone Miner. Res. 18, 1834–1841. Thomas, T., Gori, F., Khosla, S., Jensen, M. D., Burguera, B., and Riggs, B. L. (1999). Leptin acts on human marrow stromal cells to enhance differentiation to osteoblasts and to inhibit differentiation to adipocytes. Endocrinology 140, 1630–1638. Holloway, W. R., Collier, F. M., Aitken, C. J., Myers, D. E., Hodge, J. M., Malakellis, M., Gough, T. J., Collier, G. R., Nicholson, G. C. (2002). Leptin inhibits osteoclast generation. J. Bone Miner. Res. 17, 200–209. Steppan, C. M., Crawford, D. T., Chidsey-Frink, K. L., Ke, H., Swick, A. G. (2000). Leptin is a potent stimulator of bone growth in ob/ob mice. Regul. Pept. 92, 73–78. Ducy, P., Amling, M., Takeda, S., Priemel, M., Schilling, A. F., Beil, F. T., Shen, J., Vinson, C., Rueger, J. M., and Karsenty, G. (2000). Leptin inhibits bone formation through a hypothalamic relay: A central control of bone mass. Cell 100, 197–207. Takeda, S., Elefteriou, F., Levasseur, R., Liu, X., Zhao, L., Parker, K. L., Armstrong, D., Ducy, P., and Karsenty, G. (2002). Leptin regulates bone formation via the sympathetic nervous system. Cell 111, 305–317. Burguera, B., Hofbauer, L. C., Thomas, T., Gori, F., Evans, G. L., Khosla, S., Riggs, B. L., and Turner, R. T. (2001). Leptin reduces ovariectomy-induced bone loss in rats. Endocrinology 142, 3546–3553. Sheidt-Nave, C., Bismar, H., Leidig-Bruckner, G., Woitge, H., Seibel, M. J., Zeigler, R., and Pfeilschifter, J. (2001). Serum interleukin 8 is a major predictor of bone loss in women specific to the first decade. J. Clin. Endocrinol. Metab. 86, 2032–2042. Abrahamsen, B., Bonnevie-Nielsen, V., Ebbesen, E. N., Gram, J., and Beck-Nielsen, H. (2000). Cytokines and bone loss in a 5-year longitudinal study-hormone replacement therapy suppresses soluble interleukin-6 receptor and increases interleukin-1 receptor antagonist: the Danish osteoporosis prevention study. J. Bone Miner. Res. 15, 1545–1554. Garnero, P., Sornay-Rendu, E., and Delmas, P. D. (2000). Low levels of serum IGF-I and occurrence of osteoporotic fractures in postmenopausal women. Lancet 355, 898–899. Boonen, S., Mohan, S., Dequecker, J., Aerssens, J., Vanderschueren, D., Verbeke, G., Broos, P., Bouillon, R., and Baylink, D. J. (1999). Downregulation of the serum stimulatory components of the insulin-like growth factor (IGF) system (IGF-I, IGF-II, IGF-binding protein (BP)-3 and IGFBP-5) in age-related (type II) femoral neck osteoporosis. J. Bone Miner. Res. 14, 2140–2158. Thomas, T., Burguera, B., Melton, L. J., Atkinson, E. J., O’Fallon, W. M., Riggs, B. L., and Khosla, S. (2001). Role of serum leptin, insulin, and estrogen levels as potential mediators of the relationship between fat mass and bone mineral density in men versus women. Bone 29, 114–120. Yamauchi, M., Sugimoto, T., Yamaguchi, T., Nakaoka, D., Kanzawa, M. Yano S., Ozuru, R., Sugishita, T., and Chihara, K. (2001). Plasma leptin concentrations are associated with bone mineral density and the presence of vertebral fractures in postmenopausal women. Clin. Endocrinol. (Oxf). 55, 341–347. Pasco, J. A., Henry, M. J., Kotowicz, M. A., Collier, G. R., Ball, M. J., Ugoni, A. M., and Nicholson, G. C. (2001). Serum leptin levels are associated with bone mass in nonobese women. J. Clin. Endocrinol. Metab. 86, 1884–1887. Blain, H., Vuillemin, A., Guillemin, F., Durant, R., Hanesse, B., de Talance, N., Doucet, B., and Jeandel, C. (2002). Serum leptin level is a predictor of bone mineral density in postmenopausal women. J. Clin. Endocrinol. Metab. 87, 1030–1035. Martini, G., Valenti, R., Giovani, S., Franci, S., Campagna, S., and Nuti, R. (2001). Influence of insulin-like growth factor and leptin on bone mass in healthy postmenopausal women. Bone 28, 113–117.

PATRICK GARNERO AND PIERRE D. DELMAS 75. Ruhl, C. E., and Everhart, J. E. (2002). Relationship of serum leptin concentration with bone mineral density in the United States population. J. Bone Miner. Res. 17, 1896–1903. 76. Roux, C., Arabi, A., Porcher, R., and Garnero, P. (2003). Serum leptin as a determinant of bone resorption in healthy postmenopausal women. Bone 33, 847–852. 77. Kontogianni, M. D., Dafni, U. G., Routsias, J. G., and Skopouli, F. N. (2004). Blood leptin and adiponectin as possible mediators of the relation between fat mass and BMD in perimenopausal women. J. Bone Miner. Res. 19, 546–551. 78. Yano, K., Tsuda, E., Washida, N., Kobayashi, F., Goto, M., Harada, A. et al. (1999). Immunological characterization of circulating osteoprotegerin/osteoclastogenesis inhibitory factor: increased serum concentrations in postmenopausal with osteoporosis. J. Bone Miner. Res. 14, 518–527. 79. Rogers, A., Saleh, S., Hannon, R. A., Greenfield, D., and Eastell, R. (2002). Circulating estradiol and osteoprotegerin as determinants of bone turnover and bone density in postmenopausal women. J. Clin. Endocrinol. Metab. 87, 4470–4475. 80. Fahrleitner, A., Prenner, G., Leb, G., Tscheliessnigg, K. H., PiswangerSolkner, C., Obermayer-Pietsch, B., Portugaller, H. R., Berghold, A., Dobnig, H. (2003). Serum osteoprotegerin is a major determinant of bone density development and prevalent vertebral fracture status following cardiac transplantation. Bone 32, 96–106. 81. Browner, W. S., Lui, L. Y., and Cummings, S. R. (2001). Associations of serum osteoprotegerin levels, diabetes, stroke, bone density, fractures and mortality in elderly women. J. Clin. Endocrinol. Metab. 86, 631–637. 82. Khosla, S., Arrighi, H. M., Melton, III, L. J., Atkinson, E. J., O’Fallon, W. M., Dunstan, C., and Riggs, B. L. (2002). Correlates of osteoprotegerin levels in women and men. Osteoporos. Int. 13, 394–399. 83. Schett, G., Kiechl, S., Redlich, K., Oberhollenzer, F., Weger, S., Egger, G., Mayr, A., Jocher, J., Xu, Q., Pierschmann, P., Teitelbaum, S., Smolen, J., and Willeit, J. (2004). Soluble RANKL and the risk of non traumatic fracture. JAMA 291, 1108–1113. 84. Szulc, P., Hofbauer, L. C., Heufelder, A. E., Rothe, S., and Delmas, P. D. (2001). Osteoprotegerin serum levels in men: correlation with age, estrogen and testosterone status. J. Clin. Endocrinol. Metab. 86, 3162–3165. 85. Tomoyasu, A., Masaaki, G., Fujisi, N., Mochizuki, S., Yasuda, H., Morinaga, T. et al. (1998). Characterization of monomeric and homodimeric forms of osteoclastogenesis inhibitory factor. Biochem. Biophys. Res. Commun. 245, 382–387. 86. Anonymous. (1993). Consensus development conference: diagnosis, prophylaxis and treatment of osteoporosis. Am. J. Med. 94, 646–650. 87. Marshall, D., Johnell, O., and Wedel, H. (1996). Meta-analysis of how well measures of bone mineral density predict occurrence of osteoporotic fractures. Brit. Med. J. 312, 1254–1259. 88. Miller, P. D., Siris, E. S., Barret-Connor, E., Faulkner, K. G., Wehren, L. E., Abbott, T. A., Chen, Y-T., Berger, M. L., Santora, A. C., and Sherwood, L. M. (2002). Prediction of fracture risk in postmenopausal white women with peripheral bone densitometry: Evidence from the National Osteoporosis Risk Assessment. J. Bone Miner. Res. 17, 2222–2230. 89. Broe, K. D., Hannan, M. T., Kiely, D. K., Cali, C. M., Cupples, L. A., and Kiel, D. P. (2000). Predicting fractures using bone mineral density: a prospective study of long-term care residents. Osteoporos. Int. 11, 765–771. 90. Cummings, S. R., Nevitt, M. C., Browner, W. S., Stone, K., Fox, K. M., Ensrud, K. E., Cauley, J., Black, D., and Vogt, T. M. (1995). Risk factors for hip fracture in white women. N. Engl. J. Med. 332, 767–773. 91. Bouxsein, M. L., Palermo, L., Yeung, C., and Black, D. M. (2002). Digital X-ray radiogrammetry predicts hip, wrist and vertebral fracture risks in elderly women. A prospective analysis from the study of osteoporotic fractures. Osteoporos. Int. 13, 358–365. 92. Schott, A. M., Cormier, C., Hans, D., Favier, F., Hausherr, E., DargentMolina, P., Delmas, P. D., Ribot, C., Sebert, J. L., Breart, G., and

Chapter 37 Laboratory Assessment of Postmenopausal Osteoporosis

93.

94.

95.

96.

97. 98.

99. 100.

101.

102.

103. 104.

105.

106.

107.

108. 109.

110.

Meunier, P. J. (1998). How hip and whole body bone mineral density predict hip fracture in elderly women: the EPIDOS prospective study. Osteoporos. Int. 8, 247–254. Duboeuf, F., Hans, D., Schott, A. M., Kolzki, P. O., Favier, F., Marcelli, C., Meunier, P. J., and Delmas, P. D. (1997). Different morphometric and densitometric parameters predict cervical and trochanteric hip fracture: the EPIDOS study. J. Bone Miner. Res. 12, 1895–1902. Bolotin, H. H., Sievänen, H., Grashuis, J. L., Kuiper, J. W., and Järvinen, T. L. N. (2001). Inaccuracies inherent in patients-specific dual energy X-ray absorptiometry bone mineral density measurements: Comprehensive phantom-based evaluation. J. Bone Miner. Res. 16, 417–426. Bolotin, H. H. (2001). Inaccuracies inherent in dual-energy X-ray absorptiometry in vivo bone mineral densitometry may flaw osteopenic/ osteoporotic interpretations and mislead assessment of antiresorptive effectiveness. Bone 28, 548–555. Bolotin, H. H., Sievänen, H., and Grashuis, J. L. (2003). Patient-specific DXA bone mineral density inaccuracies: Quantitative effects of nonuniform extraosseous fat distributions. J. Bone Miner. Res. 18, 1020–1027. Muller, M. E., Webber, C. E., and Bouxtein, M. L. (2003). Predicting the failure load of the distal radius. Osteoporos. Int. 14, 345–352. Garnero, P., Sornay-Rendu, E., Chapuy, M. C., and Delmas, P. D. (1996). Increased bone turnover in late postmenopausal women is a major determinant of osteoporosis. J. Bone Miner. Res. 11, 337–349. Stepan, J. J. (2000). Prediction of bone loss in postmenopausal women. Osteoporos. Int. Suppl.6, S45–S54. Johansen, J. S., Riis, B. J., Delmas, P. D., and Christiansen, C. (1988). Plasma BGP: an indicator of spontaneous bone loss and effect of estrogen treatment in postmenopausal women. Eur. J. Clin. Invest. 18, 191–195. Garnero, P., Somay Rendu, E., Duboeuf, F., and Delmas, P. D. (1999). Markers of bone turnover predict postmenopausal bone loss over 4 years: The OFELY study. J. Bone Miner. Res. 14, 399–406. Bauer, D. C., Sklarin, P. M., Stone, K. T., Black, D. M., Nevitt, M. C., Ensrud, K., Arnaud, C. D., Genant, H. K., Garnero, P., Delmas, P. D., Lawaetz, H., and Cummings, S. R. (1999). Biochemical markers of bone turnover and prediction of hip bone loss in older women: The study of osteoporotic fractures. J. Bone Miner. Res. 8, 1404–1410. Ross, P. D., and Knowlton, W. (1998). Rapid bone loss is associated with increased levels of biochemical markers. J. Bone Miner. Res. 13, 297–302. Hansen, M. A., Kirsten, O., Riss, B. J., and Christiansen, C. (1991). Role of peak bone mass and bone loss in postmenopausal osteoporosis: 12 years study. Brit. Med. J. 303, 961–964. Stone, K. L., Seeley, D. G., Lui, L. Y., Cauley, J. A., Ensrud, K., Browner, W. S., Nevitt, M. C., and Cummings, S. R. (2003). BMD at multiple sites and risk of fracture of multiple types: long-term results from the Study of Osteoporotic Fractures. J. Bone Miner. Res. 18(11), 1947–1954. Schuit, S. C. E., van der Klift, M., Weel, A. E. A. M., de Laet, C. E. D. H., Buerger, H., Seeman, E., Hofman, A., Uitterlinden, A. G., van Leeuwen, J. P. T. M., and Pols, H. A. P. (2004). Fracture incidence and association with bone mineral density in elderly men and women: the Rotterdam study. Bone 31, 195–202. Riis, S. B. J., Hansen, A. M., Jensen, K., Overgaard, K., and Christiansen, C. (1996). Low bone mass and fast rate of bone loss at menopause-equal risk factors for future fracture. A 15 year follow-up study. Bone 19, 9–12. Garnero, P. (2000). Markers of bone turnover for the prediction of fracture risk. Osteoporos. Int. Suppl 6: S55–S65. Garnero, P., Hausherr, E., Chapuy, M. C., Marcelli, C., Grandjean, H., Muller, C., Cormier, C., Breart, G., Meunier, P. J., and Delmas, P. D. (1996). Markers of bone resorption predict hip fracture in elderly women: the EPIDOS prospective study. J. Bone Miner. Res. 11, 1531–1538. Gerdhem, P., Ivaska, K. K., Alatalo, S. L., Hallen, J. M., Hellman, J., Isaksson, A., Pettersson, K., Väänänen, H. K., Akesson, K., and

111.

112.

113.

114.

115.

116.

117.

118.

119.

120.

121.

122.

123.

124. 125.

126.

127.

627

Obrandt, K. J. (2004). Biochemical markers of bone metabolism and prediction of fracture in elderly women. J. Bone Miner. Res. 19, 386–393. Ross, P. D., Kress, B. C., Parson, R. E., Wasnich, R. D., Armour, K. A., and Mizrahi, I. A. (2000). Serum bone alkaline phosphatse and calcaneus bone density predict fractures: A prospective study. Osteoporos. Int. 11, 76–82. Sornay-Rendu, E., Munoz, F., Garnero, P., Duboeuf, F., Delmas, P. D. (2003). Markers of bone turnover and history of prior fractures as predictors of long term fracture risk in postmenopausal women without osteoporosis: the OFELY Study. J. Bone Miner. Res. 18, Supp. 2: S66. van Daele, P. L. A., Seibel, M. J., Burger, H., Hofman, A., Grobbee, D. E., van Leeuwen, J. P. T. M., Birkenhager, J. C., and Pols, H. A. P. (1996). Case-control analysis of bone resorption markers, disability, and hip fracture risk: The Rotterdam study. Brit. Med. J. 312, 482–483. Qvist, P., Christgau, S., Pedersen, B. J., Schlemmer, A., and Christiansen, C. (2002). Circadian variation in the serum C-terminal telopeptide of type I collagen (Serum CTx): Effects of gender, age, menopausal status, posture, daylight, serum cortisol and fasting. Bone 31, 57–61. Bjarnason, N. H., Henriksen, E. E., Alexandersen, P., Christgau, S., Henriksen, D. B., and Christiansen, C. (2002). Mechanism of circadian variation in bone resorption. Bone 30, 307–313. Henriksen, D. B., Alexandersen, P., Bjarnason, N. H., Vilsboll, T., Hartmann, B., Henriksen, E. E., Byrjalsen, I., Krarup, T., Holst, J. J., and Christiansen, C. (2003). Role of gastrointestinal hormones in postprandial reduction of bone resorption. J. Bone Miner. Res. 18, 2180–2189. Chapurlat, R. D., Garnero, P., Breart, G., Meunier, P. J., and Delmas, P. D. (2000). Serum type I collagen breakdown product (serum CTX) predicts hip fracture risk in elderly women: the EPIDOS study. Bone 27, 283–286. Bauer, D. C., Black, D. M., Ensrud, K., Qvist, P., and Williams, E. (1999). Serum markers of bone turnover and fractures of the hip and spine: A prospective study. J. Bone Miner. Res. 14 (Suppl 1): S147. Batge, B., Deibold, J., Stein, H., Bodo, M., and Muller, P. K. (1992). Compositional analysis of the collagenous bone matrix. A study on adult normal and osteopenic bone tissue. Eur. J. Clin. Invest. 22, 805–812. Kowitz, J., Knippel, M., Schuhr, T., and Mach, J. (1992). Alteration in the extent of collagen I hydroxylation, isolated from femoral heads of women with a femoral neck fracture caused by osteoporosis. Calcif. Tissue Int. 60, 501–505. Bailey, A. J., Wotton, S. F., Sims, T. J., and Thompson, P. W. (1992). Post-translational modifications in the collagen of human osteoporotic femoral head. Biochem. Biophys. Res. Commun. 185, 801–805. Oxlund, H., Moselkilde, L., and Ortoft, G. (1996). Reduced concentration of collagen reducible cross-links in human trabecular bone with respect to age and osteoporosis. Bone 19, 479–484. Banse, X., Sims, T. J., and Bailey, A. J. (2002). Mechanical properties of adult vertebral cancellous bone: correlation with collagen intermolecular cross-links. J. Bone Miner. Res. 17, 1621–1628. Wang, X., Shen, X., Li, X., and Agrawal, C. M. (2002). Age-related changes in collagen network and toughness of bone. Bone 31, 1–7. Cloos, P. A. C., and Fledelius, C. (2000). Collagen fragments in urine derived from bone resorption are highly racemized and isomerized: a biological clock of protein aging with clinical potential. Biochem. J. 345, 473–480. Garnero, P., Fledelius, C., Gineyts, E., Serre, C. M., Vignot, E., and Delmas, P. D. (1997). Decreased β isomerisation of C-telopeptides of type I collagen in Paget’s disease of bone. J. Bone Miner. Res. 12, 1407–1415. Garnero, P., Cloos, P., Sornay-Rendu, E., Qvist, P., and Delmas, P. D. (2002). Type I collagen racemization and isomerization and the risk

628

128.

129.

130. 131.

132.

133.

134. 135.

136.

137.

PATRICK GARNERO AND PIERRE D. DELMAS of fracture in postmenopausal women: The OFELY prospective study. J. Bone Miner. Res. 17, 826–833. Garnero, P., Qvist, P., Munoz, M., Sornay-Rendu, E., Christiansen, C., and Delmas, P. D. (2004). Type I collagen C-telopeptide isomerization predicts the long-term risk of non-vertebral fracture in postmenopausal women, independently of bone mineral density and bone turnover: a non invasive index of bone collagen quality. J. Bone Miner. Res. 19 (Suppl 1), S19. Garnero, P., Borel, O., Gineyts, E., Duboeuf, F., Christiansen, C., and Delmas, P. D. (2004). The degree of posttranslational modifications of collagen is an important determinant of the toughness of cortical bone. J. Bone Miner. Res. 19 (Suppl 1), S220. Kang, A. H., and Trelstad, R. L. (1971). A collagen defect in homocystinuria. J. Clin. Invest. 52, 2571–2578. Lubec, B., Fang-Kircher, S., Lubec, T., Blom, H. J., and Boers, G. H. (1996). Evidence for McKusick’s hypotheis of deficient collagen cross-linking in patients with homocystinuria. Biochem. Bipohys. Acta. 1315, 159–162. McLean, R. R., Jacques, P. F., Selhub, J., Tucker, K. L., Samelson, E. J., Broe, K. E., Hannan, M. T., Cupples, L. A., and Kiel, D. P. (2004). Homocysteine as a predictive factor for hip fracture in older persons. N. Engl. J. Med. 350, 2042–2049. van Meurs, J. B. J., Dhonukshe-Rutten, R. A. M., and Pluijm, S. M. F. et al. (2004). Homocysteine levels and the risk of osteoporotic fracture. N. Engl. J. Med. 350, 2033–2041. Raisz, L. G. (2004). Homocysteine and osteoporotic fractures. Culprit or bystander? N. Engl. J. Med. 350, 2089–2090. Shiraki, M., Shiraki, Y., Aoki, C., and Miura, M. (2000). Vitamin K2 (menatetrenone) effectively prevents fractures and sustains lumbar spine bone mineral density in osteoporosis. J. Bone Miner. Res. 15, 515–521. Szulc, P., Chapuy, M. C., Meunier, P. J., and Delmas, P. D. (1993). Serum undercarboxylated osteocalcin is a marker of the risk of hip fracture in elderly women. J. Clin. Invest. 91, 1769–1774. Vergnaud, P., Garnero, P., Meunier, P. J., Breart, G., Kamilhagi, K., Delmas, P. D. (1997). Undercarboxylated osteocalcin measured with a specific immunoassay predicts hip fracture in elderly women: the EPIDOS study. J. Clin. Endocrinol. Metab. 82, 719–724.

138. Luukinen, H., Kakonen, S. M., Pettersson, K., Koski, K., Laippala, P., Lovgren, T., Kivela, S. L., and Vaananen, H. K. (2000). Strong prediction of fractures among older adults by the ratio of carboxylated to total serum osteocalcin. J. Bone Miner. Res. 15, 2473–2478. 139. Liu, G., and Peakcock, M. (1998). Age-related changes in serum undercarboxylated osteocalcin and its relationships with bone density, bone quality, and hip fracture. Calcif. Tissue Int. 62, 286–289. 140. Sugiyama, T., and Kawai, S. (2001). Carboxylation of osteocalcin may be related to bone quality: a possible mechanism of bone fracture prevention by vitamin K. J. Bone Miner. Metab. 19, 146–149. 141. Garnero, P., Dargent-Molina, P., Hans, D., Schott, A. M., Bréart, G., Meunier, P. J., and Delmas, P. D. (1998). Do markers of bone resorption add to bone mineral density and ultrasonographic heel measurement for the prediction of hip fracture in elderly women? The EPIDOS prospective study. Osteoporos. Int. 8, 563–569. 142. Johnell, O., Oden, A., De Laet, C., Garnero, P., Delmas, P. D., and Kanis, J. A. (2002). Biochemical markers and the assessment of fracture probability. Osteoporos. Int. 13, 523–526. 143. McClung, M.R., Geusens, P., Miller, P. D., Zippel, H., Bensen, W. G., Roux, C., Adami, S., Fogelman, I., Diamond, T., Eastell, R., Meunier, P. J., and Reginster, J. Y. (2001). Hip Intervention Program Study Group. Effect of risedronate on the risk of hip fracture in elderly women. N. Engl. J. Med. 344, 333–340. 144. Kanis, J. A., Johnell, O., Black, D. M., Downs, R. W. Jr, Sarkar, S., Fuerst, T., Secrest, R. J., and Pavo, I. (2003). Effect of raloxifène on the risk of new vertebral fracture in post-menopausal women with osteopenia or osteoporosis: a reanalysis of the multiple outcomes of raloxifene evaluation trial. Bone 33, 293–300. 145. Black, D. M., Thompson, D., Quandt, S., Palermo, L., Ensrud, K., and Johnell, O. (2002). Alendronate reduces the risk of vertebral fracture in women with BMD T-scores above −2.5: results from the Fracture Intervention Trial (FIT). Osteoporos. Int. 13(Suppl 1): S27. 146. Bauer, D. C., Garnero, P., Hochberg, M. C., Santora, A., Delmas, P. D., Ewing, S. K., and Black, D. M. (2003). Pre-treatment bone turnover and fracture efficacy of alendronate: the Fracture Intervention Trial. J. Bone Miner. Res. 18(Suppl 2): S55.

Chapter 38

Monitoring Anabolic Treatment John P. Bilezikian and Mishaela R. Rubin

IV. Actions of Parathyroid Hormone to Improve Bone Quality V. Conclusions References

I. Introduction II. Cellular and Regulatory Mechanisms of the Anabolic Actions of Parathyroid Hormone III. Pharmacokinetics of Teriparatide in Human Subjects

I. INTRODUCTION

to reconstruct skeletal microarchitecture, an endpoint not shared by any of the antiresorptives [12]. The most promising anabolic agent to date, and available generally throughout the world at this time, is parathyroid hormone (PTH). The first form of PTH to be studied and approved in human subjects with osteoporosis is the aminoterminal fragment of human parathyroid hormone [hPTH (1-34)]. This peptide is known generically as teraparatide. Teriparatide appears to contain all the classical anabolic properties of PTH(1-84), the full-length molecule. Teriparatide can refer either to synthetic human PTH(1-34) or to the recombinant human form of PTH(1-34). The full-length molecule PTH(1-84) has also been the subject of major clinical trials. In this chapter, when reference is made to full-length PTH, the term PTH(1-84) will be used. When reference is made to any form of parathyroid hormone, the term, “parathyroid hormone” or PTH will be utilized. We will focus on means by which parathyroid hormone treatment for osteoporosis can be monitored. The chapter will first consider mechanisms by which PTH confers its anabolic properties on the human skeleton. Then, key properties that are affected by parathyroid hormone, with due reference to the major clinical investigations that have been published, will be considered. Some of the mechanisms

Anabolic agents represent an important new advance in the therapy of osteoporosis. Until now, the mainstay of therapy for osteoporosis has been antiresorptive in mechanism. Drugs such as estrogen, raloxifene, alendronate, risedronate, ibandronate, and calcitonin all inhibit osteoclast-mediated bone loss and, thus, reduce bone turnover [1–8]. The reduction in bone turnover leads to a reduction in the frequency at which new bone remodeling units are activated [9, 10]. This mechanism, combined with a greater inhibition of bone resorption than bone formation, leads to better bone balance and a smaller remodeling space [11]. The antiresorptives, thus, increase bone density, mineralization density, and now shown for the bisphosphonates, maintain skeletal microarchitecture. The concept of an anabolic agent is based upon a therapeutic mechanism entirely distinct from inhibition of bone resorption. The mechanism of anabolic agents is targeted upon processes associated with bone formation. By stimulating bone formation to a greater extent and earlier than bone resorption, the anabolics have the potential not only to increase bone density but also to improve the microarchitecture of bone. Such actions even have the potential Dynamics of Bone and Cartilage Metabolism

Department of Medicine, College of Physicians and Surgeons, Columbia University, New York, NY, USA

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by which parathyroid hormone improves bone quality are not readily amenable to monitoring yet but they are discussed because in the near future some of them will become available. They also underscore the potential for tracking the clinical actions of parathyroid hormone by a variety of quantitative measures.

II. CELLULAR AND REGULATORY MECHANISMS OF THE ANABOLIC ACTIONS OF PARATHYROID HORMONE The catabolic actions of PTH on the skeleton, particularly at sites of cortical bone (distal third of the radius) are seen under conditions of continuous exposure, such as in primary hyperparathyroidism (PHPT) [13, 14]. In PHPT, however, the cancellous skeleton, best represented by the lumbar spine, typically does not show catabolic consequences, especially when cohorts of subjects with asymptomatic PHPT are described. It is particularly noteworthy that in postmenopausal women with PHPT, who are not taking estrogen therapy, the lumbar spine is actually quite well preserved [15]. Thus, even in the context of continuous exposure of the skeleton to PTH, clues to the anabolic potential of PTH can be gleaned (Fig. 1). More specific anabolic actions of PTH can be appreciated through studies in which PTH is administered intermittently, once daily, and in low dosage. When PTH is administered in this way the anabolic features of PTH action predominate [16–27]. The site of PTH’s anabolic actions is seen best in the cancellous-rich lumbar spine when conventional bone mineral density (BMD) is used as a marker of efficacy. With PTH, bone formation rate increases, due possibly to direct stimulation of osteoblast function [28, 29] and to

Figure 1

inhibition of apoptosis that extends the life of the osteoblast [30]. In contrast to antiresorptive agents, the individual remodeling unit is stimulated, not suppressed. As a result, more bone is formed within each remodeling unit [21, 31]. PTH may initially stimulate bone formation directly without a requirement for prior resorption [32]. This concept argues for an initial action of PTH that does not require ongoing active bone remodeling, a biological event that is initiated by osteoclast activation. Evidence for this comes from the finding of active formation surfaces in the absence of an underlying scalloped cement line [33, 34], particularly at the periosteal surfaces where bone formation normally is extremely low [35]. This may occur by activation of lining cells on previously quiescent bone surfaces [36, 37], as well as by migration of osteoblasts engaged in remodeling-based formation outside the borders of the bone remodeling unit [38–40]. Not all studies, however, have been able to demonstrate direct activation of lining cells [41]. Other studies have linked the magnitude of the densitometric effect of PTH to baseline levels of bone turnover, suggesting that PTH may require an element of bone remodeling in order for it to preferentially stimulate osteoblasts. Whether there is a direct initial action of PTH to stimulate osteoblasts– or their precursors– or rather the need for osteoclast-mediated bone turnover, the earliest biochemical evidence of PTH action is an increase in markers of bone formation followed later by an increase in markers of bone resorption. These cellular actions provide a basis for using markers of bone turnover to monitor the actions of PTH. Despite the attention given to the osteoblast and the osteoclast in the cellular actions of PTH, it is also important to consider the osteocyte. Osteocytes are believed to function as mechanosensors, regulating the bone remodeling

Pattern of change in bone density in primary hyperparathyroidism. In postmenopausal women with mild disease a typical pattern is shown in which bone density of the lumbar spine is relatively well preserved in contrast to the distal radius (1/3) in which bone density is preferentially reduced. (Adapted from reference [15].)

Chapter 38 Monitoring Anabolic Treatment

process [42]. PTH appears to modulate the response of osteocytes to mechanical shear and strain forces [43, 44], due in part to preservation of osteocyte viability [30]. Under conditions of mechanical loading (i.e. physical activity), PTH is most effective [45, 46]. In contrast, unloading weakens the anabolic actions of PTH on trabecular bone by a PTH-independent effect to reduce bone formation and to increase bone resorption [45]. Possible cellular mechanisms for the anabolic effects of PTH include stimulation of growth factors [47], especially IGF-I [48, 49]. PTH does not increase bone formation in animals that have been genetically manipulated and lack IGF-I synthetic and secretory capacities [50, 51]. Variability in IGF-I could theoretically contribute to variability in the skeletal response to PTH treatment. For example, in a study of PTH(1-84) treatment in postmenopausal women, nearly 20% of the women had increases in spine BMD by QCT of greater than 60% after 1 year. The distribution curve describing the response has a right-sided “tail” on the exuberance side with very few subjects showing a reduction in bone density [52]. Although one candidate for this variability is IGF-I, in the study of Sellmeyer it was not possible to define factors to predict the exuberant response to PTH. Healthy inbred strains of mice with differences in skeletal synthesis of IGF-I also tend to respond differently to PTH [53]. In 39 iliac crest biopsy samples from the largest clinical trial of PTH(1-34), immunoreactivity for IGF-I was sparse, while the expression of IGF-II in cancellous and cortical bone was greater with teriparatide treatment than placebo [54]. Similarly, studies in mice with IGF-I receptor null mutation suggest that the IGF-I receptor appears to be necessary for PTH to stimulate osteoprogenitor cell proliferation and differentiation [55]. Another possible mechanism of PTH’s anabolic actions involves PTH-mediated increases in RANK ligand (RANKL) [56], a signaling molecule that can bind to and interact with RANK on the surface of neighboring osteoclasts and osteoclast precursors. Absence of the RANKL– RANK pathway abolishes the ability of PTH to increase osteoclastic bone resorption. Along with the increase in RANKL, PTH decreases OPG, a natural inhibitor of RANKL, by the osteoblast. Continuous exposure to PTH therefore increases the probability that the osteoblast or osteoblast precursor will stimulate osteoclastic bone resorption. If an osteoblast is exposed to high concentrations of PTH for only a few hours, it appears that the RANKL– OPG–RANK interaction is altered [57]. Lane and colleagues showed that in 51 postmenopausal women on HRT and chronic glucocorticoid therapy, RANKL, as well as the osteoblast cytokines IL-6 and IL-6sR, increased within 1 month of initiation of PTH therapy, suggesting that PTH rapidly stimulates osteoblast maturation and

631 function [57]. Although these results seem to argue against a direct effect of PTH on osteoblasts, it is likely that in time, PTH does require active osteoclastogenesis to continue to exert its anabolic effects on osteoblasts. For example, c-fosdeficient mice do not show an anabolic skeletal response to PTH [58]. c-fos is an essential transcription factor for osteoblasts and osteoclasts and absence of c-fos results in osteopetrosis (lack of bone resorption), confirming that osteoclast activation is a prerequisite for normal bone remodeling. Other possible mechanisms for the anabolic action of PTH include stimulation of unique subsets of “bone forming” genes, including gene expression of osteocalcin and tartrate-resistant acid phosphatase (TRAP) [27]. In mice, basic fibroblast growth factor (FGF-2) has also been found to be a critical mediator of the anabolic effect of PTH on bone [59] (see also Chapters 15, 16, and 20).

III. PHARMACOKINETICS OF TERIPARATIDE IN HUMAN SUBJECTS Data from the largest study in postmenopausal women show that between 4 and 6 hours after injection, the serum calcium peaks, but the level remains within the normal physiological range, with the average increment being about 0.8 mg/dl. The increase in serum calcium eventually returns to baseline prior to the next dose [60]. In the major phase III trial with teriparatide, there was a very small, insignificant incidence of hypercalcemia when the approved 20 µg dose was used. Peak circulating concentration of PTH, equivalent to about 2–2.5 times the usual circulating level of endogenous PTH, is reached by 45–60 minutes after administration, followed by complete disappearance in 4–6 h [61]. During the time when teriparatide is detectable in the circulation, PTH(1-84), the native secretory form, is not detected [62]. As the level of teriparatide falls, the level of endogenous PTH rises. The total amount of PTH to which a subject is exposed when receiving teriparatide is actually the same amount of PTH that would be secreted under normal circumstances over a 24-hour period. The big difference, thus, is not the cumulative exposure to PTH, but rather to the skeletal dynamics triggered by the intermittent, pulsatile administration of PTH. PTH(1-84) has a longer half-life, peaking at 120 min and disappearing from the circulation after 8 h [63]. Recombinant PTHrelated protein, PTHrP, which is also being investigated as a potential therapy for osteoporosis, has a very rapid half-life, peaking at 15 min and disappearing from the circulation after only 1–2 h [64]. It is not known whether these differences in pharmacokinetics will be important in terms of any differences that might be observed with the clinical use of these agents.

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IV. ACTIONS OF PARATHYROID HORMONE TO IMPROVE BONE QUALITY A. PTH and Bone Density Bone density is an important bone property that is affected by effective therapies for osteoporosis. As was the case for antiresorptive agents, it was entirely reasonable to assume that changes in bone mineral density, one of the most important predictors of fracture risk, would correlate with reduction in fracture risk. The seminal clinical trial of Neer et al. tested daily administration of 20 or 40 µg of subcutaneously injected teriparatide in 1637 women with severe postmenopausal osteoporosis (i.e. low bone mineral density and fractures) using a randomized, placebo-controlled, double-blinded protocol [60]. Median follow-up was 21 months. For the two doses of PTH, the increase in spine bone mineral density averaged 10–14%. Post-hoc analysis demonstrated that in 96% of individuals there was an increase at the spine, at least above baseline, and in 72% the increase was 5% [65]. The increase in femoral neck bone mineral density was also significant but the change was smaller (approximately 3%) and took longer to occur. At the 20 µg dose there was no change in cortical bone density (distal third of the radius site) but total body bone mineral density increased significantly.

In another randomized, double-blind placebo-controlled trial, 40 µg of hPTH(1-34) or 10 mg alendronate was administered daily for 14 months to 146 postmenopausal osteoporotic women [66]. The increases in BMD were twice as great with PTH as with alendronate. The first randomized, controlled trial of PTH in men with idiopathic osteoporosis was carried out by Kurland et al. [67]. Twenty three men, 30–68 years old, with osteoporosis as defined by Z-scores less than – 2.0 SD at the lumbar spine or femoral neck, and no definable cause for bone loss, were randomized to 400 U/d of teriparatide (equivalent to about 25 µg/d) or placebo in a randomized, double-blind experimental protocol for 18 months. The men who received teriparatide demonstrated an impressive 13.5% increase in lumbar spine bone density (Fig. 2). Hip bone density increased but typically was slower to change and not as robust, although significant. Cortical bone density at the distal radius did not increase in comparison to placebo. In the larger teriparatide study in men that was designed to be the counterpart to the study by Neer et al. of postmenopausal women, Orwoll et al. reported on 437 men with idiopathic or hypogonadal osteoporosis [68]. They were randomly assigned in a blinded fashion either to placebo, or to teriparatide, 20 µg or 40 µg daily, for a mean of 11 months [68]. BMD increased significantly at the lumbar spine in the treatment groups by 6% and 9%, respectively, regardless of gonadal status. Over the 11 months

Figure 2 Teraparatide increases bone mineral density at cancellous sites in men with osteoporosis. The cancellous bone of the lumbar spine responds to teraparatide therapy with a marked increase in bone density. In contrast the distal radius (1/3) site does not change. The hip regions have an intermediate response. (Adapted from reference 67.)

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of the study, the increases in bone density at the lumbar spine and the hip tracked closely along the time-dependent increase in bone density at those sites in the study of postmenopausal women by Neer et al. In a phase II trial of PTH(1-84), an agent that has been the subject of a limited number of studies but is in active development as another anabolic agent for osteoporosis, subjects were administered either placebo, or three of five doses: 50, 75, or 100 µg for 12 months [69]. There were time- and dose-related increases in lumbar spine BMD with the 100 µg dose associated with the largest increase in BMD. The pivotal clinical trial with teriparatide showed major reductions in vertebral and non-vertebral fracture incidence [60]. As compared with placebo, teraparatide reduced the risk of one or more new vertebral fractures by 65% and 69%, at 20 µg and 40 µg, respectively. This decline in fracture incidence compares favorably with the relative risk reduction for fractures in women treated with bisphosphonates [70]. The overall incidence of new nonvertebral fractures was reduced by 35% and 40%, respectively. When fragility fractures were considered apart from total fracture incidence, the reduction in new nonvertebral fractures was 53% and 54%, respectively. Hip fracture incidence was not analyzed separately because the study was not sufficiently powered to examine this point. In the male study, fracture incidence could not be assessed because of the short (11-month) duration of the trial. However, fractures were assessed in a follow-up observational study in which 279 men from the original cohort could be assessed 18 months at the end of the treatment period by lateral thoracic and lumbar radiographs. A total of 25–30% of the men reported the use of an antiresorptive therapy during the follow-up period, with the men having been treated with placebo utilizing antiresorptive therapy to a greater extent than those who were treated with either dose of teriparatide (36% vs 25%). Nevertheless, the incidence of moderate to severe vertebral fractures was significantly reduced below the placebo group when the 20 µg and 40 µg doses were combined (6.8% vs. 1.1%; P < 0.02) [71]. PTH(1-84) has also been shown to reduce vertebral fracture incidence in a group of postmenopausal women with osteoporosis [72]. The facile assumption that the reduction in fracture risk is directly associated with the increase in bone density is based upon the powerful relationship between reductions in bone density and increases in fracture risk. A number of studies have shown, however, that improvements in bone mineral density account for only a minor component of overall reduction in fracture risk [73, 74]. This would appear to be true not only for antiresorptives but also for teriparatide and PTH(1-84). Increases in bone density

appear to make a limited contribution to overall fracture efficacy with PTH. This observation underscores the fact that the efficacy of PTH as a therapy for osteoporosis resides in its effects on properties of bone other than bone density as measured by DXA. The results also lead to a cautionary note in direct comparison studies among therapies for osteoporosis. For example, one cannot conclude that teriparatide is more effective than alendronate because changes in bone density are much greater with the anabolic agent [66]. A similar cautionary note is advised in interpreting the results of the recent study in which densitometric differences were compared between alendronate and risedronate [75]. The greater increase in bone density with alendronate does not allow one to conclude that it is more efficacious. Although increases in bone density do not predict fracture efficacy, nevertheless, it is commonly used to monitor the effects of teriparatide therapy [76]. Since increases in bone density do not correlate well with reductions in fracture incidence, what are the reasons for following teraparatide therapy with measurements of bone density? Among the most important reasons, bone densitometry is readily available and does give some insight into issues such as: compliance, possible intercurrent events and fracture efficacy. If subjects do not take PTH properly, this may well become evident when bone density in follow-up does not change. If a medical event has intervened that places patients at risk for further bone loss, this may become evident only after noting the decline in bone density upon follow-up testing. Another important point is that compliance with the medication is likely to be enhanced when patients “see” a positive result, namely that bone density is improving. Finally, it is at this time the best way we have to monitor the effects of therapy, albeit an imperfect one. When bone density is used to track the course of teriparatide or PTH(1-84) therapy, it is important to note that major changes in BMD will occur primarily in the lumbar spine. In the hip, the changes will be much more modest. At the distal radius, it is in fact unlikely that BMD will improve over baseline values.

B. Teriparatide and Bone Turnover Similar to the antiresorptive agents [77, 78], it is likely that the therapeutic actions of PTH are primarily based upon its actions to influence bone turnover. A very important feature of PTH therapy that distinguishes it from antiresorptive therapy is that it stimulates bone turnover. Initially, bone formation markers rise rapidly, as early as 1 week after teriparatide therapy has been initiated. At a variable time thereafter, bone resorption markers increase. The difference in time course between the initial stimulation

634 of bone formation and the subsequent stimulation of bone resorption has given rise to the concept of the “anabolic window,” a period of time when the anabolic actions of PTH are maximal (Fig. 3). Consistent with this concept, an increase in the bone formation rate, as confirmed by tetracycline labeling of bone biopsies, has been reported on the periosteal surface of iliac cortical bone in patients treated for only 1 month with teriparatide [32]. Other data suggest that the resorptive phase of the remodeling cycle is essentially bypassed for the first 6 months of therapy [79]. This allows for a substantial gain in bone mass before the number of resorption cavities begins to increase. The subsequent actions of PTH to stimulate bone remodeling can be viewed as the second phase of PTH actions. During this second phase of PTH action, the osteoclast is clearly involved. In contrast to the antiresorptive agents, PTH increases the remodeling space with greater remodeling units being activated. Each remodeling unit experiences a greater restitutive osteoblast response. The bone remodeling unit is thus more favorably balanced. The third phase of PTH action may relate directly to the activated osteoblasts in the remodeling unit that literally spill onto the surface of bone further expanding the scope of the osteoblastic response to PTH. The latter description has been shown in classical studies by Reeve et al. [80] and more recently by Lindsay et al. [34]. Can baseline bone turnover markers be used to predict the response to PTH? Kurland has shown that the baseline level of pyridinoline and the 3 month change in osteocalcin predict changes in bone density accounting for 70% of the variance in bone density after 18 months [67]. Moreover, pyridinoline, that was not associated with bone formation markers at baseline, became highly correlated with bone formation markers after PTH treatment. At first glance, this is an unexpected observation because

Figure 3

The Anabolic Window. Based upon the difference in kinetics of changes between bone formation and bone resorption markers, an “anabolic window” is formed during which the actions of parathyroid hormone are believed to be maximally anabolic.

John P. Bilezikian and Mishaela R. Rubin

it implies that those with greater baseline bone turnover markers will have a greater response to PTH therapy. This finding can be explained, however, if considered within the context of the bone remodeling unit. If PTH depends upon the presence of functional osteoblasts or their precursors (see above), then greater access to these cells by PTH will be associated with a greater response to PTH. One might expect, therefore, that low bone turnover states might be associated with an initial sluggish or delayed response to PTH. This might be due to pathophysiological processes leading to low bone turnover or to pharmacological suppression of bone turnover due to agents such as the bisphosphonates (see below). While these relationships between bone turnover and changes in bone density are of interest, one would like to have data that relate these initial bone turnover data to fracture risk. For this question, one must turn to the larger clinical trials with teriparatide. Delmas et al. have studied patients from the pivotal clinical trial of teriparatide [81]. Although fracture risk was higher in those with the highest bone turnover, there was no relationship between the percentile of five different bone turnover markers (BSAP, P1CP, P1NP, NTX, DPD) and relative risk reduction of osteoporotic fractures. Since bone markers change quickly and in a robust manner with both teriparatide and PTH(1-84), a reasonable question follows: can changes in bone turnover markers be used to monitor the effect of PTH? One of the problems in using bone markers to monitor the response to antiresorptive agents is that the reductions, although significant, may not exceed the least significant change of the measurement. Given the fact that bone turnover markers have an intrinsic high biological variability and that the measurements themselves are subject to assay variability, bone turnover markers have not gained acceptance as a way to monitor therapy with antiresorptive drugs. The situation is theoretically different with PTH because the increases in bone turnover markers can be two- to four-fold above baseline, a difference that should easily exceed the least significant change of the measurement. Furthermore, Dobnig et al. have shown that early changes in bone formation markers, but not bone resorption markers, after teriparatide administration are associated with increases in structural parameters of bone as assessed by histomorphometry and by microCT analysis of bone biopsy specimens [82]. By quantitative histomorphometry, changes in P1CP and BSAP were positively correlated with the following structural parameters: mean wall thickness and trabecular bone volume; and by microCT: trabecular thickness and trabecular bone volume/total bone volume. The increase in P1CP at 3 months was inversely correlated with marrow star volume, an index of trabecular connectivity. The correlation with bone formation markers, not bone resorption markers,

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is consistent with concepts that the primary and most important actions of teriparatide therapy are those that relate to bone formation. The secondary effects to stimulate bone remodeling may not be a key aspect of the anabolic actions of PTH. A strategy has therefore been proposed by which responders to PTH can be identified early by using bone turnover markers. PINP, a marker of bone formation that changes early in the course of therapy, is measured after 3 months of PTH. The patient is considered a responder if the change is more than 15 µg/L [83]. Changes in a sensitive bone formation marker, like P1NP, may thus be used to gain an early indication of PTH effectiveness. If an individual does not show this short-term increase in P1NP, reasons for the delayed response should be considered. In subjects who are in a low turnover state, as identified perhaps by baseline low turnover markers, there might be a delay in the response to teriparatide. Under these circumstances, substantial changes in bone formation markers may take a longer period of time to reach their peak values. In subjects whose bone turnover markers are suppressed in the context of previous treatment with an antiresorptive, there might also be a delay (see below). In any event, if P1NP has not changed substantially after 3 months, it would seem prudent to evaluate reasons for it and to repeat the bone marker measurement in another 3 months by which time one would expect clear changes to be apparent. Virtually all studies that have been conducted with teriparatide have shown that both formation and resorption indices eventually decline towards or to baseline levels [79, 84]. The actual time course of this decline is quite variable from study to study and within groups of study subjects. The eventual fall in bone turnover marker levels with continued use of teriparatide may signal a waning of the anabolic effect on bone density, although in some clinical trials BMD continues to increase despite this downturn [67]. The waning of the effects of teriparatide on bone turnover markers could be used as an index of waning efficacy. On the other hand, there is no good evidence that the reduction in bone turnover markers is associated with any reduction in the effectiveness of PTH. Yet, there may still be a utility to measuring bone turnover markers as a guide to the length of therapy.

feature in osteoporosis, accounts importantly for increased fracture risk [21] (see also Chapter 13). Increased osteoclast activity that is associated with high bone turnover states in postmenopausal women leads to perforation of cancellous plates, loss of trabeculae and trabecular discontinuity [87]. Since much of the strength of cancellous bone is derived from its microarchitectural organization, loss of trabecular connectivity predisposes to vertebral fracture. Preserving the trabecular network is even more important for bone strength than maintaining trabecular thickness [88]. When the trabecular network becomes discontinuous due to perforations, more likely to occur in the context of excessive remodeling, bone strength is affected, without any necessary change in bone density. Bisphosphonates have been shown to maintain microarchitecture, which probably contributes to improvements in bone strength [89, 90]. Recent work in human subjects with risedronate shows that plate-like structures of trabeculae are maintained [91]. Recent data with alendronate also confirm that microarchitecture is maintained [92]. There is no evidence at this time, however, that the bisphosphonates actually improve microarchitectural qualities of bone. The situation is very different with PTH in which microarchitectural features of bone are clearly improved. Histomorphometric analysis of paired biopsies before and after treatment with PTH, using standard histomorphometry and three-dimensional microcomputed tomographic analyses, have shown not only quantitative improvements in cancellous bone indices but also major improvements in indices of connectivity (Fig. 4) [93]. There is increasing connectivity of trabeculae, thickening of trabeculae and conversion from a more rod-like to a more normal plate-like appearance [21, 93]. Trabecular elements become connected or reconnected. Marrow star volume is reduced documenting PTH-enhanced connectivity [21, 93, 94]. Another potential mechanism for enhanced connectivity is an effect of PTH to increase tunneling of thickened trabeculae, thus resulting in increased trabecular number and connectivity [21]. It is unclear whether therapy stimulates the budding of new trabeculae de novo, or alternatively, leads to the thickening of trabeculae that are bifurcated by tunneling resorption [26].

D. PTH and Bone Geometry C. Parathyroid Hormone and Microarchitecture Assessment of skeletal microarchitecture is not feasible yet in the clinical arena. Nevertheless, it is useful to review the effects of PTH to improve skeletal microarchitecture, in part, because non-invasive imaging instruments are being developed at this time to access this kind of information [85, 86]. Microarchitectural deterioration, a common

Again, measurement of bone geometry as an index of PTH action is not yet feasible in the clinical arena. But, like considerations of microarchitecture, such measurements may become available clinically in the near future. Therefore, it is useful to review the effects of PTH in this regard. With PTH-driven increases in bone turnover, expectations of negative effects on cortical bone deserve

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Figure 4 After therapy with teraparatide, microarchitecture of bone is enhanced. Depiction by microCT analysis of bone biopsy specimens from a patient before and after therapy is shown (reproduced from reference 21 with permission).

specific consideration. It has been postulated that removal of cortical bone by PTH provides calcium to supply new trabecular bone during the initial period of therapy [95], possibly through increases in cortical porosity. It is well established now that PTH does not increase bone density at the distal third site of the radius. In some situations, there is a small decline in bone density at this site. This densitometric observation, combined with PTH-mediated increases in intracortical porosity, has raised concern that perhaps the cortical skeleton is at risk. Nevertheless, there is no detrimental effect of PTH on the mechanical properties of cortical bone [25]. One reason to account for this observation is actions of PTH to favorably alter bone geometry. The inner endocortical surface of bone is a site of PTH-induced resorption and increased cortical porosity. When compensated by changes in the outer diameter of bone by the actions of PTH to stimulate periosteal bone apposition, the net effect, that is a change in the ratio of the outer to inner diameter of bone, leads to increased, not decreased, bone strength. Moreover, in time, periosteal apposition leads actually to increases in cortical area and cortical thickness [25]. Thus, expansion of the inner diameter of tubular bone due to PTH-induced endocortical remodeling [16] has little effect on calculated bending strength [23], because the effects are offset by an increase in both cross-sectional area and in cortical thickness, both consequences of periosteal bone apposition [21, 96]. Both animal and human studies confirm expectations that cortical bone is actually stronger under the influence of intermittent PTH administration. Ovariectomized monkeys treated with teriparatide develop greater cortical porosity at the femur but show increased bone strength, because

enhanced cross-sectional area and increased cortical thickness, as well as enhanced trabecular bone volume, more than compensate for the porosity [97]. In human subjects, histomorphometric analysis of bone biopsy specimens before and after 18–26 months of teriparatide therapy similarly show a net anabolic effect on cortical bone [21, 93]. Instead of an increase in cortical porosity, surprising gains are seen on the endocortical surface. A decrease in the eroded perimeter is consistent with a reduction in resorption at the endocortical surface [21]. The positive effect of increased endocortical wall width and cortical thickness may be enhanced by a reduction in resorption on that surface due to a marked decrease in eroded perimeter. Anabolic action may additionally occur at the subperiosteal surface, but it is not possible to evaluate wall width of newly formed bone units there. The concept that PTH increases cortical thickness was confirmed when 101 postmenopausal women, treated with PTH, underwent peripheral quantitative computed tomography (pQCT) of the proximal radius after a median 18 months of treatment to assess specific changes in cortical bone density [98]. Similar to the non-human primate model, PTH treatment results in significantly greater periosteal and endocortical circumferences and cortical area. These differences result in greater polar and axial moments of inertia and torsional bone strength index, biomechanical indicators of skeletal resistance to bending and torsional loading. Cross-sectional diameter thus increases, so that the ratio of outer to inner diameter increases with consequent increases in strength [99, 100]. These geometrical changes contrast with those seen with primary hyperparathyroidism, in which both an expansion

Chapter 38 Monitoring Anabolic Treatment

of inner and outer diameter occurs. In the case of intermittent PTH therapy, the outer diameter increases and the inner diameter decreases. Greater periosteal distribution of cortical bone, reflected in the moments of inertia, may contribute to the reduction in non-vertebral fractures that are not explained by changes in BMD. An increase in cortical thickness was similarly found in 40 PTH-treated women who underwent digital X-ray radiogrammetry, a technology that also measures changes in radial cortical thickness and bone size [101, 102]. Larger vertebral size likely contributes to fracture risk reduction [103]. When 51 postmenopausal women on chronic glucocorticoid therapy and estrogen were randomized to teriparatide for 1 year vs. placebo, vertebral cross-sectional area increased after 1 year by 4.8%. This improvement in cross-sectional area was maintained 1 year after PTH was discontinued [104]. Similar findings have recently been reported for changes in cross-sectional area of the femoral neck [105]. These observations help to substantiate the densitometric observations at a structural level, suggesting that PTH may be improving the skeleton in ways that are distinctly different from the anitresorptives, and help to allay concerns that PTH may have adverse effects upon the cortical skeleton. Furthermore, fracture data from the study of Neer et al. [60] clearly indicate a substantial reduction in fractures of the non-vertebral skeleton. It is this observation by Neer et al. that needs to be emphasized, even though the precise mechanisms by which PTH reduces non-vertebral fractures are not clear. It is apparent that changes in bone geometry account, as least in part, for these beneficial effects at non-vertebral sites.

E. PTH and Mineralization Mineralization is mainly a passive process, which follows the deposition of the organic matrix (see also Chapter 12). It is enhanced when the rate of bone remodeling is reduced. There are two phases to mineralization. The first, a rapid phase, follows soon after osteoid is formed. The second, a slower phase, progresses until mineralization is complete. By inhibiting bone turnover, antiresorptive drugs provide a greater time period for secondary mineralization [106–108]. Increases in bone mineral content have a profound effect on bone strength [109]. Using newer methods such as quantitative back-scattered electron imaging (qBEI), it has been shown that alendronate and risedronate increase bone mineralization [110, 111]. With conventional technology such as DXA, this property of the bisphosphonates to enhance mineralization density is measured as part of the increase in bone mineral density. It cannot be measured separately with DXA.

637 With teriparatide, the initial effects are to reduce mineralization density because the bone remodeling space, comprised of undermineralized bone, is increased. Mineralization density is reduced also because PTH erodes fully mineralized endocortical bone. The compensatory increase in periosteal bone is likely to be undermineralized, at least initially. A dynamic is thus created that tends to change bone density as measured by DXA in the opposite direction. The increase in bone size tends to increase areal bone density while the increase in undermineralized periosteal bone tends to decrease it. The reduction in mineralization density is likely to be a transient one because, over time, one would expect both the remodeling space and the undermineralized periosteal bone to be more fully mineralized. How changes in bone density, bone turnover, microarchitecture, bone geometry, and mineralization all contribute to an improvement in bone strength when teriparatide is used as a therapy for osteoporosis can be viewed in the following way. A dynamic is established among these determinants of bone strength. The increase in bone size and, where seen, in bone density tends to increase bone strength. The reduction in bone mineralization and, where seen, in bone density tends to decrease bone strength. The increase in bone density of cancellous bone is dominated by bone accrual while the reduction in bone density at the cortical skeleton is dominated by a reduction in mineralization density. At both sites, however, bone strength is improved with important and determinant microarchitectural improvements being seen at both cancellous and cortical sites.

F. Sequential and Combination Therapy with Teriparatide and an Antiresorptive Agent In this section, we consider three important questions regarding PTH therapy for osteoporosis. They are relevant because the antecedent therapeutic history, concomitant use of anabolic and antiresorptive therapy, and subsequent effects on measurable parameters such as bone density and bone turnover are all important to issues of monitoring. Question 1: Does the Previous Use of an Antiresorptive Influence the Response to Teriparatide?

Many subjects who are going to be treated with teriparatide have previously been treated with a bisphosphonate or other antiresorptive. It is important to determine whether these individuals are as likely to respond to teriparatide – and what the nature of the response will be – as those who have never previously been treated with an antiresorptive. Cosman et al. studied postmenopausal women who had

638 been previously treated for at least 1 year with estrogen [79]. These patients were then treated with teriparatide for 3 years. The group receiving teraparatide had significant increases in bone density of the spine (13%), the hip (4.4%) and the total body (3.7%). It is noteworthy that bone density in the vertebral spine began to increase with no delay and continued to increase in a linear fashion for the entire 3-year study. At the distal radius there was no significant change in bone density. With the discordance between early changes in bone formation markers (osteocalcin) and later changes in bone resorption markers, as literally shown in all studies with teriparatide, there was a return of both sets of indices to baseline values within 2.5 years of initiation of treatment. It is of interest that the return of bone markers to baseline values was not associated with a reduction in the rate of the 3-year gain in BMD at the vertebral spine. Roe et al. have obtained data from a study in postmenopausal women that was similar in design to the study of Cosman et al. Women who were all on estrogen replacement therapy for at least 1 year received either teriparatide or no further therapy except estrogen. This was not a blinded study because women in the placebo group did not receive an injection. Bone density at the lumbar spine after 2 years increased by approximately 28%. Using QCT, a technology that measures rather specifically cancellous bone of the lumbar spine, increases in vertebral bone mineral density approached 80% after 2 years [112]. It is clear the pretreatment with estrogen does not impede the subsequent response to teriparatide. Ettinger et al. studied previous exposure either to raloxifene or alendronate followed by teriparatide [113]. Both groups of subjects had been treated for an average of 28 months with the antiresorptive prior to being given teraparatide. In most respects the subjects, who were not part of a randomized, matched cohort, were similar. However, women who had previously been treated with alendronate had much lower bone turnover markers than those previously treated with raloxifene. After raloxifene, the increase in BMD of the lumbar spine was linear and progressive whereas the increase in BMD of the lumbar spine after alendronate was delayed for approximately 6 months. At the hip, the alendronate-treated subjects actually had an initial decline in bone density for the first 6 months. From these results, one might be tempted to conclude that antiresorptive agents that do not have a profound effect on bone turnover permit teraparatide to act promptly and in a robust manner whereas more potent antiresorptive agents have an initial suppressive effect on teraparatide action. However, the results of Cosman et al. raise an additional point because, in their study, women with postmenopausal osteoporosis who had been previously

John P. Bilezikian and Mishaela R. Rubin

treated with alendronate for the same duration as the study by Ettinger et al. responded with the robust changes in BMD and biochemical markers [114]. It is not clear why these results are discrepant but it is distinctly possible that it is not so much the specific antiresorptive that is used prior to teriparatide that dictates the response but rather the extent to which bone turnover is reduced. Bone turnover markers appear to have been more markedly reduced in the study of Ettinger et al. than in the study by Cosman et al. Question 2: Does Combination Therapy with Teriparatide and an Antiresorptive Lead to Greater Effects than Either Drug Alone?

There is an attractive theoretical basis for thinking that combination therapy with an antiresorptive and PTH would be more beneficial than monotherapy with either an antiresorptive or PTH. If bone resorption is inhibited and bone formation is stimulated, as could be the case with simultaneous combination therapy, the results might be much better than with either agent alone. Important data contrary to this reasoning have been provided by Black et al. [115] and Finkelstein et al. [116]. These two groups independently completed trials with PTH alone, alendronate alone, or the combination of PTH and alendronate. The study of Black et al. involved postmenopausal women treated with 100 µg of PTH(1-84). The study of Finkelstein et al. involved men treated with 40 µg of teriparatide. Both studies utilized DXA and QCT to measure areal or volumetric BMD respectively. With either measurement, monotherapy with PTH exceeded densitometric gains with combination therapy or alendronate alone at the lumbar spine. Measurement of trabecular bone by QCT, in fact, showed that combination therapy was actually associated with substantially smaller increases in BMD than monotherapy with PTH(1-84). Bone turnover markers followed the expected course for anabolic (increases) or antiresorptive (decreases) therapy alone. For combination therapy, however, bone markers followed the course of alendronate, not PTH, with reductions in bone formation and bone resorption markers, suggesting that the reduced response to combination therapy in comparison to PTH might be due to the dominating effects of the antiresorptive agent on bone dynamics (Fig. 5). Under other circumstances, combination therapy conceivably could be more beneficial than monotherapy. For example, Deal et al. in a short, 6-month study have reported that the combination of teriparatide and raloxifene has in some respects more beneficial effects than monotherapy with teriparatide alone [117]. Bone formation as assessed by P1NP increased similarly in both groups, but bone resorption as measured by CTX was

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Figure 5

The effect of combination therapy with PTH(1-84) and alendronate on bone turnover markers. The data show changes in P1NP and CTX in response to therapy with PTH(1-84) or alendronate alone or together (reproduced from reference 115 with permission).

reduced in the combination group. BMD increased to a similar extent in the lumbar spine and femoral neck in both groups, but the increase in total hip BMD was significantly greater in the combination group. One reason why combination therapy with alendronate does not appear to provide more beneficial results than monotherapy with PTH alone may be related to the impression that the effects of PTH on bone formation depend, at least in part, on its ability to increase bone resorption. As noted earlier, PTH may depend upon the bone remodeling cycle in which there is a pool of committed osteoblasts. If bone turnover and bone formation are reduced, as for example in the presence of a powerful antiresorptive, PTH may be less effective since it cannot access this pool of osteoblasts. This is consistent with the idea that some of the improvement in bone mass requires the release of IGF-I and other growth factors from the skeleton by osteoclast action. Perhaps the reason why a less potent antiresorptive agent like estrogen or raloxifene does not impede the responsiveness of bone to PTH is because bone resorption is not inhibited to the same degree as with alendronate, thus still providing for a pool of accessible osteoblasts. On the other hand, it is possible that alendronate itself has a direct effect to inhibit bone formation. Recent gene array studies suggest that alendronate might inhibit expression of bone formation marker genes [118] and biochemical pathways utilized by PTH [119]. Bearing this in mind, one can only speculate how previous or concomitant exposure to another bisphosphonate, risedronate, might affect the subsequent or concomitant response to PTH. It is possible that previous risedronate use may not impair subsequent effects of teriparatide on BMD to the same extent as alendronate because risedronate does not reduce bone turnover

to the extent that alendronate does. Risedronate also may be released from the bone surface after discontinuation more quickly than in the case of alendronate [120]. Besides alendronate, no clinical information currently exists regarding interactions between risedronate and other bisphosphonates and parathyroid hormone. It should also be borne in mind that the data that are available with regard to combination therapy utilize surrogate markers, such as BMD and bone turnover markers. Whether combining PTH with an antiresorptive drug is more efficacious in preventing fragility fractures remains to be seen. 1. Combination Therapy with Teriparatide and an Antiresorptive Agent in Glucocorticoid-induced Osteoporosis

Glucocorticoid-induced osteoporosis (GIO) is characterized by prolonged suppression of bone formation and transient increases in bone resorption. A secondary increase in PTH is no longer considered by many to be of major pathophysiologic importance in GIO [121]. Hence, the use of PTH in combination with an antiresorptive to treat GIO has an appealing rationale. Lane et al. conducted a 12-month, randomized, controlled trial of 51 postmenopausal women on previous hormone replacement therapy and glucocorticoids (>5 mg/day prednisone), who were randomized to teraparatide for one year or not (placebo injections were not used). By QCT, vertebral bone density increased by 35% and by DXA, 11%. Total hip BMD increased by 2% in the PTH group, while there was no difference in forearm BMD between groups. Bone markers showed increased bone formation in the first 3 months, while resorption markers peaked, later, at 6 months [122]. Buxton et al. have implicated changes in RANK L, OPG,

640 IL-6, and IL-6sR in the processes that lead to osteoblast stimulation by PTH in the presence of glucocorticoids [123]. Question 3: What are the Consequences of Ending PTH Therapy? Is an Antiresorptive Agent Necessary?

In the countries in which teraparatide has been approved for therapy of osteoporosis, a short 18–24-month treatment period is recommended. Obvious concerns follow thereafter with regard to the consequences of discontinuing therapy. Most of the published data pertaining to this concern are observational trials. The pivotal phase III trial of teriparatide [60] (see above) was followed by an observational, follow-up period in which study subjects were given the option of switching to a bisphosphonate or not taking any further medications. Of the original cohort in the Neer et al. study [60], 77% of the women were followed. A majority (60%) was treated with antiresorptive therapy after discontinuation of PTH. Three groups were identified: one set of study subjects elected to begin antiresorptive therapy immediately. In these individuals, the gains in bone density were maintained. A second group of subjects did not begin antiresorptive therapy until 6 months after discontinuation of teriparatide. In these individuals, there were major reductions in bone density during these 6 months but as soon as antiresorptive therapy was instituted, there were no further reductions in bone density. A third group of subjects elected not to take any therapy for the duration of the observational 30-month follow-up period. Reductions in bone density were progressive throughout. Despite these data from bone mass measurements, the effect of previous therapy with teriparatide and/or subsequent therapy with a bisphosphonate to prevent fractures persisted for as long as 31 months after the discontinuation of teraparatide treatment [124]. Non-vertebral fragility fractures were reported by proportionately fewer women previously treated with PTH (with or without a bisphosphonate) as compared with those treated with placebo (P < 0.03). Similarly, changes from baseline in lumbar spine BMD remained significantly higher in the original PTH-treated groups. The data, however, do not indicate whether the administration of bisphosphonate was important in the subsequent maintenance of bone mass and fracture reduction. In a logistic regression model, bisphosphonate use for 12 months or longer was said to add little to overall risk reduction of new vertebral fractures in this post-treatment period. It is hard to be sure of this conclusion because the data were not actually analyzed separately into those who did or did not follow teriparatide treatment with antiresorptive therapy. In the study of Rittmaster et al., all subjects were treated with alendronate after PTH(1-84) and showed no change or increases in BMD. Bone turnover declined but still

John P. Bilezikian and Mishaela R. Rubin

remained above that of the original placebo group [125]. However, these studies did not have an arm in which the antiresorptive was discontinued. In addition there were no fracture data available in the post-treatment period with PTH(1-84). In the studies of Lindsay et al. and Lane et al. the 3-year or 1-year period of teriparatide therapy respectively was followed by continuation of estrogen use [95, 126, 127]. In the following year, bone density was well maintained. In neither of these studies did the protocol include a group of subjects who were withdrawn from estrogen also. One cannot be sure, therefore, that bone density would have been maintained had the antiresorptive been discontinued. In the study of Kurland et al., men were given a choice between following teriparatide therapy with or without an antiresorptive [128]. Those who chose the bisphosphonate showed further increases of 5% in the lumbar spine over the next 12 months. Those who chose no further treatment had progressive losses, up to 3.7%, in lumbar spine bone density. When the entire period of teriparatide therapy (18 months) and either bisphosphonate or no therapy (12 months) was reviewed, the changes between those who did or did not receive a bisphosphonate was even more dramatic. Among the men who followed teriparatide therapy with a bisphosphonate, the cumulative gain was over 16%; in the men who did not follow teriparatide therapy with a bisphosphonate, the cumulative gain was reduced to only 4% [128]. Men who delayed the start of bisphosphonate therapy until 1 year after teriparatide discontinuation had gains in lumbar spine BMD over the next year, but these gains only matched the loss they had experienced in the previous year. In a larger study of male osteoporosis, 355 of the original cohort of 437 men who were treated with teriparatide, 20 or 40 µg daily, were followed up for 30 months [71]. The decline in BMD after PTH discontinuation was prevented by antiresorptive treatment. Here again, however, it is difficult to know whether the post-teriparatide use of bisphosphonate was instrumental in demonstrating antifracture efficacy of teriparatide. The relatively small numbers of subjects in the groups that either received bisphosphonate or did not, precluded a specific analysis of the effect of bisphosphonate per se. In all the studies summarized above, the follow-up design was observational and, in many cases, control subjects who were followed without antiresorptive therapy were either not available or not analyzed separately. The PaTH study has provided data in a rigorously controlled, blinded fashion to address the need for post-PTH antiresorptive therapy [129]. According to the prospective design, subjects who had received PTH(1-84) for 12 months were randomly divided into two groups. One group received alendronate,

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10 mg daily, and the other group received a placebo tablet for an additional 12 months. In the subjects who received alendronate, there was a further 4.9% gain in lumbar spine BMD while those who received placebo experienced substantial declines [129]. The results of this study definitively establish the importance of following PTH or teraparatide therapy with an antiresorptive. In the study of Black et al. bone markers fell whether the subjects were treated with placebo or alendronate in the post PTH(1-84) period. The reduction in bone turnover after PTH(1-84) was discontinued is probably based upon a different mechanism than the reduction in bone turnover when alendronate follows PTH(1-84). When bone turnover markers fall after PTH(1-84) in the presence of alendronate, it is likely that bone resorption falls more quickly than reductions in bone formation. This difference in time course should provide a post-PTH “window” during which bone formation is favored. On the other hand, when PTH is stopped without further antiresorptive therapy, it is likely that bone formation declines faster than bone resorption because PTH has been primarily driving processes associated with bone formation. Under these circumstances, bone resorption will predominate and BMD will fall. A depiction of this concept is shown in Figure 6. Further mechanistic explanations follow from this discussion about why the anabolic effects of PTH on bone mass acquisition are lost so quickly after the drug is discontinued. Teriparatide stimulates bone formation, albeit with lower mineralization density initially, enabling increased secondary mineralization and maturation of new bone when bisphosphonate is subsequently used. Without antiresorptive therapy, bone that is induced by PTH actions is

Figure 6

Changes in bone density after PTH. After therapy with parathyroid hormone is discontinued, bone markers are reduced whether or not antiresorptive therapy is instituted. The diagram depicts the concept that when PTH is discontinued without antiresorptive therapy, bone formation markers fall more rapidly leading to a decrease in bone density. When PTH is followed by antiresorptive therapy, bone resorption markers fall more rapidly leading to an increase in bone density.

likely to be rapidly resorbed, particularly if bone resorption is still increased in the early post-PTH period. In patients treated with PTH, there is evidence that new bone matrix is not yet fully mineralized because of the high rate of bone turnover [130]. Using quantitative backscattered electron imaging, Roschger et al. have shown that a greater than normal amount of matrix is found at lower mineralization densities because of the higher amounts of newly formed bone that have not yet had time to undergo complete secondary mineralization [130]. This observation is exactly opposite to mineralization density using the same technique applied to an antiresorptive agent, like alendronate. This is further illustrated by the resolution of cortical porosity that occurs after PTH withdrawal in ovariectomized cynomolgus monkeys [25]. This delay provides an explanation for the observation that apparent bone density, as measured by DXA, increases following parathyroidectomy [131]. As the bone turnover rate is decreased, more time is available for secondary mineralization, allowing for continued maturation of newly formed mineral that was rapidly deposited during ongoing PTH treatment but required further consolidation [127], resulting in an increase in apparent bone density.

G. PTH and Osteosarcoma Long-term studies with high-dose teriparatide administration to 6-week-old Fisher rats for 24 months demonstrate a dose-related appearance of osteogenic sarcoma. This effect is related not only to dose but also to duration of use, consistent with lifetime exposure in a growing rodent to about 75 human year equivalents. In a second rat toxicity study with teriparatide, a “no effect” dose was defined, still larger and with greater exposure than any human being will ever experience. No neoplasms were found when a 5 U/kg treatment was initiated at 6 months of age and continued for up to 70% of the rat life span [132]. Vahle et al. have reported the details of the experimental protocol for the toxicity studies with teriparatide [133] and Tashjian and Chabner [134] have provided a comprehensive commentary on issues related to human safety of this drug. Osteosarcoma has also been reported in a similar carcinogenicity study with PTH(1-84). Although there was no difference in PTH(1-84) at the dose of 10 µg/kg/d compared to controls, there was a dose-related incidence of osteosarcoma in the mid (50 µg) and high (100 µg) dose groups over 2 years [135]. There is great uncertainty about whether this toxicity study in a rodent model has relevance to human physiology. The rat responds to PTH with an exaggerated anabolic response, perhaps because the rat does not have an osteon

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remodeling system. In fact, the rat skeleton is in a rather constant state of modeling with growth through open epiphyses occurring essentially thoughout life. All non-human primate studies to date have not shown the occurrence of osteogenic sarcoma in monkeys exposed to protocols that are similar to the rat studies. Moreover, among the millions of individuals who have been exposed to chronically elevated levels of PTH in primary or seconday hyperparathyroidism or parathyroid cancer, there has been only one reasonably well-documented report of osteogenic sarcoma [136]. Considering the millions of individuals world-wide who have primary hyperparathyroidism, it is surprising, in fact, that so few cases have been reported. One might have expected more, simply on chance association alone. Further, there have been no reports of osteogenic sarcoma among any of the patient cohorts in any of the clinical trials with PTH, amounting to over 2500 patients [137]. It is reasonable to assume that teriparatide and PTH(1-84) are safe when administered to human subjects. However, the rat toxicity data have induced a cautionary note by the FDA, a “black box” warning on the use of the drug [138]. The black box warning is not a feature of the registration of teriparatide in other countries/regions such as Europe, South Africa, Brazil, Australia, and Hong Kong. Contraindications to the use of teriparatide include the following: any hypercalcemic condition, osteosarcoma, metastatic bone disease, Paget’s disease of bone, pregnancy, X-ray therapy to the skeleton or to soft tissues in which a skeletal port is included, and children.

molecule that antagonizes the parathyroid cell calcium receptor, thus stimulating the endogenous release of PTH [148]. This approach could represent a novel endogenous delivery system for intermittent PTH administration. The use of a calcilytic would have to meet the challenge of rapidly increasing the secretory rate of endogenous PTH for a very short period of time. Also for future consideration is the possibility that parathyroid hormone could be used to accelerate fracture healing, as demonstrated in rodent models [149].

H. Future Considerations

References

In the future, PTH may be modified for easier and more targeted delivery. Oral or transdermal delivery systems may become available. Preliminary data on a transdermal delivery system of PTH(1-34) have been presented [139]. PTH has also been formulated to provide systemic exposure following oral administration in rats and cynomolgus monkeys [140]. An oral delivery formulation of PTH(1-34) is being evaluated in a phase II clinical trial [141]. A bioactive recombinant PTH analog was also presented as a potential future oral agent [142]. A cyclic PTH(1-31) is under investigation [143] as well as PTHrP(1-36) [144]. Less frequent administration of PTH, such as once weekly, might also be an effective treatment option [145–147]. It is also possible that repeated, shorter cycles of PTH treatment followed by bisphosphonates, while bone markers are still elevated (closer to the 1-year point, rather than the 3-year point), might be even more effective [114]. Apart from forms and ways to administer exogenous PTH, Gowen et al. have described an oral calcilytic

V. CONCLUSIONS The advent of anabolic therapy in the form of teriparatide is revolutionizing our approach to the therapy of osteoporosis. For the first time, a drug is available that not only improves bone density, features of bone turnover, and reduces fracture incidence, but it also significantly improves microarchitectural and geometric properties of bone. These changes in bone qualities are attractive considering the goal of therapy for osteoporosis, namely to improve the basic underlying abnormalities that give rise to skeletal fragility. Clearly, we are on the threshold of a new paradigm in the treatment of osteoporosis. For women and for men, we can look forward to a period of unprecedented further advances in our approach to a disease that is surely to affect us all, if we live long enough.

1. Lindsay, R., and Cosman, F. (1996). The pharmacology of estrogens in osteoporosis. In “Principles of Bone Biology” (J. P. Bilezikian, L. G. Raisz, and G. A. Rodan, eds), pp. 1063–1068. Academic Press, San Diego, CA. 2. Fleisch, H. (1996). Bisphosphonates: mechanisms of action and clinical use. In “Principles of Bone Biology” (J. P. Bilezikian, L. G. Raisz, and G. A. Rodan, eds), pp. 1037–1052. Academic Press, San Diego, CA. 3. Azria, M., and Avioli, L. (1996). Calcitonin. In “Principles of Bone Biology” (J. P. Bilezikian, L. G. Raisz, and G. A. Rodan, eds), pp. 1083–1098. Academic Press, San Diego, CA. 4. Fogelman, I., Ribot, C., Smith, R., Ethgen, D., Sod, E., and Reginster, J. Y. (2000). Risedronate reverses bone loss in postmenopausal women with low bone mass: results from a multinational, double-blind, placebocontrolled trial. BMD-MN Study Group. J. Clin. Endocrinol. Metab. 85, 1895–1900. 5. Kurland, E. S., Rosen, C. J., Cosman, F., McMahon, D., Chan, F., Shane, E., Lindsay, R., Dempster, D., and Bilezikian, J. P. (1997). Insulin-like growth factor-I in men with idiopathic osteoporosis. J. Clin. Endocrinol. Metab. 82, 2799–2805. 6. Greenspan, S. L., Parker, R. A., Ferguson, L., Rosen, H. N., MaitlandRamsey, L., and Karpf, D. B. (1998). Early changes in biochemical markers of bone turnover predict the long-term response to alendronate therapy in representative elderly women: a randomized clinical trial. J. Bone Miner. Res. 13, 1431–1438.

643

Chapter 38 Monitoring Anabolic Treatment 7. Chesnut, C. H., Skag, A., Christiansen, C., Recker, R., Stakkestad, J. A., Hoiseth, A., Felsenberg, D., Huss, H., Gilbride, J., Schimmer, R. C., and Delmas, P. D. (2004). Effects of oral ibandronate administered daily or intermittently on fracture risk in postmenopausal osteoporosis. J. Bone Min. Res. 19, 1241–1249. 8. Delmas, P. D., Recker, R. R., Chesnut, D. H., Skag, A., Stakkestad, J. A., Emkey, R., Gilbride, J., Schimmer, R. C., and Christiansen, C. (2004). Daily and intermittent oral ibandronate normalize bone turnover and provide significant reduction in vertebral fracture risk: results from the BONE study. Osteoporosis Int. 15, 792–798. 9. Manolagas, S. C. (2000). Birth and death of bone cells: basic regulatory mechanisms and implications for the pathogenesis and treatment of osteoporosis. Endocr. Rev. 21, 115–137. 10. Plotkin, L. I., Weinstein, R. S., Parfitt, A. M., Roberson, P. K., Manolagas, S. C., and Bellido, T. (1999). Prevention of osteocyte and osteoblast apoptosis by bisphosphonates and calcitonin. J. Clin. Invest. 104, 1363–1374. 11. Seeman, E. (2003). Bone quality. Osteoporos. Int. 14, (Suppl 5), 3–7. 12. Rubin, M. R., and Bilezikian, J. P. (2004). Anabolic therapy of osteoporosis in women and in men. In “Principles of Gender-Specific Medicine” (M. L. Legato, ed), pp. 995–1009. Elsevier, Academic Press, San Diego, CA. 13. Albright, F., Aub, J. C., and Bauer, W. (1934). Hyperparathyroidism: a common and polymorphic condition as illustrated by seventeen proven cases from one clinic. JAMA 102, 1276–1287. 14. Bilezikian, J. P., and Silverberg, S. J. (2004). Management of asymptomatic primary hyperparathyroidism. N. Eng. J. Med. 350, 1746–1751. 15. Silverberg, S. J., Shane, E., De La Cruz, L., Dempster, D. W., Feldman, F., Seldin, D., Jacobs, T. P., Siris, E. S., Cafferty, M., Parisien, M. V., Lindsay, R., Clemens, T. L., and Bilezikian, J. P. (1989). Skeletal disease in primary hyperparathyroidism. J. Bone Min. Res. 4, 283–291. 16. Tam, C. S., Heersche, J. N., Murray, T. M., and Parsons, J. A. (1982). Parathyroid hormone stimulates the bone apposition rate independently of its resorptive action: differential effects of intermittent and continuous administration. Endocrinology 110, 506–512. 17. Lane, N. E., Kimmel, D. B., Nilsson, M. H., Cohen, F. E., Newton, S., Nissenson, R. A., and Strewler, G. J. (1996). Bone-selective analogs of human PTH(1-34) increase bone formation in an ovariectomized rat model. J. Bone Miner. Res. 11, 614–625. 18. Shen, V., Dempster, D. W., Birchman, R., Xu, R., and Lindsay, R. (1993). Loss of cancellous bone mass and connectivity in ovariectomized rats can be restored by combined treatment with parathyroid hormone and estradiol. J. Clin. Invest. 91, 2479–2487. 19. Hodsman, A. B., Kisiel, M., Adachi, J. D., Fraher, L. J., and Watson, P. H. (2000). Histomorphometric evidence for increased bone turnover without change in cortical thickness or porosity after 2 years of cyclical hPTH(1-34) therapy in women with severe osteoporosis. Bone 27, 311–318. 20. Finkelstein, J. S., Klibanski, A., Arnold, A. L., Toth, T. L., Hornstein, M. D., and Neer, R. M. (1998). Prevention of estrogen deficiency-related bone loss with human parathyroid hormone-(1-34): a randomized controlled trial. JAMA 280, 1067–1073. 21. Dempster, D. W., Cosman, F., Kurland, E. S., Zhou, H., Nieves, J., Woelfert, L., Shane, E., Plavetic, K., Muller, R., Bilezikian, J., and Lindsay, R. (2001). Effects of daily treatment with parathyroid hormone on bone microarchitecture and turnover in patients with osteoporosis: a paired biopsy study. J. Bone Miner. Res. 16, 1846–1853. 22. Alexander, J. M., Bab, I., Fish, S., Muller, R., Uchiyama, T., Gronowicz, G., Nahounou, M., Zhao, Q., White, D. W., Chorev, M., Gazit, D., and Rosenblatt, M. (2001). Human parathyroid hormone 1-34 reverses bone loss in ovariectomized mice. J. Bone Miner. Res. 16, 1665–1673. 23. Brommage, R., Hotchkiss, C. E., Lees, C. J., Stancill, M. W., Hock, J. M., and Jerome, C. P. (1999). Daily treatment with human recombinant

24.

25.

26.

27.

28. 29.

30.

31.

32.

33.

34.

35.

36.

37. 38. 39.

parathyroid hormone-(1–34), LY333334, for 1 year increases bone mass in ovariectomized monkeys. J. Clin. Endocrinol. Metab. 84, 3757–3763. Sato, M., Westmore, M., Clendenon, J., Smith, S., Hannum, B., Zeng, G. Q., Brommage, R., and Turner, C. H. (2000). Three-dimensional modeling of the effects of parathyroid hormone on bone distribution in lumbar vertebrae of ovariectomized cynomolgus macaques. Osteoporos. Int. 11, 871–880. Burr, D. B., Hirano, T., Turner, C. H., Hotchkiss, C., Brommage, R., and Hock, J. M. (2001). Intermittently administered human parathyroid hormone(1–34) treatment increases intracortical bone turnover and porosity without reducing bone strength in the humerus of ovariectomized cynomolgus monkeys. J. Bone Miner. Res. 16, 157–165. Jerome, C. P., Burr, D. B., Van Bibber, T., Hock, J. M., and Brommage, R. (2001). Treatment with human parathyroid hormone (1–34) for 18 months increases cancellous bone volume and improves trabecular architecture in ovariectomized cynomolgus monkeys (Macaca fascicularis). Bone 28, 150–159. Ilda-Klein, A., Zhou, H., Lu, S. S., Levine, L. R., Ducayen-Knowles, M., Dempster, D., Nieves, J., and Lindsay, R. (2002). Anabolic action of parathyroid hormone is skeletal site specific at the tissue and cellular levels in mice. J. Bone Miner. Res. 17, 808–816. Hock, J. M. (1999). Stemming bone loss by suppressing apoptosis. J. Clin. Invest. 104, 371–373. Nishida, S., Yamaguchi, A., Tanizawa, T., Endo, N., Mashiba, T., Uchiyama, Y., Suda, T., Yoshiki, S., and Takahashi, H. E. (1994). Increased bone formation by intermittent parathyroid hormone administration is due to the stimulation of proliferation and differentiation of osteoprogenitor cells in bone marrow. Bone 15, 717–723. Jilka, R. L., Weinstein, R. S., Bellido, T., Roberson, P., Parfitt, A. M., and Manolagas, S. C. (1999). Increased bone formation by prevention of osteoblast apoptosis with parathyroid hormone. J. Clin. Invest. 104, 439–446. Bradbeer, J. N., Arlot, M. E., Meunier, P. J., and Reeve, J. (1992). Treatment of osteoporosis with parathyroid peptide (hPTH 1-34) and oestrogen: increase in volumetric density of iliac cancellous bone may depend on reduced trabecular spacing as well as increased thickness of packets of newly formed bone. Clin. Endocrinol. (Oxf) 37, 282–289. Dempster, D., Zhou, H., Cosman, F., Nieves, J., Adachi, J. D., Fraher, L. J., Watson, P. H., Lindsay, R., and Hodsman, A. B. (2001). PTH treatment directly stimulates bone formation in cancellous and cortical bone in humans (abstract). 23rd Annual Meeting of the ASBMR, 1171. Hodsman, A. B., Hanley, D. A., Ettinger, M. P., Bolognese, M. A., Fox, J., Metcalfe, A. J., and Lindsay, R. (2003). Efficacy and safety of human parathyroid hormone-(1–84) in increasing bone mineral density in postmenopausal osteoporosis. J. Clin. Endocrinol. Metab. 88, 5212–5220. Lindsay, R., Zhou, H., Cosman, F., Bostrom, M., Cruz, J., Nieves, J., and Dempster, D. (2003). Short term response to parathyroid hormone (1–34hPTH) in human iliac crest bone using a unique quadruple (double double) tetracycline labeling regimen and single biopsy. J. Bone Miner. Res. (Supp 2), S54. Balena, R., Shih, M. S., and Parfitt, A. M. (1992). Bone resorption and formation on the periosteal envelope of the ilium: a histomorphometric study in healthy women. J. Bone Miner. Res. 7, 1475–1482. Leaffer, D., Sweeney, M., Kellerman, L. A., Avnur, Z., Krstenansky, J. L., Vickery, B. H., and Caulfield, P. J. (1995). Modulation of osteogenic cell ultrastructure by RS-23581, an analog of human parathyroid hormone (PTH)-related peptide-(1–34), and bovine PTH-(1–34). Endocrinology 136, 3624–3631. Augat, P., Fuerst, T., and Genant, H. K. (1998). Quantitative bone mineral assessment at the forearm: a review. Osteoporos. Int. 8, 299–310. Parfitt, A. M. (2002). Parathyroid hormone and periosteal bone expansion. J. Bone Miner. Res. 17, 1741–1743. Parfitt, A. M. (2002). Recent contributions of iliac bone histomorphometry to understanding the anabolic effect of parathyroid hormone.

644

40.

41.

42. 43.

44.

45.

46.

47.

48.

49.

50.

51.

52.

53. 54.

55.

56.

57.

Available online at http://www.bonekey-ibms.org/cgi/content/full/ ibmske;2002017v1, Dobnig, H., and Turner, R. T. (1995). Evidence that intermittent treatment with parathyroid hormone increases bone formation in adult rats by activation of bone lining cells. Endocrinology 136, 3632–3638. Jilka, R. L., Gubrij, I., Ali, A., Wynne, R., Weinstein, R. S., O’Brien, C., and Manolagas, S. C. (2004). Ablation of osteoblasts, but not lining cells, in 3.6Col1a1-tk transgenic mice prevents the anabolic effect of PTH: evidence against the lining cell activation hypothesis. J. Bone Miner. Res. 19(Supp 1), SA527. Noble, B. S., and Reeve, J. (2000). Osteocyte function, osteocyte death and bone fracture resistance. Mol. Cell Endocrinol. 159(1–2), 7–13. Fermor, B., and Skerry, T. M. (1995). PTH/PTHrP receptor expression on osteoblasts and osteocytes but not resorbing bone surfaces in growing rats. J. Bone Miner. Res. 10, 1935–1943. van der Plas, A., Aarden, E. M., Feijen, J. H., de Boer, A. H., Wiltink, A., Alblas, M. J., de Leij, L., and Nijweide, P. J. (1994). Characteristics and properties of osteocytes in culture. J. Bone Miner. Res. 9, 1697–1704. Tanaka, S., Sakai, A., Tanaka, M., Otomo, H., Okimoto, N., Sakata, T., and Nakamura, T. (2004). Skeletal unloading alleviates the anabolic action of intermittent PTH(1–34) in mouse tibia in association with inhibition of PTH-induced increase in c-fos mRNA in bone marrow cells. J. Bone Miner. Res. 19, 1813–1820. Hefferan, T., Evans, G., Zhang, M., Morey-Holton, E., and Turner, C. H. (2004). The anabolic bone response to low dose PTH is attenuated by reduced physical activity. J. Bone Miner. Res. 19(supp 1), SA534. Juppner, H., and Kronenberg, H. M. (2003). Parathyroid Hormone. In Primer on Metabolic Bone Diseases and Disorders of Mineral Metabolism (C. Rosen, ed), ASBMR, pp. 117–124. Rosen, C. J. (2004). Insulin-like growth factor I and bone mineral density: experience from animal models and human observational studies. Best Pract. Res. Clin. Endocrinol. Metab. 18, 423–435. Rosen, C. J. (2003). Insulin-like growth factor I and calcium balance: evolving concepts of an evolutionary process. Endocrinology 144, 4679–4681. Canalis, E., Centrella, M., Burch, W., and McCarthy, T. L. (1989). Insulin-like growth factor I mediates selective anabolic effects of parathyroid hormone in bone cultures. J. Clin. Invest. 83, 60–65. Bikle, D. D., Sakata, T., Leary, C., Elalieh, H., Ginzinger, D., Rosen, C. J., Beamer, W., Majumdar, S., and Halloran, B. P. (2002). Insulin-like growth factor I is required for the anabolic actions of parathyroid hormone on mouse bone. J. Bone Miner. Res. 17, 1570–1578. Sellmeyer, D. E., Palermo, L., Bouxsein, M. L., Bilezikian, J. P., Greenspan, S., Ensrud, K., Black, D., and Rosen, C. J. (2004). Heterogeneity in skeletal response to full-length parathyroid hormone (PTH). J. Bone Min. Res. 19(Suppl I), S96, also submitted for publication, JCEM, 2005. Zeng, G. E. A. (1998). Bone effects in mice of ovariectomy and rhPTH(1-34) are highly strain dependent. Bone 23, 448. Eriksen, E. F., Zeng, Q., Donley, D., and Ma, Y. L. (2004). Local IGF II expression in human bone is greater in women treated with teriparatide than with placebo: a quantitative immunohisotchemical study of bone biopsies from the fracture prevention trial. J. Bone Miner. Res. 19(supp 1), SA159. Wang, Y., Nishida, S., Burghardt, A., Elalieh, H., Majumdar, S., Halloran, B. P., Clemens, T., and Bikle, D. D. (2004). The IGF-I receptor is required for the anabolic actions of parathyroid hormone on bone. J. Bone Miner. Res. 19(supp 1), SA531. Hofbauer, L. C., and Heufelder, A. E. (2000). The role of receptor activator of nuclear factor-kappaB ligand and osteoprotegerin in the pathogenesis and treatment of metabolic bone diseases. J. Clin. Endocrinol. Metab. 85, 2355–2363. Buxton, E. C., Yao, W., and Lane, N. E. (2004). Changes in serum receptor activator of nuclear factor-kappaB ligand, osteoprotegerin, and interleukin-6 levels in patients with glucocorticoid-induced osteoporosis

John P. Bilezikian and Mishaela R. Rubin

58.

59.

60.

61.

62.

63. 64.

65. 66.

67.

68.

69.

70.

71.

72.

treated with human parathyroid hormone (1–34). J. Clin. Endocrinol. Metab. 89, 3332–3336. Demiralp, B., Chen, H. L., Koh, A. J., Keller, E. T., and McCauley, L. K. (2002). Anabolic actions of parathyroid hormone during bone growth are dependent on c-fos. Endocrinology 143, 4038–4047. Hurley, M. M., Okada, Y., Sobue, T., Zhang, X. Z., Xiao, L., Tanaka, Y., Ito, M., Okimoto, N., Nakamura, T., Coffin, J., Rosen, C. J., and Doetschman, T. (2002). The anabolic effect of parathyroid hormone is impaired in bones of FGF2 null mice. J. Bone Miner. Metab. 17(Suppl 1), 1061. Neer, R. M., Arnaud, C. D., Zanchetta, J. R., Prince, R., Gaich, G. A., Reginster, J. Y., Hodsman, A. B., Eriksen, E. F., Ish-Shalom, S., Genant, H. K., Wang, O., and Mitlak, B. H. (2001). Effect of parathyroid hormone (1–34) on fractures and bone mineral density in postmenopausal women with osteoporosis. N. Engl. J. Med. 344(19), 1434–1441. Schwietert, H. R., Groen, E. W., Sollie, F. A., and Jonkman, J. H. (1997). Single-dose subcutaneous administration of recombinant human parathyroid hormone [rhPTH(1–84)] in healthy postmenopausal volunteers. Clin. Pharmacol. Ther. 61, 360–376. Lindsay, R., Nieves, J., Henneman, E., Shen, V., and Cosman, F. (1993). Subcutaneous administration of the amino-terminal fragment of human parathyroid hormone-(1–34): kinetics and biochemical response in estrogenized osteoporotic patients. J. Clin. Endocrinol. Metab. 77(6), 1535–1539. Fraher, L. E. A. (1993). A comparison of the pharmacokinetics of PTH in healthy young and osteoporotic subjects. J. Bone Miner. Res. 8S:253. Henry, J. G., Mitnick, M., Dann, P. R., and Stewart, A. F. (1997). Parathyroid hormone-related protein-(1–36) is biologically active when administered subcutaneously to humans. J. Clin. Endocrinol. Metab. 82, 900–906. Product Monograph: Forteo. 2004, Eli Lilly: Canada. Body, J. J., Gaich, G. A., Scheele, W. H., Kulkarni, P. M., Miller, P. D., Peretz, A., Dore, R. K., Correa-Rotter, R., Papaioannou, A., Cumming, D. C., and Hodsman, A. B. (2002). A randomized doubleblind trial to compare the efficacy of teriparatide [recombinant human parathyroid hormone (1–34)] with alendronate in postmenopausal women with osteoporosis. J. Clin. Endocrinol. Metab. 87, 4528–4535. Kurland, E. S., Cosman, F., McMahon, D. J., Rosen, C. J., Lindsay, R., and Bilezikian, J. P. (2000). Parathyroid hormone as a therapy for idiopathic osteoporosis in men: effects on bone mineral density and bone markers. J. Clin. Endocrinol. Metab. 85, 3069–3076. Orwoll, E. S., Scheele, W. H., Paul, S., Adami, S., Syversen, U., Diez-Perez, A., Kaufman, J. M., Clancy, A. D., and Gaich, G. A. (2003). The effect of teriparatide [human parathyroid hormone (1–34)] therapy on bone density in men with osteoporosis. J. Bone Miner. Res. 18, 9–17. Hodsman, A. B., Hanley, D. A., Ettinger, M. P., Bolognese, M. A., Fox, J., Metcalfe, A. J., and Lindsay, R. (2003). Efficacy and safety of human parathyroid hormone (1–84) in increasing bone mineral density in postmenopausal osteoporosis. J. Clin. Endocrinol. Metab. 88, 5212–5220. Guyatt, G. H., Cranney, A., Griffith, L., Walter, S., Krolicki, N., Favus, M., and Rosen, C. (2002). Summary of meta-analyses of therapies for postmenopausal osteoporosis and the relationship between bone density and fractures. Endocrinol. Metab. Clin. North Am. 31, 659–679, xii. Kaufman, J. M., Orwoll, E., Goemaere, S., San Martin, J., Hossain, A., Dalsky, G. P., Lindsay, R., and Mitlak, B. H. (2004). Teriparatide effects on vertebral fractures and bone mineral density in men with osteoporosis: treatment and discontinuation of therapy. Osteoporos. Int. Ettinger, M., Greenspan, S., Marriott, T., Hanley, D., Zanchetta, J., Bone, H., and Lindsay, R. (2004). PTH(1-84) prevents first vertebral fracture in postmenopausal women with osteoporosis: results from the

645

Chapter 38 Monitoring Anabolic Treatment

73.

74.

75.

76.

77.

78.

79.

80.

81.

82.

83.

84.

85.

86.

87.

Top Study. American College of Rheumatology Annual Meeting, 2004: p. L17. Cummings, S. R., Karpf, D. B., Harris, F., Genant, H. K., Ensrud, K., LaCroix, A. Z., Black, D. M. (2002). Improvement in spine bone density and reduction in risk of vertebral fractures during treatment with antiresorptive drugs. Am. J. Med. 112, 281–289. Watts, N. B., Cooper, C., Lindsay, R., Eastell, R., Manhart, M. D., Barton, I. P., van Staa, T., and Adachi, J. (2004). Relationship between changes in bone mineral density and vertebral fracture risk associated with risedronate. J. Clin. Dens. 7, 255–261. Rosen, C. J., Hochberg, M. C., Bonnick, S. L., McClung, M., Miller, P., Broy, S., Kagan, R., Chen, E., Petruschke, R. A., Thompson, D. E., and de Papp, A. E. (2005). Treatment with once-weekly alendronate 70 mg compared with once-weekly risedronate 35 mg in women with postmenopausal osteoporosis: a randomized double-blind study. J. Bone Min. Res. 20, 141–151. Melton, L. J., 3rd, Atkinson, E. J., O’Fallon, W. M., Wahner, H. W., and Riggs, B. L. (1993). Long-term fracture prediction by bone mineral assessed at different skeletal sites. J. Bone Miner. Res. 8(10), 1227–1233. Riggs, B. L., and Melton, L. J., 3rd, (2002). Bone turnover matters: the raloxifene treatment paradox of dramatic decreases in vertebral fractures without commensurate increases in bone density. J. Bone Miner. Res. 17, 11–14. Hochberg, M. C., Greenspan, S., Wasnich, R. D., Miller, P., Thompson, D. E., and Ross, P. D. (2002). Changes in bone density and turnover explain the reductions in incidence of nonvertebral fractures that occur during treatment with antiresorptive agents. J. Clin. Endocrinol. Metab. 87, 1586–1592. Cosman, F., Nieves, J., Woelfert, L., Formica, C., Gordon, S., Shen, V., and Lindsay, R. (2001). Parathyroid hormone added to established hormone therapy: effects on vertebral fracture and maintenance of bone mass after parathyroid hormone withdrawal. J. Bone Miner. Res. 16, 925–931. Reeve, J., Meunier, P. J., Parsons, J. A., Bernat, M., Bijvoet, O. L., Courpron, P., Edouard, C., Klenerman, L., Neer, R. M., Renier, J. C., Slovik, D., Vismans, F. J., and Potts, J. T. Jr. (1980). Anabolic effect of human parathyroid hormone fragment on trabecular bone in involutional osteoporosis: a multicentre trial. BMJ 280(6228), 1340–1344. Delmas, P., Licata, A., Crans, G., Chen, P., Misurski, D., Wagman, R., and Mitlak, B. H. (2004). Fracture risk reduction during treatment with teriparatide is independent of pretreatment bone turnover. J. Bone Miner. Res. 19(supp 1), 1170. Dobnig, H., Sipos, A., Jiang, Y., Fahrleitner-Pammer, A., Ste-Marie, L., Gallahger, J. C., Pavo, I., Wang, J., and Eriksen, E. F. (2005). Early changes in biochemical markers of bone formation predict improvements in bone structure during teriparatide therapy. J. Clin. Endocrin. Metab. 90(7), 3970–3977. Eastell, R., Chen, P., and Krege, J. (2004). Identification of responders to teriparatide therapy by procollagen type I N-propeptide (PINP) using the least significant change approach. J. Bone Miner. Res. 19(supp 1), SA434. Kurland, E. S., Cosman, F., Rosen, C. J., Lindsay, R., and Bilezikian, J. (2000). Parathyroid hormone (PTH 1–34) as a treatment for idiopathic osteoporosis in men: changes in bone mineral density, bone markers and optimal duration of therapy (abstract). ASBMR 22nd Annual Meeting, F435. Wehrli, F. W., Saha, P. K., Gomberg, B. R., Song, H. K., Snyder, P. J., Benito, M., Wright, A., and Weening, R. (2002). Role of magnetic resonance for assessing structure and function of trabecular bone. Top Magn. Reson. Imaging 13, 335–355. Genant, H. K., Gordon, C., Jiang, Y., Link, T. M., Hans, D., Majumdar, S., and Lang, T. F. (2000). Advanced imaging of the macrostructure and microstructure of bone. Horm. Res. 54 Suppl 1, 24–30. Parfitt, A. M., Mathews, C. H., Villanueva, A. R., Kleerekoper, M., Frame, B., and Rao, D. S. (1983). Relationships between surface,

88.

89.

90.

91.

92.

93.

94.

95.

96.

97.

98.

99.

100.

101.

volume, and thickness of iliac trabecular bone in aging and in osteoporosis. Implications for the microanatomic and cellular mechanisms of bone loss. J. Clin. Invest. 72, 1396–1409. Silva, M. J., and Gibson, L. J. (1997). Modeling the mechanical behavior of vertebral trabecular bone: effects of age-related changes in microstructure. Bone 21, 191–199. Borah, B., Dufresne, T. E., Chmielewski, P. A., Gross, G. J., Prenger, M. C., and Phipps, R. J. (2002). Risedronate preserves trabecular architecture and increases bone strength in vertebra of ovariectomized minipigs as measured by three-dimensional microcomputed tomography. J. Bone Miner. Res. 17, 1139–1147. Eriksen, E. F., Melsen, F., Sod, E., Barton, I., and Chines, A. (2002). Effects of long-term risedronate on bone quality and bone turnover in women with postmenopausal osteoporosis. Bone 31, 620–625. Borah, B., Dufresne, T. E., Chmielewski, P. A., Johnson, T. D., Chines, A., and Manhart, M. D. (2004). Risedronate preserves bone architecture in postmenopausal women with osteoporosis as measured by three-dimensional microcomputed tomography. Bone 34, 736–746. Recker, R., Masarachia, P., Santora, A., Howard, T., Chavassieux, P., Arlot, M., Rodan, G., Wehren, L., and Kimmel, D. (2005). Trabecular bone microarchitecture after alendronate treatment of osteoporotic women. Curr. Med. Res. Opin. 21, 185–194. Jiang, Y., Zhao, J. J., Mitlak, B. H., Wang, O., Genant, H. K., and Eriksen, E. F. (2003). Recombinant human parathyroid hormone (1–34) [teriparatide] improves both cortical and cancellous bone structure. J. Bone Miner. Res. 18, 1932–1941. Eriksen, E. F. (2002). Primary hyperparathyroidism: lessons from bone histomorphometry. J. Bone Miner. Res. 17 Suppl 2, N95–N97. Lindsay, R., Nieves, J., Formica, C., Henneman, E., Woelfert, L., Shen, V., Dempster, D., and Cosman, F. (1997). Randomised controlled study of effect of parathyroid hormone on vertebral-bone mass and fracture incidence among postmenopausal women on oestrogen with osteoporosis. Lancet 350(9077), 550–555. Mashiba, T., Burr, D. B., Turner, C. H., Sato, M., Cain, R. L., and Hock, J. M. (2001). Effects of human parathyroid hormone (1–34), LY333334, on bone mass, remodeling, and mechanical properties of cortical bone during the first remodeling cycle in rabbits. Bone 28, 538–547. Sato, M., Westmore, M., Ma, Y. L., Schmidt, A., Zeng, Q. Q., Glass, E. V., Vahle, J., Brommage, R., Jerome, C. P., and Turner, C. H. (2004). Teriparatide [PTH(1–34)] strengthens the proximal femur of ovariectomized nonhuman primates despite increasing porosity. J. Bone Miner. Res. 19, 623–629. Zanchetta, J. R., Bogado, C. E., Ferretti, J. L., Wang, O., Wilson, M. G., Sato, M., Gaich, G. A., Dalsky, G. P., and Myers, S. L. (2003). Effects of teriparatide [recombinant human parathyroid hormone (1–34)] on cortical bone in postmenopausal women with osteoporosis. J. Bone Miner. Res. 18, 539–543. Hirano, T., Burr, D. B., Turner, C. H., Sato, M., Cain, R. L., and Hock, J. M. (1999). Anabolic effects of human biosynthetic parathyroid hormone fragment (1–34), LY333334, on remodeling and mechanical properties of cortical bone in rabbits. J. Bone Miner. Res. 14, 536–545. Ejersted, C., Andreassen, T. T., Oxlund, H., Jorgensen, P. H., Bak, B., Haggblad, J., Torring, O., and Nilsson, M. H. (1993). Human parathyroid hormone (1–34) and (1–84) increase the mechanical strength and thickness of cortical bone in rats. J. Bone Miner. Res. 8, 1097–1101. Hyldstrup, L., Jorgensen, J. T., and Gaich, G. A. (2001). Assessment of effects of LY333334 [recombinant human parathyroid horomone (1–34)] on cortical bone using digital X-ray radiogrammetry. 1st Joint Meeting of the International Bone and Mineral Society and the European Calcified Tissue Society, Madrid, p. SC33 S.

646 102. Uusi-Rasi, K., Beck, T., Oreskovic, T. L., Sato, M., Borgado, C., and Zanchetta, J. R. (2002). Teriparatide [rhPTH(1–34)] improves the structural geometry of the hip. J. Bone Miner. Res. 17(supp 1), S208. 103. Mosekilde, L. (1990). Sex differences in age-related changes in vertebral body size, density and biomechanical competence in normal individuals. Bone 11, 67–73. 104. Rehman, Q., Lang, T. F., Arnaud, C. D., Modin, G. W., and Lane, N. E. (2003). Daily treatment with parathyroid hormone is associated with an increase in vertebral cross-sectional area in postmenopausal women with glucocorticoid-induced osteoporosis. Osteoporos. Int. 14, 77–81. 105. Semanick, L., Uusi-Rasi, K., Zanchetta, J. R., et. al., (2003). Teriparatide [rhPTH(1–34)] treatment improves the structure of the proximal femur in women with osteoporosis. J. Bone Miner. Res. 18(supp 1), S40. 106. Meunier, P. J., and Boivin, G. (1997). Bone mineral density reflects bone mass but also the degree of mineralization of bone: therapeutic implications. Bone 21, 373–377. 107. Boivin, G. Y., Chavassieux, P. M., Santora, A. C., Yates, J., and Meunier, P. J. (2000). Alendronate increases bone strength by increasing the mean degree of mineralization of bone tissue in osteoporotic women. Bone 27, 687–694. 108. Boivin, G., and Meunier, P. J. (2002). Changes in bone remodeling rate influence the degree of mineralization of bone. Connect. Tissue Res. 43(2–3), 535–537. 109. Hernandez, C. J., Beaupre, G. S., Keller, T. S., and Carter, D. R. (2001). The influence of bone volume fraction and ash fraction on bone strength and modulus. Bone 29(1), 74–78. 110. Roschger, P., Rinnerthaler, S., Yates, J., Rodan, G. A., Fratzl, P., and Klaushofer, K. (2001). Alendronate increases degree and uniformity of mineralization in cancellous bone and decreases the porosity in cortical bone of osteoporotic women. Bone 29, 185–191. 111. Borah, B., Ritman, E. L., Dufresne, R. E., Liu, S., Chmielewski, P. A., Jorgensen, S. M., Reyes, D. A., Turner, R. T., Phipps, R. J., Manhart, M. D., and Sibonga, J. D. (2004). Five year risedronate therapy normalizes mineralization. J. Bone Min. Res. 19(Suppl 1), S308. 112. Roe, E., Sanchez, S., del Puerto, G., Pierini, E., Bacchetti, P., Cann, C., and Arnaud, C. D. (1999). Parathyroid hormone 1–34 (hPTH 1–34) and estrogen produce dramatic bone density increases in postmenopausal osteoporosis – results from a placebo-controlled randomized trial. J. Bone Min. Res. 14(suppl 1), S137. 113. Ettinger, B., San Martin, J., Crans, G., and Pavo, I. (2004). Differential effects of teriparatide on BMD after treatment with raloxifene or alendronate. J. Bone Miner. Res. 19(5), 745–751. 114. Cosman, F., Nieves, J., Luckey, M., Zion, M., Woelfert, L., and Lindsay, R. (2003). Daily vs cyclic PTH combined with alendronate vs alendronate alone for treatment of osteoporosis. Bone Miner. 18, S32. 115. Black, D., Greenspan, S., Ensrud, K., Palmero, L., McGowan, J., Lang, T., Garnero, P., Bouxsein, M., Bilezikian, J., and Rosen, C. (2003). The effects of PTH and alendronate alone or in combination in postmenopausal osteoporosis. N. Engl. J. Med. 349, 1207–1215. 116. Finkelstein, J. S., Hayes, A., Hunzelman, J. L., Wyland, J. J., Lee, H., and Neer, R. M. (2003). The effects of parathyroid hormone, alendronate, or both in men with osteoporosis. N. Engl. J. Med. 349, 1216–1226. 117. Deal, C., Omizo, M., Schwartz, E., Eriksen, E. F., Cantor, P., Wang, J., Glass, E. V., Myers, S., and Krege, J. (2004). Raloxifene in combination with teriparatide reduced teriparatide-induced stimulation of bone resorption but not formation in postmenopausal women with osteoporosis. J. Bone Miner. Res. 19(supp 1), 1169. 118. Onyia, J. E. A. (2002). Gene array analysis of the bone effects of raloxifene and alendronate show that alendronate strongly inhibits the expression of bone formation marker genes. J. Bone Miner. Res. 17(supp 1), S157.

John P. Bilezikian and Mishaela R. Rubin 119. Katz, R., Sun, Q., and Bilezikian, J. P. (2004). Bisphosphonates differentially affect osteoblast survival in vitro. J. Bone Min. Res. 19(Suppl 1), S477. 120. Watts, N., McClung, M., and Olsynski, W., (2004). Effects of risedronate discontinuation on bone turnover and bone mineral density in postmenopausal women with osteoporosis. J. Clin. Densitom. 7, 37. 121. Rubin, M. R., and Bilezikian, J. P. (2002). Clinical review 151: The role of parathyroid hormone in the pathogenesis of glucocorticoid-induced osteoporosis: a re-examination of the evidence. J. Clin. Endocrinol. Metab. 87, 4033–4041. 122. Lane, N. E., Sanchez, S., Modin, G. W., Genant, H. K., Ini, E., and Arnaud, C. D. (1998). Parathyroid hormone treatment can reverse corticosteroid-induced osteoporosis. Results of a randomized controlled clinical trial. J. Clin. Invest. 102, 1627–1633. 123. Buxton, E., Yao, W., and Lane, N. (2004). Changes in serum receptor activator of nuclear factor-kB ligand, osteoprotegerin, and interleukin-6 levels in patients with glucocorticoid-induced osteoporosis treated with human parathyroid hormone (1–34). J. Clin. Endocrinol. Metab. 89, 3332–3336. 124. Lindsay, R., Scheele, W. M., Neer, R., Pohl, G., Adami, S., Mautalen, C., Reginster, J-I., Stepan, J. J., Myers, S. L., and Mitlak, B. H., (2004). Sustained vertebral fracture risk reduction after withdrawal of teriparatide in postmenopausal women with osteoporosis. Arch. Intern. Med. 164, 2024–2030. 125. Rittmaster, R. S., Bolognese, M., Ettinger, M. P., Hanley, D. A., Hodsman, A. B., Kendler, D. L., and Rosen, C. J. (2000). Enhancement of bone mass in osteoporotic women with parathyroid hormone followed by alendronate. J. Clin. Endocrinol. Metab. 85, 2129–2134. 126. Lindsay, R., Scheele, W. H., Clancy, A. D., Hossain, A., and Mitlak, B. H. (2002). Reduction in Nonvertebral Fragility Fractures and Increase in Spinal Bone Density Is Maintained 31 Months after Discontinuation of Recombinant Human Parathyroid Hormone (1–34) in Postmenopausal Women with Osteoporosis. 84th Annual Meeting of the Endocrine Society, San Francisco, CA, pp. OR35–36. 127. Lane, N. E., Sanchez, S., Modin, G. W., Genant, H. K., Pierini, E., and Arnaud, C. D. (2000). Bone mass continues to increase at the hip after parathyroid hormone treatment is discontinued in glucocorticoidinduced osteoporosis: results of a randomized controlled clinical trial. J. Bone Miner. Res. 15, 944–951. 128. Kurland, E. S., Heller, S. L., Diamond, B., McMahon, D. J., Cosman, F., and Bilezikian, J. P. (2004). The importance of bisphosphonate therapy in maintaining bone mass in men after therapy with teriparatide [human parathyroid hormone(1–34)]. Osteoporos. Int. 15, 992–997. 129. Black, D. M., Bilezikian, J. P., Palmero, L., Hue, T., Ensrud, K., Greenspan, S., Lang, T., McGowan, J., and Rosen, C. (2005). The effect of one year of alendronate following one year of PTH1-84: second year results from the PTH and alendronate (PaTH) trial. N. Eng. J. Med. 353(6), 555–565. 130. Misof, B. M., Roschger, P., Cosman, F., Kurland, E. S., Tesch, W., Messmer, P., Dempster, D. W., Nieves, J., Shane, E., Fratzl, P., Klaushofer, K., Bilezikian, J., and Lindsay, R. (2003). Effects of intermittent parathyroid hormone administration on bone mineralization density in iliac crest biopsies from patients with osteoporosis: a paired study before and after treatment. J. Clin. Endocrinol. Metab. 88, 1150–1156. 131. Dempster, D. W., Parisien, M., Silverberg, S. J., Liang, X. G., Schnitzer, M., Shen, V., Shane, E., Kimmel, D. B., Recker, R., Lindsay, R., and Bilezikian, J. P. (1999). On the mechanism of cancellous bone preservation in postmenopausal women with mild primary hyperparathyroidism. J. Clin. Endocrinol. Metab. 84, 1562–1566.

Chapter 38 Monitoring Anabolic Treatment 132. Vahle, J. L., Long, G. G., Sandusky, G., Westmore, M., Ma, Y. L., and Sato, M. (2004). Bone neoplasms in F344 rats given teriparatide [rhPTH(1-34)] are dependent on duration of treatment and dose. Toxicol. Pathol. 32, 426–438. 133. Vahle, J. L., Sato, M., Long, G. G., Young, J. K., Francis, P. C., Engelhardt, J. A., Westmore, M. S., Linda, Y., and Nold, J. B. (2002). Skeletal changes in rats given daily subcutaneous injections of recombinant human parathyroid hormone (1–34) for 2 years and relevance to human safety. Toxicol. Pathol. 30, 312–321. 134. Tashjian, A. H., Jr., and Chabner, B. A. (2002). Commentary on clinical safety of recombinant human parathyroid hormone 1–34 in the treatment of osteoporosis in men and postmenopausal women. J. Bone Miner. Res. 17, 1151–1161. 135. Wilker, C., Jolette, J., Smith, S., Doyle, N., Hardisty, J., Metcalfe, A. J., Marriott, T. B., Fox, J., and Wells, D. (2004). A no observable carcinogenic effect dose level identified in Fischer 344 rats following daily treatment with PTH(1–84) for 2 years: role of the C-terminal PTH receptor? J. Bone Miner. Res. 19(supp 1), SA435. 136. Betancourt, M., Wirfel, K. L., Raymond, A. K., Yasko, A. W., Lee, J., and Vassilopoulou-Sellin, R. (2003). Osteosarcoma of bone in a patient with primary hyperparathyroidism: a case report. J. Bone Miner. Res. 18, 163–166. 137. Palmer, M., Adami, H. O., Krusemo, U. B., and Ljunghall, S. (1988). Increased risk of malignant diseases after surgery for primary hyperparathyroidism. A nationwide cohort study. Am. J. Epidemiol. 127, 1031–1040. 138. FORTEO (teriparatide [rDNA origin] injection). 2002, Eli-Lilly, p. 1, package insert. 139. Gopalakrishnan, V., Hwang, S., Loughre, H., Alexandre, B., Deai, D., Haiying, L., Abribat, T., Vachon, L., and Daddona, P. (2004). Administration of ThPTH to humans using Macroflux transdermal technology results in the rapid delivery of biologically active PTH. J. Bone Miner. Res. 19(supp 1), M484. 140. Leone-Bay, A., Sato, M., Paton, D., Hunt, A. H., Sarubbi, D., Carozza, M., Chou, J., McDonough, J., and Baughman, R. A. (2001). Oral delivery of biologically active parathyroid hormone. Pharm. Res. 18, 964–970.

647 141. Mehta, N. M. (2001). Oral delivery of PTH analogs by a solid dosage formulation. J. Bone Miner. Res. 16(supp 1), S540. 142. Mehta, N. M. (2002). Biological activity of recombinant PTH analog 7841. J. Bone Miner. Metab. 17(Suppl 1), SA 362. 143. Chen, Z., Xu, P., Barbier, J. R., Willick, G., and Ni, F. (2000). Solution structure of the osteogenic 1–31 fragment of the human parathyroid hormone. Biochemistry 39(42), 12766–12777. 144. Horwitz, M. J., Tedesco, M. B., Sereika, S. M., Hollis, B. W., Garcia-Ocana, A., and Stewart, A. F. (2003). Direct comparison of sustained infusion of human parathyroid hormone-related protein(1–36) [hPTHrP-(1–36)] versus hPTH-(1–34) on serum calcium, plasma 1,25-dihydroxyvitamin D concentrations, and fractional calcium excretion in healthy human volunteers. J. Clin. Endocrinol. Metab. 88(4), 1603–1609. 145. Black, D. M., and Rosen, C. J. (2002). Parsimony with PTH: is a single weekly injection of PTH superior to a larger cumulative dose given daily? J. Bone Miner. Res. 17(suppl 1), SA367. 146. Okimoto, N., Tsurukami, H., Okazaki, Y., Nishida, S., Sakai, A., Ohnishi, H., Hori, M., Yasukawa, K., and Nakamura, T. (1998). Effects of a weekly injection of human parathyroid hormone (1–34) and withdrawal on bone mass, strength, and turnover in mature ovariectomized rats. Bone 22, 523–531. 147. Fujita, T., Inoue, T., Morii, H., Morita, R., Norimatsu, H., Orimo, H., Takahashi, H. E., Yamamoto, K., and Fukunaga, M. (1999). Effect of an intermittent weekly dose of human parathyroid hormone (1–34) on osteoporosis: a randomized double-masked prospective study using three dose levels. Osteoporos. Int. 9, 296–306. 148. Gowen, M., Stroup, G. B., Dodds, R. A., James, I. E., Votta, B. J., Smith, B. R., Bhatnagar, P. K., Lago, A. M., Callahan, J. F., DelMar, E. G., Miller, M. A., Nemeth, E. F., and Fox, J. (2000). Antagonizing the parathyroid calcium receptor stimulates parathyroid hormone secretion and bone formation in osteopenic rats. J. Clin. Invest. 105, 1595–1604. 149. Alkhiary, Y. (2003). PTH (1–34) enhances experimental fracture healing. J. Bone Miner. Res. 18, S24.

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Chapter 39

Monitoring of Antiresorptive Therapy Christian Meier Tuan V. Nguyen

Endokrinologische Praxis & Labor, University Hospital Basel, Switzerland

Markus J. Seibel

Bone Research Program, ANZAC Research Institute, The University of Sydney, Australia

Bone and Mineral Research Program, Garvan Institute of Medical Research, University of New South Wales, Sydney, Australia

IV. Interpretation of Changes in Bone Turnover Markers References

I. Introduction II. Effects of Pre-treatment Bone Turnover and Mineral Density on Therapeutic Outcomes III. The Role of Markers of Bone Turnover in Monitoring Anti-resorptive Osteoporosis Therapy

I. INTRODUCTION

and/or bone turnover markers reliably predict the expected or true reduction in fracture risk?” While this appears to be a straightforward question, one should be aware of a few caveats that need careful consideration. Firstly, the precise mechanisms by which antiosteoporotic agents reduce fracture risk remain largely obscure. Therefore, the biological significance of a change in any given surrogate endpoint is not self-evident and any inference needs to be interpreted with extreme caution. For example, it has become quite clear that changes in BMD after 1–3 years of antiresorptive therapy fail to predict outcomes across the range of antiresorptive agents, and only partly explain the observed effects within one given agent. While the situation is less clear for markers of bone turnover, “bigger” is certainly not always better when it comes to predicting osteoporosis treatment outcomes. In other words: a change in a surrogate marker, even if it goes into the expected direction, does not automatically infer a “response” in the clinical outcome of interest, nor does the absence of a change in a surrogate endpoint mean

The aim of monitoring treatment is to ascertain therapeutic efficacy, to predict therapeutic outcomes, to screen for unwanted drug effects, and to maintain and encourage patient compliance. Many chronic disorders require longterm therapy and, as a consequence, the intended clinical outcome often does not become evident before many years have passed. In these circumstances, monitoring usually resorts to the use of surrogate endpoints. Fleming and DeMets have pointed out that “the effect of a therapeutic intervention on a surrogate endpoint must reliably predict the effect on the actual clinical outcome in order for the surrogate to be considered an acceptable substitute” [1]. In the case of osteoporosis treatment, the most commonly used surrogate endpoints are bone mineral density (BMD) and bone turnover. In most instances, the change in either bone density or bone turnover will be assessed over time, and the main question asked goes like this: “Following the initiation of therapy, do serial measurements of BMD Dynamics of Bone and Cartilage Metabolism

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650 lack of therapeutic efficacy. Secondly, any surrogate endpoint is subject to random variability (‘background noise’). This and the imprecision of the analytical procedure itself has the potential to significantly affect the result of measurement. The following chapter will review the current knowledge on the use of bone turnover markers in monitoring the effects of antiresorptive treatment in osteoporosis. (The effects of anabolic treatments on bone turnover markers are reviewed in Chapter 39.) We will first discuss the effects of pretreatment bone turnover on therapeutic outcomes, and then look at the role of bone turnover markers in monitoring antiresorptive therapies, paying particular attention to the associations between changes in bone turnover and changes in bone mineral density and fracture risk. In a second part, we will review the interpretation of changes in bone turnover markers based upon the concept of biologic mechanism and random error.

II. EFFECTS OF PRETREATMENT BONE TURNOVER AND MINERAL DENSITY ON THERAPEUTIC OUTCOMES A lot of effort has gone into the identification of predictors of osteoporotic fracture risk. In the treatment-naïve patient, age, gender, comorbidity, disability, calcium intake, low body mass index, low bone mass, and prevalent fractures are the most common and powerful predictors of osteoporotic fracture. More recently, it has become evident that accelerated bone turnover is also associated with a higher risk of osteoporotic fracture, independent of age, disability, and bone mass (see Chapter 38). In these studies, patients with low bone mineral density and/or high bone turnover were shown to be at the highest risk for osteoporotic fractures. Determinants of fracture risk also have the potential to affect treatment outcomes, and again, much work has been devoted to establishing the relationship between baseline variables such as age, gender, bone mass or prevalent fracture status, and therapeutic results. Bone turnover and bone markers were no exception, with some evidence suggesting an association between pretreatment bone turnover and therapy-induced changes in bone mineral density. Thus, several studies point to a relationship between the rates of baseline bone turnover and the subsequent change in BMD during different antiresorptive treatment regimens. Using the whole-body retention of a technetium-labeled bisphosphonate to assess the rate of bone turnover, Civitelli et al. [2] demonstrated that after 12 months of treatment with subcutaneous calcitonin, osteoporotic patients with high bone turnover had significantly greater changes in

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lumbar spine (but not in hip) BMD than similar patients with low bone turnover. Two later studies demonstrated that treatment with calcitonin was more effective in women with high bone turnover at baseline when the rate of turnover was assessed using biochemical markers of bone turnover [3, 4]. Similar results suggesting a significant relationship between baseline bone turnover and the response in BMD were reported for alendronate: in postmenopausal women, urinary type I collagen crosslinked N-telopeptide (NTX-I) was significantly correlated with the percent increase in total body, vertebral, and trochanteric BMD after 1–2 years treatment [5]. Another study confirmed that when osteoporotic women were stratified by baseline bone turnover rates, the increase in spine BMD during alendronate therapy was significantly greater in women with high than with low bone turnover (+7.9% vs. +3%) [6]. However, another report suggested that changes in BMD during treatment with alendronate are independent of baseline bone turnover [7]. Several prospective studies have investigated the effectiveness of oral or transdermal estrogen replacement therapy on BMD in early postmenopausal women in relation to pretherapeutic bone turnover [8–11]. When assessed by whole-body retention of technetium-labeled bisphosphonates after 2 years of estrogen treatment, gains in lumbar spine BMD were significantly greater in patients with high bone turnover than in those with low or normal bone metabolism (6.6% vs. 2.7%) [10]. Chesnut et al. [9] and Rosen et al. [8] demonstrated that individuals within the highest quartile of baseline bone turnover experienced the greatest gain in BMD after 6 and 12 months of treatment with calcium and HRT. In this study, baseline urinary NTX-I and serum OC showed the highest predictive value for a change in spinal BMD after 1 year of either HRT or calcium supplementation. In contrast, in a 3-year prospective study on the effects of HRT on spine and hip BMD Stevenson et al. [12] were unable to distinguish between responders and non-responders by means of either pretreatment or follow-up measures in bone turnover. Both groups showed the same baseline values of bone formation and bone resorption, and the change in bone markers in response to HRT was identical in the affected and unaffected groups. Thus, while there is some evidence suggesting an association between pretreatment bone turnover and therapyinduced changes in BMD, little is known whether a similar relationship exists between baseline bone turnover and the reduction in osteoporotic fracture risk during antiresorptive therapy. Such an association would be of great interest as changes in BMD during antiresorptive treatment explain only the smaller part of the effects of therapy on fracture incidence [13]. A recent analysis of the risedronate phase III clinical programs has demonstrated that this bisphosphonate reduces the risk of vertebral fractures in women with

Chapter 39 Monitoring of Antiresorptive Therapy

postmenopausal osteoporosis regardless of pretreatment bone resorption rates [14]. After 3 years of oral risedronate, the relative risk of fracture was reduced by 48% in patients with high bone resorption at baseline, and by 46% in women with low pretreatment bone resorption (Fig. 1). At 1 year, the number of patients needed to treat to avoid one fracture (NNT) was substantially lower in the group of patients with high baseline bone resorption than for their counterparts with normal bone turnover (NNT 15 vs. 25). This result was mostly due to the higher fracture incidence and therefore greater underlying risk in the placebo group with high baseline bone resorption rates, and was less pronounced when analyzing the entire 3-year study period. The findings of this study do not support treating patients solely based on their bone resorption rate, as treatment efficacy was not significantly influenced by baseline bone resorption status.

651 In summary, there is some evidence favoring a relationship between the rate of bone turnover at baseline and the subsequent response of BMD to antiresorptive treatment. When treated with calcitonin, HRT or bisphosphonates, subjects with accelerated pretreatment bone turnover appear to gain BMD at a faster rate than patients with normal or low bone turnover. For HRT, this association appears to holds true only for patients with markedly increased bone turnover rates. The effects of a widely used third-generation aminobisphosphonate, risedronate, appear to be independent of baseline bone resorption. No results have been published so far for other agents with antiresorptive activity. Considering other factors such as the degree of preanalytical variability of bone turnover markers (see Chapter 34), it remains unclear whether the measurement of pretreatment bone turnover should guide treatment decisions in individual patients.

III. THE ROLE OF MARKERS OF BONE TURNOVER IN MONITORING ANTIRESORPTIVE OSTEOPOROSIS THERAPY The major domain for the clinical use of biochemical bone markers is the monitoring of osteoporosis therapy. This application includes keeping an eye on both therapeutic efficacy (i.e. the prediction of the therapeutic response in regards to changes in BMD and reduction in fracture risk) and patient compliance.

A. Effect of Antiresorptive Agents on Markers of Bone Turnover Most antiresorptive agents induce a profound decrease in bone turnover with maximal suppression of bone resorption markers within 1–6 months of treatment, and a somewhat delayed decrease in bone formation markers. The magnitude of suppression in bone markers appears to depend on the dose and potency of the drug chosen as well as the marker used. The following paragraph summarizes the therapeutic efficacy of different antiresorptive regimens. 1. Hormone Replacement Therapy (HRT)

Figure 1

Cumulative incidence of new vertebral fractures by bone resorption status (urinary DPD). (A) Below normative mean (NM). (B) Above normative mean (NM). The dotted lines represent risedronate-treated patients, whereas the solid lines represent patients receiving placebo. (From [13].)

Estrogen deficiency is associated with a significant increase in bone remodeling as reflected by a 50–100% increase in both bone formation and bone resorption markers [15–19]. It is now widely accepted that this acceleration in bone turnover, together with an imbalance between bone resorption and bone formation, is ultimately causing the bone loss observed after the menopause. Many studies,

652 including large randomized placebo-controlled trials evaluating the effect of estrogen on bone turnover have consistently demonstrated that hormone replacement therapy can reverse these changes by decreasing markers of bone turnover into premenopausal levels. Irrespective of the route of administration, a rapid and sustained decrease in bone resorption markers is observed during the initial weeks of HRT, reaching a plateau within 3–6 months. Due to the normal coupling of bone resorption and formation processes, bone formation markers usually follow with a short delay [8, 9, 11, 20–41]. After discontinuation of HRT, markers of bone resorption and, at a later stage, markers of bone formation return to postmenopausal (i.e. elevated) levels [32, 36, 42]. As shown in early postmenopausal women treated with conjugated estrogens (0.625 mg/day), urinary NTX-I levels significantly decrease by 23% as early as 2 weeks after commencing replacement therapy. After 3 months, urinary NTX-I levels decreased by 39% [9], whereas in the same cohort, urinary DPD concentrations showed a less pronounced effect with a mean decrease of 20% [8]. Similar results were reported in the PEPI trial [34]: the numerically greatest suppression after 12 months of HRT was observed for serum type I collagen C-terminal crosslinked telopeptide (CTX-I; −60%) and urinary NTX-I (−50%), while less pronounced effects were seen when deoxypyridinoline (DPD, −30%) or pyridinoline (PYD, −25%) were measured. When looking at these numbers, one should not forget that such a numerical result has little value if not viewed against the background of the marker’s non-specific variability. As a rule, markers with pronounced reductions in response to bisphosphonate therapy (signal) also exhibit the greatest degree of non-specific variability (noise). Calculating the respective signal-to-noise ratios for the various markers can be helpful in distinguishing between spurious and true differences in the magnitude of response (see Chapter 34 and below). Comparable antiresorptive effects have been observed for formulations using alternate routes of administration, including transdermal [24, 30, 40], percutaneous [20], and intranasal [35, 41] estrogen applications. With both conjugated estrogens [38] and transdermal 17β-estradiol [43], a dose-dependent suppressive effect on bone resorption markers has been observed in early postmenopausal women. Lindsay et al. [38] reported on significant reductions in urinary NTX-I levels after 24 months of treatment with either 0.625 mg conjugated equine estrogens (CEE) and 0.3 mg CEE daily. However, the reduction in urinary NTX-I levels was significantly smaller for women taking CEE at 0.3 mg/d (−34%) as compared to women on 0.625 mg/d (−55%). In older postmenopausal women, however, the response of biochemical markers of bone

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turnover was similar for lower or higher estrogen doses [33, 36, 39]. Prestwood et al. [36] observed similar rates of reduction in urinary bone resorption markers (CTX-I, NTX-I, DPD) irrespective whether the daily dose of micronized 17β-estradiol was 0.25 mg (−30–66%), 0.5 mg (−20–85%), or 1.0 mg (−23–71%). Correspondingly, when based on equivalence testing, the effect of 17β-estradiol 0.25 mg/d was similar to that of 1.0 mg/d in decreasing markers of bone turnover, demonstrating that lower than usual doses of estrogen may equally reduce bone resorption and, thereby, prevent bone loss in older postmenopausal women. Compared to bone resorption markers, bone formation markers show a delayed decrease in response to estrogen replacement [8, 34, 38]. Rosen et al. [8] observed a progressive reduction in OC levels with significantly lower levels at 1 month (−15%) and maximal suppression after 12 months of oral HRT (−60%). In contrast, serum bone alkaline phosphatase (BALP) levels increased early during treatment (up to month 3) and then declined with a maximal decrease of 25% after 12 months of HRT (end of study). The differential pattern of change in bone formation markers during the first 3 months of HRT may depend on the route of estrogen administration, taking into account the first-pass effect of oral estrogens on liver proteins such as IGF-I [44]. Much like seen with bone resorption markers, the effect of oral estrogens on bone formation markers appears to be dose- and age-dependent. In early postmenopausal women, the decline in serum OC levels was more pronounced in women treated with CEE 0.625 mg/d as compared to 0.3 mg/d (−37 vs. −22%) [38]. In contrast, no such effect was seen in elderly women treated with micronized 17βestradiol. Specifically, serum levels of BALP (−10–12%), OC (−17–22%), and PINP (−22–34%) decreased to the same extent, irrespective whether the women were treated with either 0.25, 0.5, or 1.0 mg of micronized 17β-estradiol daily [36]. 2. Soy Isoflavones

Phytoestrogenic agents have received some attention due to their presumed positive effects on bone mass and the cardiovascular system. Of the several isoflavones found in soybeans and their derivative foods, genistein has been shown to be the most efficacious [45]. In a recent randomized placebo-controlled study, genistein treatment over 1 year significantly reduced the urinary excretion of DPD (−44%) and of PYD (−42%). The effects on bone resorption markers were similar to those of continuous HRT with 1.0 mg of micronized 17β-estradiol. However, in contrast to the suppressive effects of estrogens on bone formation markers (BALP −20%, OC −22%), 12 months of genistein treatment resulted in an increase in BALP (+25%)

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Chapter 39 Monitoring of Antiresorptive Therapy

and OC (+37%) [46]. The mechanisms leading to these observations remain unclear. 3. Raloxifene

Raloxifene is a benzothiopene-derived selective estrogen receptor modulator (SERM) with estrogen agonistic effects on the skeleton. Studies in different cohorts including early menopausal [47], late postmenopausal [48–50] as well as older, institutionalized women [51] demonstrate that raloxifene induces a sustained decrease in markers of bone resorption and formation, reaching a plateau after 6–12 months of treatment. Delmas et al. [47] reported significant reductions in urinary CTX-I by 34%, BALP by 23%, and OC by 15% after 24 months of continuous treatment with raloxifene 60 mg/d. Two recent studies comparing the effects of raloxifene and estrogen treatment on bone turnover suggest that raloxifene results in a less pronounced reduction in bone markers. Median reductions in serum OC were 27% for raloxifene 60 mg/d, as compared to 47% in estrogen-treated women (CEE 0.625 mg/d). Similarly, serum BALP decreased by 12% in the raloxifene group and by 32% in the CEE-treated group. Urinary CTX-I was significantly suppressed only in women receiving raloxifene 150 mg/d (−25%) or CEE (−60%), but not in women on raloxifene 60 mg/d [52, 53]. 4. Bisphosphonates

Treatment of postmenopausal women, and of men, with bisphosphonates such as alendronate [5, 6, 54–61], risedronate [62–66], ibandronate [67–71], pamidronate [72], and zoledronate [73] results in suppression of bone turnover. As with HRT, bone resorption markers decrease earlier than markers of bone formation, consistent with a direct effect of bisphosphonates on osteoclasts. Several studies have shown that the absolute amount of metabolic suppression achieved by any bisphosphonate depends on the dose, the route of administration and the biochemical marker chosen. Independent of these factors, the effects of bisphosphonates on bone turnover seem more pronounced than those of either HRT [74, 75] or raloxifene [76] (Fig. 2). Most studies would suggest that the suppressive effect of bisphosphonates on bone turnover is “largest” when measured by the amino- and carboxyterminal telopeptides of type I collagen (NTX-I, CTX-I), and that less pronounced effects are observed when using urinary DPD and PYD as resorption markers. Furthermore, treatment with intravenous bisphosphonates leads to a reduction in resorption markers as early as 12–72 hours after the injection [72, 77, 78], while oral bisphosphonates act much slower. For example, postmenopausal women treated with oral alendronate had a 58% decrease in urinary NTX-I levels after 4 weeks of treatment [58], and achieved the maximal

Figure 2

Change in bone turnover markers during treatment with alendronate 70 mg one weekly (■) and raloxifene 60 mg daily (▲) for 12 months. (A) Urinary N-telopeptide of type I collagen (NTX) absolute values, (B) serum bone-specific alkaline phosphatase (BSAP) absolute values. Apparently, the effects of alendronate on both markers are more pronounced than those of raloxifene. However, both antiresorptive agents reduce both bone markers into their respective reference range (area between dashed lines). (From [76].)

response after 6–12 months of treatment. At this point in time, mean reductions were 50–80% for urinary NTX-I, 35–50% for urinary CTX-I, 20–50% for total DPD and PYD, and 20–25% for free DPD and PYD [5, 54–56, 58, 61, 76]. When looking at these numbers, one should not forget that a numerical result alone has little value if not viewed against the background of that marker’s nonspecific variability. As a rule of thumb, markers showing a pronounced response (signal) to bisphosphonate therapy also exhibit the greatest degree of non-specific variability (noise). Calculating the respective signal-to-noise ratios for the various markers can be helpful in distinguishing between spurious and true differences in the magnitude of response (see Chapter 34 and below). True differences in marker response have been attributed to the fact that during bone resorption, collagen is predominantly released in the form of larger peptides. Osteoclasts cultured on human bone slices generate NTX-I and peptide-bound DPD, but not free DPD [79]. This implies that free DPD (and PYD) is not produced directly by osteoclasts, but at a later stage of processing [58].

654 Randall et al. [80] proposed that the proportion of peptidebound crosslink components, such as NTX-I, is increased in high bone turnover states, partly due to a rate-limiting step in their degradation to free crosslinks. Although intermittent bisphosphonate treatment (i.e. with cyclic oral etidronate or intravenous ibandronate) results in a rapid decrease of bone resorption markers, this change tends to be sustained for only a short time and bone resorption markers often return to sub-baseline levels within a few weeks [68, 81]. Due to the coupling of bone resorption and bone formation, markers of bone formation decrease in parallel to markers of bone resorption, but with a delay of several weeks to months. In postmenopausal women treated with alendronate 5 mg/d, a decrease in serum OC by 30–50%, BALP by 35–45%, and PICP by 35–40% has been reported [54, 55, 58, 61]. Similar effects were described in women treated with alendronate 10 mg/d [5, 56, 76] and different doses of oral ibandronate [69]. Upon cessation of most bisphosphonates given for 2 or 3 years, indices of bone turnover increase to attain pretreatment values within 3–12 months [82, 83]. However, following a group of postmenopausal women treated with the highest doses of oral ibandronate, Ravn et al. [69] observed that serum OC and BALP values remained reduced at 12 months after discontinuation of treatment. Similar data were reported for both bone formation and bone resorption markers after a single dose of zoledronate [72], indicating that high-dose bisphosphonate regimens may induce long-term suppression of bone turnover. Bone et al. [84] showed that after 5 years of alendronate treatment, NTX-I levels rise about 20–25% in the first year and then are maintained at the new reduced level for the next 4 years. Several recent studies investigated the effect of combined treatments on bone mass and bone turnover, comparing HRT alone with combinations of HRT with either alendronate [74, 83] or risedronate [85] (Fig. 3). Elderly postmenopausal women treated with HRT (2 mg estradiol) plus alendronate (10 mg/d) had a more pronounced decrease in bone turnover than women on HRT alone; mean reductions after 2 years of treatment were 78–80% vs. 60–63% for uNTX-I, and 67–75% vs. 54–60% for PINP [74]. Similar effects were observed in postmenopausal women treated with conjugated estrogens (CEE 0.625 mg) alone or in combination with risedronate 5 mg/d [85]. Although a significantly greater decrease in bone turnover markers was seen in the combined bisphosphonate-HRT groups, the differences between the various treatment groups were small and, hence, of uncertain clinical significance when it comes to differences in the change of BMD or fracture risk.

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Figure 3

Mean change in urinary N-telopeptide cross-links of collagen type I and mean change in serum bone-specific alkaline phosphatase during different treatment regimens: Placebo/placebo (n = 26 and 27), alendronate/ placebo group (n = 38 and 39), conjugated estrogen/placebo group (n = 72 and 74), combination therapy/combination therapy group (n = 35 and 36), and combination therapy/placebo group (n = 32 and 33). (From [82].)

5. Other Antiresorptive Drugs

Treatment with intranasal [86–91] and oral [92] calcitonin has been shown to significantly decrease bone turnover, albeit to a lesser extent than what has been observed during alendronate treatment [90, 93]. In postmenopausal women and men with idiopathic osteoporosis, intranasal calcitonin significantly reduced urinary DPD levels by 20–40% and urinary NTX-I levels by 20–50%. Effects on bone formation markers were less pronounced and ranged between 5% and 30% [86, 88, 91]. Some studies observed a significant increase in bone resorption markers after 2–3 months of continued calcitonin treatment, suggesting an attenuated effect of calcitonin after prolonged treatment [87, 89]. RANKL and its decoy receptor osteoprotegerin (OPG) are critical regulators of osteoclast differentiation and function. As a soluble member of the TNF receptor family, OPG blocks bone resorption by binding to RANKL and preventing its interaction with RANK, a cell surface receptor on pre-osteoclasts and osteoclasts [94, 95]. A single subcutaneous OPG has been shown to be effective in rapidly and profoundly reducing bone turnover for a sustained period of time in postmenopausal women [96]. The rapid dose-dependent decrease in urinary NTX-I levels (−50–80%) and the delayed decrease in BALP concentrations (−20–50%) indicate that OPG acts primarily to

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decrease bone resorption. Recently, similar dose-dependent antiresorptive effects have been documented in postmenopausal women, using a single subcutaneous dose of a human monoclonal antibody to RANKL [97] (Fig. 4).

B. Association Between Change in Bone Turnover and Change in BMD During Antiresorptive Treatment The ultimate goal in treating patients with osteoporosis is to reduce their fracture risk. However, because of the comparatively low short-term incidence of osteoporotic fractures, monitoring the effect of antiresorptive treatment by fracture incidence alone would be inadequate for most practical purposes. Consequently, serial measurements of changes in BMD as a surrogate marker of therapeutic efficacy are currently the standard approach to monitor osteoporosis therapy. However, changes in BMD occur slowly and therapeutic effects are usually not detectable before several years of treatment. This is partly due to the disadvantageous short-term signal-to-noise ratio of most BMD measurements. Within a year of treatment, comparatively small changes in bone density (2–4%) are contrasted by relatively high precision errors (1–3%), which renders BMD measurements unreliable for short-term assessment

Figure 4

of antiresorptive treatment efficacy [98]. Owing to their significant biological variability, biochemical markers of bone turnover also exhibit a high degree of imprecision and a signal-to-noise ratio comparable to BMD measurements. However, bone markers change much faster in response to therapeutic interventions. In fact, the decrease in bone turnover markers during antiresorptive treatment is inversely related to the subsequent increase in BMD, predominantly at the lumbar spine. Several studies in early postmenopausal women treated with HRT [8, 9, 11, 20, 27, 37, 99–103], except for one [34], have indicated that the degree of short-term reduction in bone turnover markers (after 3–6 months) correlates with the observed long-term increase in BMD at the lumbar spine and/or the mid-radius (after 1–3 years of treatment). One of the first prospective trials investigating the predictive value of changes in bone markers to assess outcome in BMD was the study by Johansen et al. [20]. This placebo-controlled study in postmenopausal women treated with oral or transdermal estrogens has indicated a significant inverse association between the changes in serum OC and the changes of BMD at the forearm and the whole body during 24 months of treatment (r = −0.40 and r = −0.39, respectively). Similar results demonstrating significant correlations with changes in lumbar spine BMD were obtained by others when bone turnover was assessed

The effect of different doses of AMG 162 treatment on bone resorption as reflected by changes in a second morning void urinary NTX/creatinine (nmol BCE/mmol creatinine) over time. Placebo, no symbol; 0.01 mg/kg AMG 162, ❍; 0.03 mg/kg AMG 162, ❒; 0.1 mg/kg AMG 162, ∆; 0.3 mg/kg AMG 162,
; 1.0 mg/kg AMG 162, *; 3.0 mg/kg AMG 162. (From [98].)

656 by serum or urinary CTX-I [37, 101, 103, urinary NTX-I [9, 103], urinary DPD [11], urinary hydroxyproline [99], or by serum osteocalcin [37, 99], bone-specific alkaline phosphatase [100] and bone sialoprotein [103]. Accordingly, early postmenopausal women with no densitometric response to long-term HRT (defined as women whose 5-year BMD change was similar to the mean BMD change of placebo-treated women) were characterized by significantly lower mean changes in total alkaline phosphatase levels during the first months of HRT as compared to densitometric responders [102]. In a study including 236 postmenopausal women treated for 1 year with conjugated equine estrogens, Rosen et al. [8] investigated the association between the change in BMD and bone turnover. Patients were classified according to their bone marker response to HRT, and stratified by quartile of the percent change from baseline to 6 months. For urinary NTX-I, those women with the greatest percent decrease from baseline (−87% to −66%) had the greatest increase in BMD (4.45 ± 0.67 g/cm2), whereas subjects that experienced the least NTX-I response to HRT had the smallest increase in spine BMD (1.36 ± 0.59 g/cm2, P < 0.005). Comparable results were found for free urinary DPD and serum BALP. Based on the same cohort, the areas under the curve derived from ROC analyses of the association between 6-month change in bone marker and the 1-year change in BMD ranged from 0.61 for OC, 0.65 for BALP to 0.79 for NTX-I, indicating that the percent change in urinary NTX-I provided a greater discrimination between gain and loss of BMD than OC or BALP [9]. Using data from two large placebo-controlled trials in which 569 postmenopausal women were treated with various doses of transdermal 17β-estradiol, Delmas et al. [37] found that the percent change and actual value of bone markers under treatment were independent predictors of BMD change. When specificity was set at 90% to have a low rate of false positives, markers of bone resorption (serum and urinary CTX-I) at either 3 or 6 months, and markers of bone formation (OC, BALP) at 6 months were highly predictive of a positive BMD response, with a probability ranging from 0.82–0.91. The corresponding sensitivity ranged from 60% to 68% and was highest in a model combining the percent decrease and the actual value of markers at 6 months. Overall, these studies suggest that short-term changes in bone turnover markers are able to reflect the degree of bone mass responsiveness after 1–3 years of HRT. Similar results have been published for raloxifene [53] and bisphosphonates. Of the latter, most studies were performed with different doses of alendronate in early [57, 61, 104, 105] and late [5, 54, 59, 60, 106, 107] postmenopausal women with or without osteoporosis.

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In contrast to these studies reporting significant correlations between changes in bone markers and BMD after 1 and 4 years, others have observed inconsistent [55] or no [58] such associations. These negative results might be attributed to lower alendronate doses used [55] or to shorter study durations with non-significant changes in BMD (after 6 months) [58]. In a 2-year follow-up study in 85 postmenopausal women treated with alendronate, a marked decrease in turnover was associated with a significant increase in BMD at the lumbar spine. In fact, highly significant negative correlations were observed between percent change in bone markers at 3 months and percent change in BMD after 2 years of treatment for bone formation markers (OC, BALP, PICP; r = −0.63 to −0.67), urinary DPD (r = −0.48), and urinary NTX-I (r = −0.53) [54]. The prediction of long-term response in bone mass during alendronate therapy is not restricted to the lumbar spine, as in other studies changes in bone markers from baseline were also correlated with the increase in BMD at the hip, trochanter, and total body [5, 57, 107]. Greenspan et al. [5] observed in a doubleblind, placebo-controlled trial of alendronate treatment for 2.5 years that at 6 months, women with the greatest drop in urinary NTX-I (65% or greater) had the largest gain in BMD at all sites compared with patients who had smaller decreases in NTX-I (P < 0.05; Fig. 5). Based on linear regression analyses, a 30% decrease in NTX-I at 6 months predicted a BMD increase of 2.8% to 4.1% for the hip regions, and of 5.8% for the spine. Furthermore, Garnero et al. [59] showed that both the absolute serum BALP level and the percent change in BALP at 6 months of treatment are independent predictors of BMD gain (defined as an increase in spine BMD of at least 3% after 2 years). Based upon these data it could be argued that in order to define treatment responsiveness, assessment of the relative change from baseline at 3–6 months may be an inaccurate means to identify BMD responders. The combination of the absolute bone marker level at 6 months (as an estimate of the current bone turnover rate) and the change from baseline (as a measure of treatment efficacy) may improve the prediction of changes in BMD [59] (Fig. 6). Further validation of such models using different bone markers is required before they can be safely applied to clinical practice.

C. Association Between Change in Bone Turnover and Change in Fracture Risk During Antiresorptive Treatment Although several randomized trials have found that antiresorptive agents improve BMD and reduce the risk of

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Chapter 39 Monitoring of Antiresorptive Therapy

Figure 5

Change in BMD at the (A) total hip and trochanter and (B) PA and lateral spine after 2.5 years of alendronate therapy grouped by percentage decreases in NTX at 6 months. Comparison of groups 1–3: total hip, trochanter, PA spine all P < 0.05; lateral spine P = 0.13. (From [4].)

fractures [48, 63, 108–112], recent studies have evidenced that the observed reduction in fracture risk is only partly explained by the documented changes in BMD. The reduction in risk was greater than predicted from improvements in BMD, and it has been estimated that change in BMD explains only 4% to 28% of the reduction in vertebral

Figure 6

fracture risk attributed to antiresorptive treatment [13, 113, 114]. It is therefore possible that changes in other determinants of bone strength, including the rate of bone turnover and its changes during antiresorptive therapy, may be more predictive of antifracture efficacy than changes in BMD. In fact, several studies confirmed that shortterm reductions in bone turnover were associated with a reduction in vertebral and/or non-vertebral fracture risk in women treated with HRT [115], raloxifene [50, 116, 117], risedronate [66], and alendronate [118]. Bjarnason et al. [116] studied the relationship between the 6 and 12 months changes in bone turnover markers and vertebral fracture risk after 3 years of raloxifene treatment in postmenopausal women. A decrease of 9.3 pg/ml in serum OC after 1 year of raloxifene treatment was associated with an odds ratio for new vertebral fractures after 3 years of 0.69 (CI 0.54–0.88, P = 0.003). Similarly, for a decrease of 5.91 µg/L in serum bone alkaline phosphatase the odds ratio was 0.75 (CI 0.62–0.92, P = 0.005; Fig. 7). Importantly, these relationships remained after adjustment for baseline vertebral fracture status and BMD. Two subsequent analyses including postmenopausal women with osteoporosis from the same cohort (MORE trial) extended and confirmed these results showing that both, 1-year percentage changes in serum PINP [50] and OC [117], are able to predict the reduction in vertebral fracture risk after 3 years of treatment. Recently, two studies have been published investigating the change in bone turnover markers and fracture risk in bisphosphonate-treated postmenopausal women [66, 118]. Using data from the VERT studies including postmenopausal women with at least one vertebral fracture,

Prediction of treatment group (alendronate vs. placebo), based on either BALP percent change (from baseline to 6 months of treatment) or BALP level (after 6 months of treatment) or their combination by logistic regression in 307 elderly women with osteoporosis. (From [58].)

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Figure 7 The relative risk of new vertebral fracture (raloxifene vs. placebo) by tertiles of change in serum OC after 6 months and 12 months and in femoral neck BMD after 12 months and 24 months. The cutoffs are indicated below the tertile notification. The p values are for interaction and indicate the presence of a differential antifracture efficacy across tertiles for a model including tertile, therapy and tertile x therapy. (From [116]).

Eastell et al. [66] found that reductions in urinary CTX-I (by 60%) and NTX-I (by 51%) at 3–6 months of risedronate treatment were significantly associated with the reduction in vertebral and non-vertebral fracture risk after 3 years (Fig. 8). The change in bone resorption markers explained more than 50% of the risedronate-related fracture risk reduction for both, vertebral and non-vertebral fractures. Interestingly, there appears to be a threshold level for the decrease in bone resorption markers (e.g., 55–60% as measured by CTX-I, and 35–40% as measured by NTX-I) below which there was no further increase in therapeutic benefit. Bauer et al. [118] reported that in alendronate-treated women, greater reductions in bone turnover were associated with fewer osteoporotic fractures. In their study, each SD reduction in the change in serum BALP at 1 year was associated with fewer spine (odds ratio 0.74; CI: 0.63, 0.87), non-spine (relative hazard [RH] 0.89; CI: 0.78, 1.00) and hip fractures (RH 0.61; CI: 0.46, 0.78) (Fig. 9A). Furthermore, alendronate-treated women with at least a 30% reduction in serum BALP had a lower risk of non-spine (RH 0.72; CI: 0.55, 0.92) and

hip fractures (RH 0.26; CI: 0.08, 0.83) relative to those with reductions < 30% (Fig. 9B). Again, this effect was at least as strong as antifracture effect observed with 1-year change in BMD (118). In summary, it is conceivable that short-term reductions in bone turnover markers after 3–6 months of antiresorptive treatment with bisphosphonates, HRT, or raloxifene are associated with positive long-term responses in BMD. In contrast, patients who show only minimal or no changes in bone markers most likely will not present with favorable BMD outcomes. In addition, and more importantly in terms of fracture risk reduction, changes in bone turnover markers after raloxifene and bisphosphonate therapy are significantly related to subsequent fracture risk with a far greater effect on fracture reduction as what has been attributed to treatment-induced changes in BMD. These data suggest that biochemical markers of bone turnover are useful tools to evaluate therapeutic effects after a relatively short period of time, and that serial measurements of bone markers may help to decide whether or not a patient responds to a specific antiresorptive treatment.

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Figure 8

Relationship between percentage change in bone resorption markers and the incidence of new vertebral fractures. The placebo group is represented by the broken line and the risedronate 5 mg group by the solid line. (From [66].)

D. Monitoring Patient Compliance Using Bone Markers Long-term compliance with treatment for osteoporosis is usually poor [119]. Several studies reported that up to 50% of postmenopausal women were not adherent to their treatment after 1–5 years of HRT [120–124]. A major cause of non-compliance were unwanted side-effects or fear of side-effects, inconvenience caused by medication, and high drug costs [125, 126]. Hence, monitoring patients on antiresorptive medication is an eminent part of patient management in order to improve adherence and persistence to therapy, and ultimately treatment effectiveness. Biochemical markers of bone turnover have been advocated to facilitate follow-up of patients receiving antiresorptive treatments for osteoporosis. As bone turnover markers, in particular indices of bone resorption, decrease rapidly after initiation of treatment within 3–6 months,

they might represent useful surrogate markers for monitoring patient compliance. Only few data, however, are available to support this theoretical approach. Using a decision analysis model, Chapurlat et al. [127] compared two strategies of follow-up: (1) treatment of a woman without specific monitoring, and (2) treatment of this woman with measurement of a serum marker of bone resorption after 3 months of treatment, with change of treatment if response to treatment as assessed by this marker was not satisfactory. It has been suggested that the approach of monitoring osteoporotic women with measurements of bone markers early during treatment course may increase effectiveness of treatment with greater qualityadjusted life years than no follow-up. In a recent study in 75 postmenopausal women treated with raloxifene, Clowes et al. [128] examined whether monitoring (nursemonitoring or marker-monitoring) enhances adherence and persistence with antiresorptive therapy, and whether

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Figure 9

(A) One-year change in bone ALP and hip fracture risk among ALN-treated women. Percent change in bone ALP and predicted risk (log OR) of hip fracture (solid line) and 95% CI (dotted lines) from logistic regression model. Individual data points represented on the x axis. Departure from linearity P value 0.33. (From [118].) (B) Fracture incidence in the placebo and the alendronate treated groups by change in serum BALP at 1 year. Black: placebo group. Gray: alendronate treated women with a change in serum BALP of less than 30% from baseline; white: alendronate treated women with a change in serum BALP of more than 30% from baseline. The graph shows the proportion of placebo and alendronate-treated groups with a fracture during the study. *, P < 0.01 vs. placebo group. (Adapted from [118].)

presenting information on biochemical response to therapy (presentation of results on the percentage change in urinary NTX-I at each visit) provided additional benefit. Survival analyses showed that the monitored group increased cumulative adherence to therapy by 57% compared with no monitoring and there was a trend for the monitored group to persist with therapy for 25% longer. However, presentation of results of effects on NTX-I levels did not improve compliance to therapy compared with nurse-monitoring alone. Nevertheless, results from the IMPACT study in postmenopausal women on risedronate have shown that a reinforcement message based on bone turnover marker response influences long-term

treatment persistence. In patients in whom a verbal feedback on the change of urinary NTX-I was provided, 1-year persistence was higher than in non-reinforced subjects. Interestingly, the message given to patients with a bone turnover marker response considered “good” was associated with significant improvement in persistence, whereas the information given to those with a poor resorption marker response led to a lower persistence [129]. Another large study investigating patient compliance using measurements of urinary CTX-I is under way and should give further evidence whether monitoring osteoporosis treatment using bone turnover markers should be encouraged in clinical practice [130].

Chapter 39 Monitoring of Antiresorptive Therapy

IV. INTERPRETATION OF CHANGES IN BONE TURNOVER MARKERS A. Assessment of Change in an Individual An observed value of a bone turnover marker is made up of two components: the biologic effect (“signal”, “true change”) and random error (“noise”). In the clinical management of osteoporosis, patients are often required to undergo repeated measurements of bone turnover markers. Because of the presence of random error in each measurement, a question of critical interest is whether a change observed in an individual represents a true biologic change or a mere random fluctuation. Currently, there are two approaches to address the question: a distributionbased approach and the Bayesian method. In the former, a range of variation within an individual in repeated measurements is determined to assess whether the change has been “significant”. In the latter, one is interested in making a probabilistic statement of an observed change, such as “given an observed change in a patient, what is the probability that a true change of a certain magnitude has occurred”. In this section, we will consider the two approaches and their assumptions.

B. The Distribution-based Approach For simplicity, a two-measurement case with only one baseline and one follow-up measurement is considered in this section. For any two measurements of bone turnover, a baseline x1 and a follow-up measured value of x2 in an ith subject, an observed change can obviously be estimated by d = x2 − x1. The value of d is considered “significant” if it exceeds the change that may be expected from random fluctuation with a certain probability. To assess the significance of d, we assume that x1 and x2 are normally distributed, and that the variance of x1 and x2 for the subject are the same (denoted by s2). Under these assumptions, it can be shown that the variance of d is: sd2 = s 2 + s 2 − 2 cov( x1 , x 2 ), where cov( x1 , x 2 ) is the covariance between the two measurements. Assuming that x1 and x2 are independent (i.e., no correlation between them, or cov(x1, x2) = 0), the variance of d can be reduced to: sd2 = 2s 2, and the standard deviation of d is: sd = 2 s. (Note that s is also referred to as the standard error of measurement.) Under the assumption of the normal distribution, 1.96 × 2 s (also known as the least significant change, LSC) covers 95% of the random variability of withinsubject differences. In other words, if d / 2s > 1.96,

661 the change between baseline and follow-up measurements can be considered “significant” at the 5% level. However, since 1.64 × 2 s covers 90% of the random variability of within-subject differences, a change of d / 2 s > 1.64 can also be considered “significant” at the 10% level. For example, if the standard error of measurement of serum type I collagen C-terminal telopeptide (sICTP), a marker of bone resorption, is 1 µg/L, and if the baseline and follow-up measured value of sICTP of a patient are 4.5 and 2.0 µg/L, respectively, then the change (−2.5 µg/L) can not be considered “significant” at the 5% level because it falls within the ± 1.96 × 2 × 1 = ± 2.77 µg/L. However, it may be considered “significant” at the 10% level as it falls outside the ±1.64 × 2 × 1 = ± 2.32 µg/L). In clinical medicine, there is no widely accepted standard of the level of confidence (P), and 10 or even 20% is often considered sufficient for decision-making. It is sometimes useful to express the standard error of measurement as percentage of the mean x , such as 100s/ x , which yields the within-subject coefficient of variation (wCV). A proper interpretation of the wCV is that assuming the measurements are normally distributed, 68% of the differences between measurements within an individual lie within wCV% of the true average (or 95% of the differences between measurements within an individual lie within 1.96wCV% of the true average). The LSC can then be expressed in terms of the wCV as follows: LSC = 1.96 2 wCV for the 5% significance level or 1.64 2 wCV for the 10% significance level. Thus, for a bone turnover marker with a wCV of 10%, a change in the marker for any individual would be considered “significant” (at the 5% significance level) if it fell outside the interval ± 28%. The above formulation essentially depends on the assumption that any two consecutive measurements in an individual are unrelated. In reality, of course, this assumption is unlikely to be true because repeated measurements within an individual are usually correlated, and, as a result, the LSC formulation can be rather conservative. That is, the LSC is more likely to underestimate the biologic effect and overestimate the effect of within-subject random variability [131]. An important point to remember is that the LSC can only indicate whether or not an observed change is due to random variability. It doesn’t indicate whether or not the change is clinically significant. The demarcation of “significant” versus “non-significant” change is, as shown above, fairly arbitrary, because it depends on what probabilistic range one is prepared to take. There is no good reason why one has to make a decision based on the 5% instead of the 10% significance level.

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C. The Bayesian Approach The logic behind the LSC formulation is actually derived from the single-subject design counterpart to the t-test of correlated means, which is often used to determine whether a group has changed to a statistically significant degree. The distribution-based method as described above can thus be considered a frequentist method. This method is built with the null hypothesis that there is no significant change, and then work out a statistic that shows the null hypothesis is unlikely (or likely) to be true (i.e., it is a variation of the concept of proof by contradiction). The frequentist inference does not attach probabilities to the data; rather it describes the behaviors of the test statistic and confidence interval under the null hypothesis. It is only when the LSC is used in conjunction with the cut-off point between the functional and dysfunctional distributions that it has any relevance to the determination of clinically significant change. If the measurement error (e.g. analytical and preanalytical variability) is unrelated to the measured value, those subjects with large positive measurement error contributing to their measurements at baseline are likely to have higher baseline measurement, on the average, than those subjects with negligible or large negative error contributing to their measurements. However, it is unlikely that subjects with large positive error in their baseline measurements will also have large positive error in their subsequent measurements, because the errors are independent. Therefore, their follow-up measurements tend to be lower (closer to the mean) than their baseline measurements. The analogous but opposite statement is also true for large negative error in baseline measurement. This means that these high-value individuals may show the smallest changes in response to intervention, a form of “regression toward the mean” phenomenon (for more detail, see Chapter 34). In addition, a decrease or increase in a measured marker may be biologically related to its initial level, i.e. a form of “horse-racing” effect. It is within these contexts that a more meaningful question has to be posed: “Given an observed change, what is the probability that the change reflects a biologic or true effect?” The question can be addressed by the Bayesian approach (for technical details and assumptions, see [132, 133] and Chapter 34). Briefly, the Bayesian approach is based upon the concept that an unknown parameter (such as the mean of change) has a probability distribution, which is referred to as prior distribution. This prior distribution is specified before any new sample data are observed. When the data have been collected, the prior probability is then combined with the new sample data (called “likelihood”) by the Bayes’ theorem to derive a new posterior distribution.

This posterior distribution attaches a probability to different values of the unknown parameter. Thus, the Bayesian approach is a way of synthesis of prior data and existing data from a study to form new data [134]. The application of the Bayesian idea to assess change in an individual can be illustrated by the following narrative example. Consider the data from the patient mentioned above, who has participated in a clinical trial in which baseline and post-intervention measurements of sICTP have been obtained, and the respective values are 4.5 µg/L and 2.0 µg/L (e.g., a 56% reduction). Suppose that it is known that the standard error of measurement (SEM) of an individual sICTP level was 1 µg/L (i.e., the variance of a single measurement is 12 = 1). In order to make an assessment of change for the patient, external information is required. Because the patient was drawn from a clinical trial in which patients with the same clinical characteristics were recruited, it seems reasonable to use the group data as prior information. Suppose that the average and standard deviation (SD) of baseline sICTP in the group were 3.0 and 1.5 µg/L (variance = 1.52 = 2.25), respectively. After the intervention, the overall mean and SD of change in sICTP for the group were −1.5 and 2.6 µg/L (variance = 2.6 2 = 6.76), respectively. The Bayesian estimate of change for the patient can be done in the following steps: (a) The “true” baseline value of sICTP for the patient (denoted by t) can be estimated as a weighted mean of the sample mean (prior data) and the patient’s observed value, where the weight is the sum of inverse variances of the sample data and individual measurement: t = (4.5 /1 + 3/2.25)/(1/1 + 1/2.25) = 4.04 µg/L. The adjusted change for the patient is then: d’ = 2 − 4.04 = −2.04 µg/L. (b) The variance of this adjusted change (denoted by sd2 ') can be shown to be a function of the group and individual variances (see Chapter 34): sd2 ' = (1 × 2.25)/(1 + 2.25) + 1 = 1.69. (c) Next, the Bayesian estimate of change for the patient (denoted by dˆ ) is the weighted average of the sample change and the individual’s adjusted change, where the weight is the sum of the inverse variances of the sample data and the individual’s variance: dˆ = (−2/1.69 − 1.5/6.76)/ (1/1.69 + 1/6.76) = −2.04 µg/L. (d) The variance of dˆ (denoted by sd2ˆ ) is: sd2ˆ = (1.69 × 6.76)/(1.69 + 6.76) = 1.35. Thus, the estimated “true” change in sICTP for the patient is normally distributed with the mean of −2.04 µg/L, and a variance of 1.35 µg/L2 (or SD of 1.35 = 1.16 µg/L). Based on these data, a posterior distribution of sICTP change for the patient can be constructed (Fig. 10). The area under the curve between any two points on the x-axis

Chapter 39 Monitoring of Antiresorptive Therapy

Figure 10

Posterior distribution of change in sICTP for an individual. The figure is constructed such that the total area under the curve is 1. Thus, the area under any two points along the x-axis is the probability that the change in sICTP lies between those limits. For example, the probability that the patient’s sICTP level has decreased by at least 1 mg/L is estimated to be 0.8 or 80%. Moreover, by this method of calculation, the probability that the patient’s sICTP level has decreased by between −1 and −2 mg/L is estimated to be 0.3 or 30%.

is the posterior probability that the change in sICTP lies between those limits. For example, the probability that the patient’s sICTP level has decreased during the follow-up period is 0.96. Because the mean of change was −2.04 µg/L; therefore, the probability that the patient’s sICTP level has decreased by 2.0 µg/L or higher was, as expected, 0.5 or 50%. If a change of 3 µg/L or higher is considered to be clinically relevant, then the probability of such change is 0.18. However, the probability that the patient’s sICTP level has decreased by 1 µg/L or higher was 0.80. Therefore, unlike the frequentist-based LSC, the Bayesian approach allows to make a direct inference on the parameter of interest. The Bayesian approach offers an attractive way to interpret data from a study in the context of external evidence (or judgment) to form conclusions in a manner that fits naturally with the clinical reasoning and decision-making process [135]. The above calculations appear to be complicated at first sight, however, the whole procedure can, in fact, be implemented in a simple computer program or a spreadsheet file.

D. Practical Implications As described in Chapter 34, most bone turnover markers exhibit significant within-subject variability, both in the short and long term. This poses a major problem in the practical use of bone markers for the clinical management of the patient with bone disorders. Whenever a change in a bone marker level is observed in an individual patient (e.g. following an invention), this change must be interpreted

663 against the background of the respective marker’s variability. Therefore, knowledge of the sources of variability and the strategies used to cope with “background noise” are essential for the correct the interpretation of changes in bone markers (see Chapter 34). A number of factors need to be taken into account when employing serial measurements of biochemical markers to determine changes in bone turnover. To minimize some of the limitations linked to pre-analytical and analytical variability, standardized sampling and sample handling are mandatory to obtain reliable results. Controllable factors such as the mode of sample collection, samples handling and storage, diurnal and menstrual rhythms, pre-sampling exercise and pre-sampling diet/fasting should be taken care of wherever possible. As reviewed earlier in this chapter, a reduction in bone turnover after initiation of antiresorptive therapy seems to be associated with a reduction in the observed fracture risk. Therefore, serial measurements of bone turnover during treatment with antiresorptive agents appear to reflect therapy-induced changes in fracture propensity. However, several questions in regard to this association are still open; for example: • What is the effect size, i.e. how close is the association between change in bone turnover and change in fracture risk, and is it similar for different antiresorptive treatments? • Is this association valid for all patients, or is it restricted to certain subgroups (i.e. patients with low BMD [118], or large changes in bone turnover)? • Is the association between the change in bone turnover and the change in fracture risk a continous relationship (i.e. is more always better), or are there threshold/ceiling effects? Are there differences between agents? • Finally, what is a valid response? This question is of utmost importance when it comes to clinical decision making and the answer effectively determines the usefulness of bone markers for monitoring antiresorptive therapy. Not surprisingly, there are several answers to this question, and all are more or less controversial. As a consequence, the use of bone turnover markers in clinical practice is also controversial, to say the least. Again, variability is a major issue. In the presence of pronounced within-subject variability and regression to the mean (RTM) effects, it can be difficult to distinguish between an individual’s therapy-induced (“true”) and other biologic/stochastic changes. As a rule of thumb, markers showing large changes in response to interventions also show substantial degrees of variability. This is partly due to the ubiquitous RTM effect, which tends to yield unexpectedly pronounced “effects” even though

664 part of the result may have nothing to do with the intervention considered. Further above, we described two mathematical approaches to the assessment of an individual’s true change: the concept of “least significant change” (LSC), and the Bayesian model. The LSC approach is based upon a null hypothesis (e.g. “The observed change is within the range of within-subject fluctuation and therefore not significant”). It tends to underestimate biologic effects and, consequently, overemphasizes stochastic effects. Because the within-subject random variability may be constant over time for one individual, but likely varies between individuals, it could be argued that the application of one single criterion to all individuals may be inappropriate. In contrast, the Bayesian model works on a ‘forward-looking’ hypothesis such as “Given an observed change, what is the probability that a 5% change has taken place?” and provides a posterior distribution of the individual’s “true” value. The two methodologies are described in more detail in Chapter 34. In clinical medicine, two further interrelated approaches are often used to assess the clinical significance of a change. These include a comparison of the actual difference to a predefined cut-off, and a comparison of the measured value to a predefined range. The first method defines certain cut-off levels that a change in a marker must exceed to be considered “clinically significant”. Some models apply the LSC or similar statistical approaches to define a-priori cut-offs (44). Other models have used the placebo and treatment groups of large RCTs and calculated the fracture incidence according to the change in a marker above or below a certain cut-off. For example, using data from the fracture intervention trial, Bauer et al. demonstrated that alendronatetreated women with a reduction in serum BALP of 30% or more at 1 year had fewer hip and non-spine (but not vertebral) fractures then alendronate-treated women with a 1-year change in serum BALP of less than 30%. However, when compared to the placebo group, there was no difference in the incidence of vertebral fractures between the alendronate-treated groups (>30% vs. <30% change at 1 year). In this instance, therefore, setting a 30% cut-off helped to define – a posteriori – a valid response to alendronate treatment for non-spine and hip fractures, but appears to be useless for vertebral fractures (Fig. 9B) [119]. Another widely used method to assess therapy-induced changes in laboratory parameters is to compare the actual marker level to a predefined range (similar to the T and Z score approach in bone mineral density). In most instances, this range will be the “reference” or “normal” range, which may or may not be standardized between methods and/or laboratories. In the bone field, most people

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would agree that patients with accelerated bone turnover are likely to benefit from antiresorptive treatment if their bone markers return to the respective “normal”, i.e. premenopausal range. Such a “normalization” would be considered a valid response, while changes that do not lead to a return of the marker into the reference range would be labeled as an invalid response (Fig. 11). Furthermore, a reduction of the marker below the reference range could be indicative of over-treatment. A recent paper by Sambrook et al. illustrates the usefulness of this approach. In a double-masked and doubleplacebo-controlled study, the authors compared the effects of alendronate 70 mg once weekly with raloxifene 60 mg daily on markers of bone turnover in 487 postmenopausal women with low bone mineral density [76]. As shown in Figure 2, both antiresorptive agents reduced serum osteocalcin and urinary NTX-I levels after 1 year of treatment, but the effect was more pronounced for alendronate. Large RCTs have shown that both alendronate and raloxifene reduce fracture risk in women with low bone mineral density. It is therefore interesting to note that in the head-to-head trial by Sambrook et al. [76], both alendronate and raloxifene returned bone markers into their respective reference ranges. The problem with this method is that the reference ranges for most markers have not been well defined. Also, a reduction into the “normal” range can only be achieved if the pretreatment values are abnormally high, which is the case in less then 50% of patients with osteoporosis. Clearly, the approach is also invalid for anabolic treatments, which increase both bone formation and resorption (see Chapter 39). Finally, standardization of most assays is presently insufficient to provide a basis for a wider application of such a reference-based approach [136].

Figure 11 Defining a valid response. Comparison with a predefined reference range (gray area). While the solid line would be considered a valid response (“normalisation”), the dashed line would be typical for a non-response. The dotted line could be indicative of overtreatment.

Chapter 39 Monitoring of Antiresorptive Therapy

References 1. Fleming, T. R., and DeMets, D. L. (1996). Surrogate end points in clinical trials: are we being misled? Ann. Intern. Med. 125, 605–613. 2. Civitelli, R., Gonnelli, S., Zacchei, F., Bigazzi, S., Vattimo, A., Avioli, L. V., and Gennari, C. (1988). Bone turnover in postmenopausal osteoporosis. Effect of calcitonin treatment. J. Clin. Invest. 82, 1268–1274. 3. Overgaard, K., Hansen, M. A., Nielsen, V. A., Riis, B. J., and Christiansen, C. (1990). Discontinuous calcitonin treatment of established osteoporosis—effects of withdrawal of treatment. Am. J. Med. 89, 1–6. 4. Nielsen, N. M., von der Recke, P., Hansen, M. A., Overgaard, K., and Christiansen, C. (1994). Estimation of the effect of salmon calcitonin in established osteoporosis by biochemical bone markers. Calcif. Tissue Int. 55, 8–11. 5. Greenspan, S. L., Parker, R. A., Ferguson, L., Rosen, H. N., Maitland-Ramsey, L., and Karpf, D. B. (1998). Early changes in biochemical markers of bone turnover predict the long-term response to alendronate therapy in representative elderly women: a randomized clinical trial. J. Bone Miner. Res. 13, 1431–1438. 6. Gonnelli, S., Cepollaro, C., Pondrelli, C., Martini, S., Montagnani, A., Monaco, R., and Gennari, C. (1999). Bone turnover and the response to alendronate treatment in postmenopausal osteoporosis. Calcif. Tissue Int. 65, 359–364. 7. Garnero, P., Sornay-Rendu, E., Chapuy, M. C., and Delmas, P. D. (1996). Increased bone turnover in late postmenopausal women is a major determinant of osteoporosis. J. Bone Miner. Res. 11, 337–349. 8. Rosen, C. J., Chesnut, C. H., 3rd, and Mallinak, N. J. (1997). The predictive value of biochemical markers of bone turnover for bone mineral density in early postmenopausal women treated with hormone replacement or calcium supplementation. J. Clin. Endocrinol. Metab. 82, 1904–1910. 9. Chesnut, C. H., 3rd, Bell, N. H., Clark, G. S., Drinkwater, B. L., English, S. C., Johnson, C. C., Jr., Notelovitz, M., Rosen, C., Cain, D. F., Flessland, K. A., and Mallinak, N. J. (1997). Hormone replacement therapy in postmenopausal women: urinary N-telopeptide of type I collagen monitors therapeutic effect and predicts response of bone mineral density. Am. J. Med. 102, 29–37. 10. Gonnelli, S., Cepollaro, C., Pondrelli, C., Martini, S., Monaco, R., and Gennari, C. (1997). The usefulness of bone turnover in predicting the response to transdermal estrogen therapy in postmenopausal osteoporosis. J. Bone Miner. Res. 12, 624–631. 11. Hesley, R. P., Shepard, K. A., Jenkins, D. K., and Riggs, B. L. (1998). Monitoring estrogen replacement therapy and identifying rapid bone losers with an immunoassay for deoxypyridinoline. Osteoporos. Int. 8, 159–164. 12. Stevenson, J. C., Hillard, T. C., Lees, B., Whitcroft, S. I., Ellerington, M. C., and Whitehead, M. I. (1993). Postmenopausal bone loss: does HRT always work? Int. J. Fertil. Menopausal. Stud. 38 Suppl 2, 88–91. 13. Cummings, S. R., Karpf, D. B., Harris, F., Genant, H. K., Ensrud, K., LaCroix, A. Z., and Black, D. M. (2002). Improvement in spine bone density and reduction in risk of vertebral fractures during treatment with antiresorptive drugs. Am. J. Med. 112, 281–289. 14. Seibel, M. J., Naganathan, V., Barton, I., and Grauer, A. (2004). Relationship between pretreatment bone resorption and vertebral fracture incidence in postmenopausal osteoporotic women treated with risedronate. J. Bone Miner. Res. 19, 323–329. 15. Epstein, S., Poser, J., McClintock, R., Johnston, C. C., Jr., Bryce, G., and Hui, S. (1984). Differences in serum bone GLA protein with age and sex. Lancet. 1, 307–310. 16. Beardsworth, L. J., Eyre, D. R., and Dickson, I. R. (1990). Changes with age in the urinary excretion of lysyl- and hydroxylysylpyridinoline, two new markers of bone collagen turnover. J. Bone Miner. Res. 5, 671–676.

665 17. Garnero, P., and Delmas, P. D. (1993). Assessment of the serum levels of bone alkaline phosphatase with a new immunoradiometric assay in patients with metabolic bone disease. J. Clin. Endocrinol. Metab. 77, 1046–1053. 18. Seibel, M. J., Woitge, H., Scheidt-Nave, C., Leidig-Bruckner, G., Duncan, A., Nicol, P., Ziegler, R., and Robins, S. P. (1994). Urinary hydroxypyridinium crosslinks of collagen in population-based screening for overt vertebral osteoporosis: results of a pilot study. J. Bone Miner. Res. 9, 1433–1440. 19. Hassager, C., Risteli, J., Risteli, L., and Christiansen, C. (1994). Effect of the menopause and hormone replacement therapy on the carboxyterminal pyridinoline cross-linked telopeptide of type I collagen. Osteoporos. Int. 4, 349–352. 20. Johansen, J. S., Riis, B. J., Delmas, P. D., and Christiansen, C. (1988). Plasma BGP: an indicator of spontaneous bone loss and of the effect of oestrogen treatment in postmenopausal women. Eur. J. Clin. Invest. 18, 191–195. 21. Stepan, J. J., Pospichal, J., Schreiber, V., Kanka, J., Mensik, J., Presl, J., and Pacovsky, V. (1989). The application of plasma tartrate-resistant acid phosphatase to assess changes in bone resorption in response to artificial menopause and its treatment with estrogen or norethisterone. Calcif. Tissue Int. 45, 273–280. 22. Uebelhart, D., Schlemmer, A., Johansen, J. S., Gineyts, E., Christiansen, C., and Delmas, P. D. (1991). Effect of menopause and hormone replacement therapy on the urinary excretion of pyridinium cross-links. J. Clin. Endocrinol. Metab. 72, 367–373. 23. Hasling, C., Eriksen, E. F., Melkko, J., Risteli, L., Charles, P., Mosekilde, L., and Risteli, J. (1991). Effects of a combined estrogengestagen regimen on serum levels of the carboxy-terminal propeptide of human type I procollagen in osteoporosis. J. Bone Miner. Res. 6, 1295–1300. 24. Reginster, J. Y., Christiansen, C., Dequinze, B., Deroisy, R., Gaspard, U., Taquet, A. N., and Franchimont, P. (1993). Effect of transdermal 17 betaestradiol and oral conjugated equine estrogens on biochemical parameters of bone resorption in natural menopause. Calcif. Tissue Int. 53, 13–16. 25. Prestwood, K. M., Pilbeam, C. C., Burleson, J. A., Woodiel, F. N., Delmas, P. D., Deftos, L. J., and Raisz, L. G. (1994). The short-term effects of conjugated estrogen on bone turnover in older women. J. Clin. Endocrinol. Metab. 79, 366–371. 26. Cicinelli, E., Galantino, P., Pepe, V., Popolizio, A., Savino, F., Balzano, G., Epifani, S., and Cantatore, F. P. (1994). Bone metabolism changes after transdermal estradiol dose reduction during estrogen replacement therapy: a 1-year prospective study. Maturitas. 19, 133–139. 27. Riis, B. J., Overgaard, K., and Christiansen, C. (1995). Biochemical markers of bone turnover to monitor the bone response to postmenopausal hormone replacement therapy. Osteoporos. Int. 5, 276–280. 28. Bonde, M., Qvist, P., Fledelius, C., Riis, B. J., and Christiansen, C. (1995). Applications of an enzyme immunoassay for a new marker of bone resorption (CrossLaps): follow-up on hormone replacement therapy and osteoporosis risk assessment. J. Clin. Endocrinol. Metab. 80, 864–868. 29. Raisz, L. G., Wiita, B., Artis, A., Bowen, A., Schwartz, S., Trahiotis, M., Shoukri, K., and Smith, J. (1996). Comparison of the effects of estrogen alone and estrogen plus androgen on biochemical markers of bone formation and resorption in postmenopausal women. J. Clin. Endocrinol. Metab. 81, 37–43. 30. Hannon, R., Blumsohn, A., Naylor, K., and Eastell, R. (1998). Response of biochemical markers of bone turnover to hormone replacement therapy: impact of biological variability. J. Bone Miner. Res. 13, 1124–1133. 31. Cooper, C., Stakkestad, J. A., Radowicki, S., Hardy, P., Pilate, C., Dain, M. P., and Delmas, P. D. (1999). Matrix delivery transdermal 17beta-estradiol for the prevention of bone loss in postmenopausal women. The International Study Group. Osteoporos. Int. 9, 358–366.

666 32. Prestwood, K. M., Thompson, D. L., Kenny, A. M., Seibel, M. J., Pilbeam, C. C., and Raisz, L. G. (1999). Low dose estrogen and calcium have an additive effect on bone resorption in older women. J. Clin. Endocrinol. Metab. 84, 179–183. 33. Recker, R. R., Davies, K. M., Dowd, R. M., and Heaney, R. P. (1999). The effect of low-dose continuous estrogen and progesterone therapy with calcium and vitamin D on bone in elderly women. A randomized, controlled trial. Ann. Intern. Med. 130, 897–904. 34. Marcus, R., Holloway, L., Wells, B., Greendale, G., James, M. K., Wasilauskas, C., and Kelaghan, J. (1999). The relationship of biochemical markers of bone turnover to bone density changes in postmenopausal women: results from the Postmenopausal Estrogen/Progestin Interventions (PEPI) trial. J. Bone Miner. Res. 14, 1583–1595. 35. Garnero, P., Tsouderos, Y., Marton, I., Pelissier, C., Varin, C., and Delmas, P. D. (1999). Effects of intranasal 17beta-estradiol on bone turnover and serum insulin-like growth factor I in postmenopausal women. J. Clin. Endocrinol. Metab. 84, 2390–2397. 36. Prestwood, K. M., Kenny, A. M., Unson, C., and Kulldorff, M. (2000). The effect of low dose micronized 17ss-estradiol on bone turnover, sex hormone levels, and side effects in older women: a randomized, double blind, placebo-controlled study. J. Clin. Endocrinol. Metab. 85, 4462–4469. 37. Delmas, P. D., Hardy, P., Garnero, P., and Dain, M. (2000). Monitoring individual response to hormone replacement therapy with bone markers. Bone 26, 553–560. 38. Lindsay, R., Gallagher, J. C., Kleerekoper, M., and Pickar, J. H. (2002). Effect of lower doses of conjugated equine estrogens with and without medroxyprogesterone acetate on bone in early postmenopausal women. JAMA 287, 2668–2676. 39. Prestwood, K. M., Kenny, A. M., Kleppinger, A., and Kulldorff, M. (2003). Ultralow-dose micronized 17beta-estradiol and bone density and bone metabolism in older women: a randomized controlled trial. JAMA 290, 1042–1048. 40. Valimaki, M. J., Laitinen, K. A., Tahtela, R. K., Hirvonen, E. J., and Risteli, J. P. (2003). The effects of transdermal estrogen therapy on bone mass and turnover in early postmenopausal smokers: a prospective, controlled study. Am. J. Obstet. Gynecol. 189, 1213–1220. 41. Nielsen, T. F., Ravn, P., Bagger, Y. Z., Warming, L., and Christiansen, C. (2004). Pulsed estrogen therapy in prevention of postmenopausal osteoporosis. A 2-year randomized, double blind, placebo-controlled study. Osteoporos. Int. 15, 168–174. 42. Gallagher, J. C., Rapuri, P. B., Haynatzki, G., and Detter, J. R. (2002). Effect of discontinuation of estrogen, calcitriol, and the combination of both on bone density and bone markers. J. Clin. Endocrinol. Metab. 87, 4914–4923. 43. Delmas, P. D., Pornel, B., Felsenberg, D., Garnero, P., Hardy, P., Pilate, C., and Dain, M. P. (1999). A dose-ranging trial of a matrix transdermal 17beta-estradiol for the prevention of bone loss in early postmenopausal women. International Study Group. Bone 24, 517–523. 44. Delmas, P. D. (2000). Markers of bone turnover for monitoring treatment of osteoporosis with antiresorptive drugs. Osteoporos. Int. 11 Suppl 6, S66–76. 45. Anderson, J. J., Anthony, M. S., Cline, J. M., Washburn, S. A., and Garner, S. C. (1999). Health potential of soy isoflavones for menopausal women. Public Health Nutr 2, 489–504. 46. Morabito, N., Crisafulli, A., Vergara, C., Gaudio, A., Lasco, A., Frisina, N., D’Anna, R., Corrado, F., Pizzoleo, M. A., Cincotta, M., Altavilla, D., Ientile, R., and Squadrito, F. (2002). Effects of genistein and hormone-replacement therapy on bone loss in early postmenopausal women: a randomized double-blind placebo-controlled study. J. Bone Miner. Res. 17, 1904–1912. 47. Delmas, P. D., Bjarnason, N. H., Mitlak, B. H., Ravoux, A. C., Shah, A. S., Huster, W. J., Draper, M., and Christiansen, C. (1997). Effects of raloxifene on bone mineral density, serum cholesterol concentrations, and uterine endometrium in postmenopausal women. N. Engl. J. Med. 337, 1641–1647.

CHRISTIAN MEIER ET AL. 48. Ettinger, B., Black, D. M., Mitlak, B. H., Knickerbocker, R. K., Nickelsen, T., Genant, H. K., Christiansen, C., Delmas, P. D., Zanchetta, J. R., Stakkestad, J., Gluer, C. C., Krueger, K., Cohen, F. J., Eckert, S., Ensrud, K. E., Avioli, L. V., Lips, P., and Cummings, S. R. (1999). Reduction of vertebral fracture risk in postmenopausal women with osteoporosis treated with raloxifene: results from a 3-year randomized clinical trial. Multiple Outcomes of Raloxifene Evaluation (MORE) Investigators. JAMA 282, 637–645. 49. Morii, H., Ohashi, Y., Taketani, Y., Fukunaga, M., Nakamura, T., Itabashi, A., Sarkar, S., and Harper, K. (2003). Effect of raloxifene on bone mineral density and biochemical markers of bone turnover in Japanese postmenopausal women with osteoporosis: results from a randomized placebo-controlled trial. Osteoporos. Int. 14, 793–800. 50. Reginster, J. Y., Sarkar, S., Zegels, B., Henrotin, Y., Bruyere, O., Agnusdei, D., and Collette, J. (2004). Reduction in PINP, a marker of bone metabolism, with raloxifene treatment and its relationship with vertebral fracture risk. Bone 34, 344–351. 51. Hansdottir, H., Franzson, L., Prestwood, K., and Sigurdsson, G. (2004). The effect of raloxifene on markers of bone turnover in older women living in long-term care facilities. J. Am. Geriatr. Soc. 52, 779–783. 52. Prestwood, K. M., Gunness, M., Muchmore, D. B., Lu, Y., Wong, M., and Raisz, L. G. (2000). A comparison of the effects of raloxifene and estrogen on bone in postmenopausal women. J. Clin. Endocrinol. Metab. 85, 2197–2202. 53. Reid, I. R., Eastell, R., Fogelman, I., Adachi, J. D., Rosen, A., Netelenbos, C., Watts, N. B., Seeman, E., Ciaccia, A. V., and Draper, M. W. (2004). A comparison of the effects of raloxifene and conjugated equine estrogen on bone and lipids in healthy postmenopausal women. Arch. Intern. Med. 164, 871–879. 54. Garnero, P., Shih, W. J., Gineyts, E., Karpf, D. B., and Delmas, P. D. (1994). Comparison of new biochemical markers of bone turnover in late postmenopausal osteoporotic women in response to alendronate treatment. J. Clin. Endocrinol. Metab. 79, 1693–1700. 55. Bone, H. G., Downs, R. W., Jr., Tucci, J. R., Harris, S. T., Weinstein, R. S., Licata, A. A., McClung, M. R., Kimmel, D. B., Gertz, B. J., Hale, E., and Polvino, W. J. (1997). Dose-response relationships for alendronate treatment in osteoporotic elderly women. Alendronate Elderly Osteoporosis Study Centers. J. Clin. Endocrinol. Metab. 82, 265–274. 56. Stepan, J. J. and Vokrouhlicka, J. (1999). Comparison of biochemical markers of bone remodelling in the assessment of the effects of alendronate on bone in postmenopausal osteoporosis. Clin. Chim. Acta. 288, 121–135. 57. Ravn, P., Hosking, D., Thompson, D., Cizza, G., Wasnich, R. D., McClung, M., Yates, A. J., Bjarnason, N. H., and Christiansen, C. (1999). Monitoring of alendronate treatment and prediction of effect on bone mass by biochemical markers in the early postmenopausal intervention cohort study. J. Clin. Endocrinol. Metab. 84, 2363–2368. 58. Braga de Castro Machado, A., Hannon, R., and Eastell, R. (1999). Monitoring alendronate therapy for osteoporosis. J. Bone Miner. Res. 14, 602–608. 59. Garnero, P., Darte, C., and Delmas, P. D. (1999). A model to monitor the efficacy of alendronate treatment in women with osteoporosis using a biochemical marker of bone turnover. Bone 24, 603–609. 60. Greenspan, S. L., Rosen, H. N., and Parker, R. A. (2000). Early changes in serum N-telopeptide and C-telopeptide cross-linked collagen type 1 predict long-term response to alendronate therapy in elderly women. J. Clin. Endocrinol. Metab. 85, 3537–3540. 61. Ravn, P., Thompson, D. E., Ross, P. D., and Christiansen, C. (2003). Biochemical markers for prediction of 4-year response in bone mass during bisphosphonate treatment for prevention of postmenopausal osteoporosis. Bone 33, 150–158. 62. Mortensen, L., Charles, P., Bekker, P. J., Digennaro, J., and Johnston, C. C., Jr. (1998). Risedronate increases bone mass in an early postmenopausal population: two years of treatment plus one year of follow-up. J. Clin. Endocrinol. Metab. 83, 396–402.

Chapter 39 Monitoring of Antiresorptive Therapy 63. Harris, S. T., Watts, N. B., Genant, H. K., McKeever, C. D., Hangartner, T., Keller, M., Chesnut, C. H., 3rd, Brown, J., Eriksen, E. F., Hoseyni, M. S., Axelrod, D. W., and Miller, P. D. (1999). Effects of risedronate treatment on vertebral and nonvertebral fractures in women with postmenopausal osteoporosis: a randomized controlled trial. Vertebral Efficacy With Risedronate Therapy (VERT) Study Group. JAMA 282, 1344–1352. 64. Raisz, L., Smith, J. A., Trahiotis, M., Fall, P., Shoukri, K., Digennaro, J., and Sacco-Gibson, N. (2000). Short-term risedronate treatment in postmenopausal women: effects on biochemical markers of bone turnover. Osteoporos. Int. 11, 615–620. 65. Brown, J. P., Kendler, D. L., McClung, M. R., Emkey, R. D., Adachi, J. D., Bolognese, M. A., Li, Z., Balske, A., and Lindsay, R. (2002). The efficacy and tolerability of risedronate once a week for the treatment of postmenopausal osteoporosis. Calcif. Tissue Int. 71, 103–111. 66. Eastell, R., Barton, I., Hannon, R. A., Chines, A., Garnero, P., and Delmas, P. D. (2003). Relationship of early changes in bone resorption to the reduction in fracture risk with risedronate. J. Bone Miner. Res. 18, 1051–1056. 67. Ravn, P., Clemmesen, B., Riis, B. J., and Christiansen, C. (1996). The effect on bone mass and bone markers of different doses of ibandronate: a new bisphosphonate for prevention and treatment of postmenopausal osteoporosis: a 1-year, randomized, double-blind, placebo-controlled dose-finding study. Bone 19, 527–533. 68. Thiebaud, D., Burckhardt, P., Kriegbaum, H., Huss, H., Mulder, H., Juttmann, J. R., and Schoter, K. H. (1997). Three monthly intravenous injections of ibandronate in the treatment of postmenopausal osteoporosis. Am. J. Med. 103, 298–307. 69. Ravn, P., Christensen, J. O., Baumann, M., and Clemmesen, B. (1998). Changes in biochemical markers and bone mass after withdrawal of ibandronate treatment: prediction of bone mass changes during treatment. Bone 22, 559–564. 70. Cooper, C., Emkey, R. D., McDonald, R. H., Hawker, G., Bianchi, G., Wilson, K., and Schimmer, R. C. (2003). Efficacy and safety of oral weekly ibandronate in the treatment of postmenopausal osteoporosis. J. Clin. Endocrinol. Metab. 88, 4609–4615. 71. Christiansen, C., Tanko, L. B., Warming, L., Moelgaard, A., Christgau, S., Qvist, P., Baumann, M., Wieczorek, L., and Hoyle, N. (2003). Dose dependent effects on bone resorption and formation of intermittently administered intravenous ibandronate. Osteoporos. Int. 14, 609–613. 72. Garnero, P., Gineyts, E., Arbault, P., Christiansen, C., and Delmas, P. D. (1995). Different effects of bisphosphonate and estrogen therapy on free and peptide-bound bone cross-links excretion. J. Bone Miner. Res. 10, 641–649. 73. Reid, I. R., Brown, J. P., Burckhardt, P., Horowitz, Z., Richardson, P., Trechsel, U., Widmer, A., Devogelaer, J. P., Kaufman, J. M., Jaeger, P., Body, J. J., Brandi, M. L., Broell, J., Di Micco, R., Genazzani, A. R., Felsenberg, D., Happ, J., Hooper, M. J., Ittner, J., Leb, G., Mallmin, H., Murray, T., Ortolani, S., Rubinacci, A., Saaf, M., Samsioe, G., Verbruggen, L., and Meunier, P. J. (2002). Intravenous zoledronic acid in postmenopausal women with low bone mineral density. N. Engl. J. Med. 346, 653–661. 74. Evio, S., Tiitinen, A., Laitinen, K., Ylikorkala, O., and Valimaki, M. J. (2004). Effects of alendronate and hormone replacement therapy, alone and in combination, on bone mass and markers of bone turnover in elderly women with osteoporosis. J. Clin. Endocrinol. Metab. 89, 626–631. 75. Johnell, O., Scheele, W. H., Lu, Y., Reginster, J. Y., Need, A. G., and Seeman, E. (2002). Additive effects of raloxifene and alendronate on bone density and biochemical markers of bone remodeling in postmenopausal women with osteoporosis. J. Clin. Endocrinol. Metab. 87, 985–992. 76. Sambrook, P. N., Geusens, P., Ribot, C., Solimano, J. A., FerrerBarriendos, J., Gaines, K., Verbruggen, N., and Melton, M. E. (2004). Alendronate produces greater effects than raloxifene on bone density and bone turnover in postmenopausal women with low bone density: results of EFFECT (Efficacy of FOSAMAX versus EVISTA Comparison Trial) International. J. Intern. Med. 255, 503–511.

667 77. Pedrazzoni, M., Alfano, F. S., Gatti, C., Fantuzzi, M., Girasole, G., Campanini, C., Basini, G., and Passeri, M. (1995). Acute effects of bisphosphonates on new and traditional markers of bone resorption. Calcif. Tissue Int. 57, 25–29. 78. Seibel, M. J., Woitge, H. W., Pecherstorfer, M., Karmatschek, M., Horn, E., Ludwig, H., Armbruster, F. P., and Ziegler, R. (1996). Serum immunoreactive bone sialoprotein as a new marker of bone turnover in metabolic and malignant bone disease. J. Clin. Endocrinol. Metab. 81, 3289–3294. 79. Apone, S., Lee, M. Y., and Eyre, D. R. (1997). Osteoclasts generate cross-linked collagen N-telopeptides (NTx) but not free pyridinolines when cultured on human bone. Bone 21, 129–136. 80. Randall, A. G., Kent, G. N., Garcia-Webb, P., Bhagat, C. I., Pearce, D. J., Gutteridge, D. H., Prince, R. L., Stewart, G., Stuckey, B., Will, R. K., Retallack, R. W., Price, R. I., and Ward, L. (1996). Comparison of biochemical markers of bone turnover in Paget disease treated with pamidronate and a proposed model for the relationships between measurements of the different forms of pyridinoline cross-links. J. Bone Miner. Res. 11, 1176–1184. 81. Meunier, P. J., Confavreux, E., Tupinon, I., Hardouin, C., Delmas, P. D., and Balena, R. (1997). Prevention of early postmenopausal bone loss with cyclical etidronate therapy (a double-blind, placebo-controlled study and 1-year follow-up). J. Clin. Endocrinol. Metab. 82, 2784–2791. 82. Rossini, M., Gatti, D., Zamberlan, N., Braga, V., Dorizzi, R., and Adami, S. (1994). Long-term effects of a treatment course with oral alendronate of postmenopausal osteoporosis. J. Bone Miner. Res. 9, 1833–1837. 83. Greenspan, S. L., Emkey, R. D., Bone, H. G., Weiss, S. R., Bell, N. H., Downs, R. W., McKeever, C., Miller, S. S., Davidson, M., Bolognese, M. A., Mulloy, A. L., Heyden, N., Wu, M., Kaur, A., and Lombardi, A. (2002). Significant differential effects of alendronate, estrogen, or combination therapy on the rate of bone loss after discontinuation of treatment of postmenopausal osteoporosis. A randomized, double-blind, placebo-controlled trial. Ann. Intern. Med. 137, 875–883. 84. Bone H. G., Hosking, D., Devogelaer, J. P., Tucci, J. R., Emkey, R. D., Tonino, R. P., Rodriguez-Portales, J. A., Downs, R. W., Gupta, J., Santora, A. C., Liberman, U. A. (2004) Ten years’ experience with alendronate for osteoporosis in postmenopausal women. N. Engl. J. Med. 18, 350;1189–1199. 85. Harris, S. T., Eriksen, E. F., Davidson, M., Ettinger, M. P., Moffett, A. H., Jr., Baylink, D. J., Crusan, C. E., and Chines, A. A. (2001). Effect of combined risedronate and hormone replacement therapies on bone mineral density in postmenopausal women. J. Clin. Endocrinol. Metab. 86, 1890–1897. 86. Overgaard, K., and Christiansen, C. (1996). A new biochemical marker of bone resorption for follow-up on treatment with nasal salmon calcitonin. Calcif. Tissue Int. 59, 12–16. 87. Kraenzlin, M. E., Seibel, M. J., Trechsel, U., Boerlin, V., Azria, M., Kraenzlin, C. A., and Haas, H. G. (1996). The effect of intranasal salmon calcitonin on postmenopausal bone turnover as assessed by biochemical markers: evidence of maximal effect after 8 weeks of continuous treatment. Calcif. Tissue Int. 58, 216–220. 88. Thamsborg, G., Jensen, J. E., Kollerup, G., Hauge, E. M., Melsen, F., and Sorensen, O. H. (1996). Effect of nasal salmon calcitonin on bone remodeling and bone mass in postmenopausal osteoporosis. Bone 18, 207–212. 89. Ongphiphadhanakul, B., Piaseu, N., Chailurkit, L., and Rajatanavin, R. (1998). Suppression of bone resorption in early postmenopausal women by intranasal salmon calcitonin in relation to dosage and basal bone turnover. Calcif. Tissue Int. 62, 379–382. 90. Downs, R. W., Jr., Bell, N. H., Ettinger, M. P., Walsh, B. W., Favus, M. J., Mako, B., Wang, L., Smith, M. E., Gormley, G. J., and Melton, M. E. (2000). Comparison of alendronate and intranasal calcitonin for treatment of osteoporosis in postmenopausal women. J. Clin. Endocrinol. Metab. 85, 1783–1788.

668 91. Trovas, G. P., Lyritis, G. P., Galanos, A., Raptou, P., and Constantelou, E. (2002). A randomized trial of nasal spray salmon calcitonin in men with idiopathic osteoporosis: effects on bone mineral density and bone markers. J. Bone Miner. Res. 17, 521–527. 92. Buclin, T., Cosma Rochat, M., Burckhardt, P., Azria, M., and Attinger, M. (2002). Bioavailability and biological efficacy of a new oral formulation of salmon calcitonin in healthy volunteers. J. Bone Miner. Res. 17, 1478–1485. 93. Adami, S., Passeri, M., Ortolani, S., Broggini, M., Carratelli, L., Caruso, I., Gandolini, G., Gnessi, L., Laurenzi, M., Lombardi, A., Norbiato, G., Pryor-Tillstson, S., Reda, C., Romanini, L., Subriji, D., Wei, L., and Yates, A. J. (1995). Effects of oral alendronate and intranasal salmon calcitonin on bone mass and biochemical markers of bone turnover in postmenopausal women with osteoporosis. Bone 17, 383–390. 94. Simonet, W. S., Lacey, D. L., Dunstan, C. R., Kelley, M., Chang, M. S., Luthy, R., Nguyen, H. Q., Wooden, S., Bennett, L., Boone, T., Shimamoto, G., DeRose, M., Elliott, R., Colombero, A., Tan, H. L., Trail, G., Sullivan, J., Davy, E., Bucay, N., Renshaw-Gegg, L., Hughes, T. M., Hill, D., Pattison, W., Campbell, P., Sander, S., Van, G., Tarpley, J., Derby, P., Lee, R., and Boyle, W. J. (1997). Osteoprotegerin: a novel secreted protein involved in the regulation of bone density. Cell 89, 309–319. 95. Tsuda, E., Goto, M., Mochizuki, S., Yano, K., Kobayashi, F., Morinaga, T., and Higashio, K. (1997). Isolation of a novel cytokine from human fibroblasts that specifically inhibits osteoclastogenesis. Biochem. Biophys. Res. Commun. 234, 137–142. 96. Bekker, P. J., Holloway, D., Nakanishi, A., Arrighi, M., Leese, P. T., and Dunstan, C. R. (2001). The effect of a single dose of osteoprotegerin in postmenopausal women. J. Bone Miner. Res. 16, 348–360. 97. Bekker, P. J., Holloway, D. L., Rasmussen, A. S., Murphy, R., Martin, S. W., Leese, P. T., Holmes, G. B., Dunstan, C. R., and DePaoli, A. M. (2004). A single-dose placebo-controlled study of AMG 162, a fully human monoclonal antibody to RANKL, in postmenopausal women. J. Bone Miner. Res. 19, 1059–1066. 98. Seibel, M. J. (2003). Biochemical markers of bone remodeling. Endocrinol. Metab. Clin. North Am. 32, 83–113, vi–vii. 99. Filipponi, P., Pedetti, M., Fedeli, L., Cini, L., Palumbo, R., Boldrini, S., Massoni, C., and Cristallini, S. (1995). Cyclical clodronate is effective in preventing postmenopausal bone loss: a comparative study with transcutaneous hormone replacement therapy. J. Bone Miner. Res. 10, 697–703. 100. Dresner-Pollak, R., Mayer, M., and Hochner-Celiniker, D. (2000). The decrease in serum bone-specific alkaline phosphatase predicts bone mineral density response to hormone replacement therapy in early postmenopausal women. Calcif. Tissue Int. 66, 104–107. 101. Bjarnason, N. H., and Christiansen, C. (2000). Early response in biochemical markers predicts long-term response in bone mass during hormone replacement therapy in early postmenopausal women. Bone 26, 561–569. 102. Komulainen, M., Kroger, H., Tuppurainen, M. T., Heikkinen, A. M., Honkanen, R., and Saarikoski, S. (2000). Identification of early postmenopausal women with no bone response to HRT: results of a five-year clinical trial. Osteoporos. Int. 11, 211–218. 103. Fall, P. M., Kennedy, D., Smith, J. A., Seibel, M. J., and Raisz, L. G. (2000). Comparison of serum and urine assays for biochemical markers of bone resorption in postmenopausal women with and without hormone replacement therapy and in men. Osteoporos. Int. 11, 481–485. 104. Ravn, P., Clemmesen, B., and Christiansen, C. (1999). Biochemical markers can predict the response in bone mass during alendronate treatment in early postmenopausal women. Alendronate Osteoporosis Prevention Study Group. Bone 24, 237–244. 105. Fink, E., Cormier, C., Steinmetz, P., Kindermans, C., Le Bouc, Y., and Souberbielle, J. C. (2000). Differences in the capacity of several

CHRISTIAN MEIER ET AL.

106.

107.

108.

109.

110.

111.

112.

113.

114.

115.

116.

117.

118.

biochemical bone markers to assess high bone turnover in early menopause and response to alendronate therapy. Osteoporos. Int. 11, 295–303. Bone, H. G., Greenspan, S. L., McKeever, C., Bell, N., Davidson, M., Downs, R. W., Emkey, R., Meunier, P. J., Miller, S. S., Mulloy, A. L., Recker, R. R., Weiss, S. R., Heyden, N., Musliner, T., Suryawanshi, S., Yates, A. J., and Lombardi, A. (2000). Alendronate and estrogen effects in postmenopausal women with low bone mineral density. Alendronate/Estrogen Study Group. J. Clin. Endocrinol. Metab. 85, 720–726. Watts, N. B., Jenkins, D. K., Visor, J. M., Casal, D. C., and Geusens, P. (2001). Comparison of bone and total alkaline phosphatase and bone mineral density in postmenopausal osteoporotic women treated with alendronate. Osteoporos. Int. 12, 279–288. Storm, T., Thamsborg, G., Steiniche, T., Genant, H. K., and Sorensen, O. H. (1990). Effect of intermittent cyclical etidronate therapy on bone mass and fracture rate in women with postmenopausal osteoporosis. N. Engl. J. Med. 322, 1265–1271. Overgaard, K., Hansen, M. A., Jensen, S. B., and Christiansen, C. (1992). Effect of salcatonin given intranasally on bone mass and fracture rates in established osteoporosis: a dose-response study. BMJ 305, 556–561. Lufkin, E. G., Wahner, H. W., O’Fallon, W. M., Hodgson, S. F., Kotowicz, M. A., Lane, A. W., Judd, H. L., Caplan, R. H., and Riggs, B. L. (1992). Treatment of postmenopausal osteoporosis with transdermal estrogen. Ann. Intern. Med. 117, 1–9. Liberman, U. A., Weiss, S. R., Broll, J., Minne, H. W., Quan, H., Bell, N. H., Rodriguez-Portales, J., Downs, R. W. Jr., Dequeker, J., and Favus, M. (1995). Effect of oral alendronate on bone mineral density and the incidence of fractures in postmenopausal osteoporosis. The Alendronate Phase III Osteoporosis Treatment Study Group. N. Engl. J. Med. 333, 1437–1443. Cummings, S. R., Black, D. M., Thompson, D. E., Applegate, W. B., Barrett-Connor, E., Musliner, T. A., Palermo, L., Prineas, R., Rubin, S. M., Scott, J. C., Vogt, T., Wallace, R., Yates, A. J., and LaCroix, A. Z. (1998). Effect of alendronate on risk of fracture in women with low bone density but without vertebral fractures: results from the Fracture Intervention Trial. JAMA 280, 2077–2082. Sarkar, S., Mitlak, B. H., Wong, M., Stock, J. L., Black, D. M., and Harper, K. D. (2002). Relationships between bone mineral density and incident vertebral fracture risk with raloxifene therapy. J. Bone Miner. Res. 17, 1–10. Li, Z., and Meredith, M. P. (2003). Exploring the relationship between surrogates and clinical outcomes: analysis of individual patient data vs. meta-regression on group-level summary statistics. J. Biopharm. Stat. 13, 777–792. Riggs, B. L., Melton, L. J., 3rd, and O’Fallon, W. M. (1996). Drug therapy for vertebral fractures in osteoporosis: evidence that decreases in bone turnover and increases in bone mass both determine antifracture efficacy. Bone 18, 197S–201S. Bjarnason, N. H., Sarkar, S., Duong, T., Mitlak, B., Delmas, P. D., and Christiansen, C. (2001). Six and twelve month changes in bone turnover are related to reduction in vertebral fracture risk during 3 years of raloxifene treatment in postmenopausal osteoporosis. Osteoporos. Int. 12, 922–930. Sarkar, S., Reginster, J. Y., Crans, G. G., Diez-Perez, A., Pinette, K. V., and Delmas, P. D. (2004). Relationship between changes in biochemical markers of bone turnover and BMD to predict vertebral fracture risk. J. Bone Miner. Res. 19, 394–401. Bauer, D. C., Black, D. M., Garnero, P., Hochberg, M., Ott, S., Orloff, J., Thompson, D. E., Ewing, S. K., and Delmas, P. D. (2004). Change in bone turnover and hip, non-spine, and vertebral fracture in alendronate-treated women: the fracture intervention trial. J. Bone Miner. Res. 19, 1250–1258.

Chapter 39 Monitoring of Antiresorptive Therapy 119. McCombs, J. S., Thiebaud, P., McLaughlin-Miley, C., and Shi, J. (2004). Compliance with drug therapies for the treatment and prevention of osteoporosis. Maturitas 48, 271–287. 120. Marwick, C. (1994). Hormone combination treats women’s bone loss. JAMA 272, 1487. 121. Cano, A. (1994). Compliance to hormone replacement therapy in menopausal women controlled in a third level academic centre. Maturitas 20, 91–99. 122. Faulkner, D. L., Young, C., Hutchins, D., and McCollam, J. S. (1998). Patient noncompliance with hormone replacement therapy: a nationwide estimate using a large prescription claims database. Menopause 5, 226–229. 123. Kotzan, J. A., Martin, B. C., and Wade, W. E. (1999). Persistence with estrogen therapy in a postmenopausal Medicaid population. Pharmacotherapy 19, 363–369. 124. Kayser, J., Ettinger, B., and Pressman, A. (2001). Postmenopausal hormonal support: discontinuation of raloxifene versus estrogen. Menopause 8, 328–332. 125. Bjorn, I., and Backsrom, T. (1999). Drug related negative side-effects is a common reason for poor compliance in hormone replacement therapy. Maturitas 32, 77–86. 126. Segal, E., Tamir, A., and Ish-Shalom, S. (2003). Compliance of osteoporotic patients with different treatment regimens. Isr. Med. Assoc. J. 5, 859–862. 127. Chapurlat, R. D., and Cummings, S. R. (2002). Does follow-up of osteoporotic women treated with antiresorptive therapies improve effectiveness? Osteoporos. Int. 13, 738–744.

669 128. Clowes, J. A., Peel, N. F., and Eastell, R. (2004). The impact of monitoring on adherence and persistence with antiresorptive treatment for postmenopausal osteoporosis: a randomized controlled trial. J. Clin. Endocrinol. Metab. 89, 1117–1123. 129. Delmas, P. D., Vrijens, B., Roux, C., Le-Moigne-Amrani, A., Eastell, R., Grauer, A., Watts, N. B., Pols, H. A., Ringe, J. D., and Cahall, D. (2003). A reinforcement message based on bone turnover marker response influences long-term persistence with risedronate in osteoporosis: The IMPACT Study. J. Bone Miner. Res. 18, S374. 130. Di Munno, O., Mazzantini, M., Braga, V., and Adami, S. (2002). The DOMINO Project: To improve the compliance of osteoporotic patients with periodic feedback of urinary CTX values. J. Bone Miner. Res. 17, S374. 131. Nguyen, T. V., and Eisman, J. A. (2000). Assessment of significant change in BMD: a new approach. J. Bone Miner. Res. 15, 369–372. 132. Congdon, P. (2002). Bayesian statistical modelling. Wiley, New York pp. 11–61. 133. Berry, D. A. (1996). Statistics: A perspective. Duxbury Press, Belmont pp. 336–368. 134. Spiegelhalter, D. J., Freedman, L. S., and Parmar, M. K. (1994). Bayesian approaches to randomised trials. J. Royal Statist. Soc. Series A 157, 357–416. 135. Howson, C., and Urbach, P. (1993). Scientific reasoning: The Bayesian Approach. La salle: Open Court. La salle. pp. 357–416. 136. Seibel, M. J., Lang, M., and Geilenkeuser, W. J. (2001). Interlaboratory variation of biochemical markers of bone turnover. Clin. Chem. 47, 1443–1450.

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Chapter 40

Age-related Osteoporosis and Skeletal Markers of Bone Turnover CLIFFORD J. ROSEN

Maine Center for Osteoporosis Research and Education, St. Joseph Hospital, 900 Broadway, Bldg #2, Bangor, Maine 04401 USA

IV. Summary References

I. Introduction II. Epidemiology and Pathogenesis of Age-Related Osteoporosis-Relationship to Bone Turnover III. Markers of Bone Turnover and Age-Related Osteoporosis – Clinical Implications

I. INTRODUCTION

or hormonal determinants, remains to be determined. Still, there is strong evidence that there are certain phenotypic characteristics of individuals who sustain age-related fractures. This chapter will focus on the metabolic and biochemical changes in the skeleton, which characterize age-related osteoporosis. By convention, the term osteoporosis will denote a bone mineral density lower than 2.5 standard deviations below young normal mean values, irrespective of fracture history [4]. Age-related osteoporotic fractures will refer to either hip or spine fractures, recognizing that other sites are also susceptible to fractures. No attempt will be made in this chapter to classify individuals based on a single biochemical measure (e.g. parathyroid hormone), since several factors, in concert, reduce skeletal strength. Still, there are final common pathways (e.g. increased bone resorption or decreased bone formation)

Advanced age represents a major risk factor for osteoporosis [1]. Although low bone density accounts for a significant proportion of the risk in older individuals, age is clearly an independent risk factor for fracture [1, 2]. Indeed, the overwhelming majority of osteoporotic fractures occur after age 65 [2, 3]. Therefore, as the general population ages, it is projected that there will be significantly more osteoporotic fractures [3]. The pathogenesis of age-related fractures is complex. A hip fracture in a 75-year-old woman can be related to profound changes in several organ systems. Whether these perturbations are part of the normal aging process, are genetically programmed, or are pathognomonic of a larger subset of individuals susceptible because of environmental Dynamics of Bone and Cartilage Metabolism

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which are activated by aging and which contribute to skeletal weakness. These pathways will be examined in this chapter.

II. EPIDEMIOLOGY AND PATHOGENESIS OF AGE-RELATED OSTEOPOROSIS-RELATIONSHIP TO BONE TURNOVER A. Scope and Severity of the Problem Based on the WHO definition of osteoporosis (i.e. a bone mineral density [BMD] more than 2.5 SD below mean for a young normal individual) 30% of all postmenopausal women likely have osteoporosis of the hip, spine, or distal forearm [5]. Age has a dramatic impact on the epidemiology of this disease. Although only about 4% of women in their sixth decade have low bone density, the figure for women 80 years or older is approximately 50% [6]. Agerelated osteoporosis is manifested as fractures of the hip, wrist, and spine in older individuals. Hip fractures are by far the most dangerous and expensive manifestation of osteoporosis. One-year mortality from this event can range from 10–30% depending on several factors including age, gender, nutritional status, body composition, and place of residence [7]. For survivors, 15–50% will require long-term institutional care after their injury [8]. Medical complications such as infections, venous thrombosis, atelectasis, pneumonia, and malnutrition all contribute to morbidity [8]. Even if individuals do return to their previous living situation, many will have impaired quality of life and functional disabilities, which persist beyond the first year.

B. Pathogenesis of Age-related Osteoporosis: Relationship to Bone Remodeling 1. Falls and Fractures

Older men and women fall frequently, and these events lead to osteoporotic fractures of the spine and hip. Forty percent of individuals over the age of 80 have sustained at least one fall within the previous year [9]. Several studies have confirmed that gait, immobility, leg and foot dysfunction, and medications are the greatest risk factors for falling [10]. All these are much more common in the elderly. But not all falls result in fracture, even in the very old. For example, among hip fracture patients, falling directly on the hip or falls straight down increase the risk of a hip fracture up to 30-fold [11]. This is in sharp contrast to the relatively modest increased risk for fracture for every one

standard deviation decline in BMD. In fact, analysis of case-control and cohort studies reveal that major risk factors for hip fracture such as body weight (or weight loss after age 50), lower extremity strength, and mobility, and previous osteoporotic fractures are completely independent of BMD [12]. In respect to those age-related changes in bone that allow for increased susceptibility to fracture following a fall, qualitative aspects of skeletal architecture are the most prominent. In vivo and in vitro studies of aging bone reveal dramatic changes in trabecular connectivity, spacing and thickness, not reflected by BMD measurements [13–15]. Hence, a previous fracture in an elderly women likely signals a major defect in bone “quality” that would raise her risk of subsequent fracture. 2. Protein-calorie Malnutrition and Osteoporotic Fractures

It is estimated that 20–30% of elderly individuals are malnourished [16]. This may be an underestimate since it is difficult to survey the most vulnerable populations. Irrespective of the magnitude of the problem, this is a particularly relevant problem in respect to age-related osteoporosis since many hip fracture patients are undernourished at the time of injury. Since all individuals become catabolic post-fracture, poor dietary intake prior to a fracture is an ominous sign. Serum albumin, an indirect indicator of protein-calorie stores, is also a very good predictor of survival following hip fracture [17]. Protein supplementation for hip fracture patients has been shown to improve outcome and reduce hospital stays [18]. Whether there is a causal association between protein calorie undernutrition and hip fractures is still speculative. But weight loss, especially after age 50, is a major risk factor for hip fracture in women [19]. In addition, low serum albumin has also been associated with a greater risk of fracture [17]. Since poor muscle mass and impaired balance are important determinants of a subsequent hip fracture, and are also features of protein-calorie malnutrition, this may be one mechanism whereby nutritional status has a major effect on the skeletal response to trauma. Another potential pathway whereby malnutrition may affect fracture risk is through the insulin-like growth factor (IGF) system. IGF-I is produced by osteoblasts and is a major anabolic factor for bone [20]. It enhances osteoblast proliferation and collagen matrix biosynthesis [20]. Serum and skeletal levels of IGF-I decline with advanced age and may contribute to the pathogenesis of age-related osteoporosis [21]. Serum IGF-I levels also drop dramatically during protein-calorie malnutrition [22]. With poor dietary intake, falling levels of serum IGF-I could contribute to an overall reduction in bone formation which, in turn, may lead to uncoupling of bone turnover [14]. Indeed, in two

Chapter 40 Age-related Osteoporosis and Skeletal Markers of Bone Turnover

large cohort studies, low serum IGF-I has been shown to be an independent risk factor for hip fracture [23, 24]. Although the precise relationship between this serum marker and fracture remains uncertain. It is clear that the major source of circulating IGF-I is the liver and numerous hormones, cytokines, and other factors regulate hepatic synthesis of IGF-I [20]. Hence, even though it is relatively easy to extrapolate changes in serum IGF-I to a state of undernutrition, and to subsequent fracture risk, the evidence that poor nutrition alters the skeletal IGF regulatory system is less clear. At least two intervention studies suggest a strong relationship between nutritional supplementation and IGF-I. In one study undernourished elders post hip fracture that were supplemented with a high protein diet showed less bone loss in the contralateral femur than those individuals on a regular diet [25]. Moreover serum IGF-I levels rose quicker in those individuals receiving protein supplementation [25]. Short courses of rhIGF-I to fasting young women and women with anorexia nervosa have led to a marked stimulation of markers of bone formation and bone resorption as well as improvement in BMD, particularly when combined with antiresorptive agents [26]. In addition to reduced bone formation, malnourished individuals also have high rates of bone resorption. The mechanisms responsible for such an effect are not known. But there is now strong evidence from several epidemiological studies in elderly women that increased bone resorption is an independent risk factor for fracture [27, 28]. However, as noted previously, individuals who fracture often live in chronic care facilities, are vitamin D deficient, are on multiple medications, and suffer from several disease states simultaneously. Therefore, from a broad perspective, true cause and effect between malnutrition and rapid bone resorption is difficult to ascertain. Certainly, secondary hyperparathyroidism, which results from calcium and vitamin D deficiency, can stimulate bone resorption and may be one pathway by which bone resorption is accentuated. 3. Calcium Intake and Age-related Osteoporotic Fractures

Older individuals consume little calcium. Most studies estimate calcium intake in healthy elders to be between 400–600 mg/day [29]. In part, this can be related to the reduction in milk intake, although use of multiple medications, changes in appetite, taste alterations, and other factors contribute to poor dietary calcium consumption. In addition to minimal calcium intake, the compensatory mechanisms, which are key to maintaining calcium homeostasis, change dramatically. The renal response to increased serum PTH is attenuated, the skin synthesizes less vitamin D, intestinal mucosal alterations reduce the capacity of the

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gastrointestinal tract to enhance calcium absorption, 1-alphahydroxylase activity declines and kidney function deteriorates [30]. Due to these age-related alterations, calcium balance studies have established that maintenance of calcium homeostasis in the elderly requires a dietary intake of approximately 1500 mg/day [31]. This is consistent with recent recommendations from the National Academy of Sciences, which support the use of 1500 mg of calcium per day in women over age 65 regardless of bone density or fracture risk. Bone turnover is accelerated in the elderly and is associated with bone loss. Although initially most investigators postulated that reduced bone formation led to uncoupling in the remodeling unit of very old individuals, it has now been established that bone resorption is markedly enhanced in both men and women beyond the eighth decade of life [28]. Markers of bone resorption, such as deoxypyridinoline, have been shown to predict rapid bone losers in older postmenopausal women and this may be an independent risk factor for fracture (see Section III B1) [32–34]. There is probably no single cause for accelerated bone resorption in the elderly, but secondary hyperparthyroidism, which results from reduced calcium and vitamin D intake, plays an important role [35]. In fact, there is a marked rise in serum PTH with age that correlates with declining renal function [36]. In addition, higher levels of PTH are associated with reduced serum 25OH vitamin D levels, particularly in the elderly [37–39]. Increased PTH concentrations are closely associated with several markers of bone turnover, although less so with changes in BMD [40, 41]. But, in one of the larger cohort studies of elders, the Baltimore Longitudinal Study of Aging, radial bone mineral density was inversely related to serum PTH [42]. Similarly, in another study, a strong inverse relationship was noted between PTH and femoral bone density [43]. However, Chapuy and coworkers found in elderly French women that only 4% of the variance in femoral BMD could be related to PTH [44]. Other cross-sectional studies have shown that hip fracture patients have higher levels of serum PTH than age-matched controls but, due to design issues, this could reflect many perturbations in other organ systems with aging [45]. On the other hand, at least one study in elderly women who sustained hip fractures in Boston, Massachusetts, revealed that a great proportion (i.e. > 50%) were either vitamin D deficient (i.e. serum 25OHD < 10 ng/ml), or insufficient (serum 25OHD < 30 ng/ml) [46]. A similar finding was noted in a large cohort of hip fracture patients in Italy [47]. During calcium deficiency states, bone formation is suppressed, almost certainly to protect the supply of calcium in the circulation. But it is unclear how this occurs within the skeleton. Older individuals are likely to be in

674 a chronic and persistent calcium deficiency state, which, in turn, can lead to secondary hyperparathyroidism. In one study it was noted that one of the IGF-binding proteins (IGFBPs) that is synthesized by osteoblasts and is inhibitory in bone, IGFBP-4, is increased in elderly individuals [48]. Furthermore, serum PTH levels correlate closely with IGFBP-4 [49]. Also, PTH can increase IGFBP-4 expression in bone cells. Moreover, in one small cross-sectional study, IGFBP-4-binding intensity was significantly higher in hip fracture patients than controls [50]. Since this binding protein is strongly inhibitory to IGF-I at high concentrations, it is conceivable that impaired bone formation during experimental calcium deficiency relates to increased IGFBP-4 production leading to reduced IGF-I bioactivity. However, the clinical use of serum IGFBP-4 as a marker of suppressed bone formation remains doubtful, particularly in light of a more recent study from the Framingham cohort demonstrating age-associated changes in IGF-I, IGFBP-4, and IGFBP-5, but little correlation with bone markers or BMD [51]. Notwithstanding that study, bone turnover is markedly altered in the elderly when dietary calcium is limited. Clearly, bone resorption is increased by calcium deficiency, but at the same time bone formation may also be suppressed. Hence, it is not surprising that bone loss could be dramatic in elders with calcium-deficient diets. The strongest evidence to support the thesis that chronic calcium deficiency in the elderly is a major risk factor for osteoporotic fractures is derived from case-control and cohort studies. Most of these studies have shown that women consuming the greatest amount of calcium have the lowest risk of hip fracture [52, 53]. Prospective calcium intervention trials in older postmenopausal women are also supportive of that premise, including a very recent study in Australia with more than 1500 elderly women [54]. In another study, women with calcium intakes below 450 mg/day and more than 5 years after menopause who were randomized to calcium supplementation exhibited virtually no bone loss from three skeletal sites [55]. Finally, another smaller study showed that calcium supplementation to osteoporotic women with calcium intakes less than 1 g/day resulted in a reduction in incident vertebral fractures [56]. Besides preserving bone mass and preventing spinal fractures, supplemental calcium, particularly in combination with vitamin D, in older postmenopausal women showed a reduction in the number of non-vertebral fractures [57]. This calcium intervention trial involved more than 3000 elderly French women (mean age 84) who were randomized to either 1200 mg of calcium phosphate plus 800 IU of vitamin D or placebo for 3 years. Calcium-supplemented women showed a reduction in all non-vertebral fractures

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and a 30% decrease in hip fractures after only 18 months of treatment [44]. In virtually all the calcium intervention trials in the elderly, calcium (with or without vitamin D) not only preserves BMD but also leads to a reduction in serum PTH [52, 53, 55–57]. These findings would suggest that calcium supplementation is effective because it can indirectly block bone resorption via suppression of PTH. However, it is not clear what threshold dose of calcium is required to suppress PTH. One recent study showed that 2400 mg/day of calcium reduced PTH values to young normal values [30]. Timing of calcium supplementation may also be critical since there is a strong diurnal variation in bone resorption that is more accentuated in the elderly [28, 58]. A complicating factor in this equation however, is that circulating 25OHD levels are also related to endogenous PTH secretion. Several groups have shown a strong inverse relationship between 25OHD and PTH, particularly when vitamin D levels are lower than 30 ng/ml [59, 60]. Further studies will be needed to determine whether calcium supplementation in the evening is more effective than calcium supplements consumed during meals. In summary, calcium deficiency promotes a state of secondary hyperparathyroidism that leads to an increase in calcium mobilization from bone stores but also results in suppression in bone formation [61]. Biochemical markers of bone resorption are increased while bone formation indices may be increased, unchanged, or reduced, depending on the surrogate marker. Serum IGF-I levels are low in age-related osteoporotic patients compared to controls, and low serum IGF-I is a risk factor for hip fractures in elderly women [24]. The combination of increased resorption and reduced bone formation likely accentuates the uncoupling of the bone remodeling unit, and is certainly considered an independent risk factor for subsequent osteoporotic fractures. 4. Vitamin D and Age-related Osteoporosis

Adequate vitamin D is essential for increasing the efficiency of calcium absorption in the gut [62] (see also Chapters 18 and 31). The presence of normal levels of vitamin D can enhance calcium absorption from 15–60% [62, 63]. As noted previously, most elderly people consume very little calcium because they do not drink milk. Since the major dietary source of vitamin D in the US is fortified milk, dietary vitamin D insufficiency is very common. It is estimated that upwards of 85% of elderly nursing home residents who do not receive a vitamin supplement have serum levels of 25 hydroxyvitamin D below 20 ng/ml [64]. Classically 12 ng/ml or lower of serum 25 hydroxyvitamin D is considered vitamin D deficiency [65].

Chapter 40 Age-related Osteoporosis and Skeletal Markers of Bone Turnover

Estimates for the number of chronic care residents who are vitamin D deficient ranges from 20% to 70% [65]. In part this can be explained by the fact that most circulating 25-hydroxyvitamin D is derived from percutaneous conversion of provitamin to previtamin D. Previtamin D is metabolically unstable and is converted within hours to vitamin D3 where it subsequently undergoes 25-hydroxylation in the liver and 1,25-hydroxylation in the kidney (see also Chapters 18 and 31). Lack of sunlight, skin coverings, or sunscreen, or living in a region where the apogee of the sun during winter is not high enough to permit the non-enzymatic conversion of provitamin D to previtamin D in the epidermis, all are factors which limit the number of elderly people who are vitamin D sufficient. Also, it should be noted that elderly individuals can only make approximately one third the vitamin D3 from skin that younger individuals can produce [66]. Thus it is the sum of age, poor dietary intake, and limited solar exposure that contribute to the high rate of vitamin D deficiency in the elderly. Lack of vitamin D reduces the efficiency of calcium absorption in the gut and leads to secondary hyperparathyroidism. A rise in PTH, which is triggered by a drop in serum 25OH vitamin D and low dietary calcium, can result in increased bone resorption and is the most likely explanation for bone loss in vitamin D-insufficient elders. In addition to chronically low dietary calcium intake, PTH is also stimulated by seasonal changes in serum 25OH vitamin D. In both cross-sectional and longitudinal studies of healthy elderly women living in northern latitudes, serum 25 hydroxyvitamin D can drop by as much as 25% during the winter months [41, 57, 60, 67]. This triggers PTH release, which in turn can lead to an increase in bone turnover as measured by serum osteocalcin and urinary N-telopeptide excretion [68]. Very recent studies from Australia revealed that elevated serum parathyroid hormone levels predict time to fall and mortality in frail elderly men and women, independent of their vitamin D status [69, 70]. The seasonal rise in PTH as a result of a decline in serum 25-hydroxyvitamin D is also responsible for a rise in serum IGFBP-4 observed during the winter in northern New England women [68]. In a recent 2-year prospective placebo-controlled trial with calcium carbonate, healthy elderly women given a placebo calcium pill demonstrated a 45% increase in urinary N-telopeptide, and 50% increase in osteocalcin during the winter months (Fig. 1A, B) [71]. These changes were directly related to a marked seasonal decline in 25OH vitamin D and a secondary increase in PTH secretion (Fig. 1C). Surprisingly, elderly women in that study lost nearly 3% of their femoral BMD over 6 winter

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Figure 1 (A) Serial changes in serum 25OHD over a 24-month period in elderly women residing in northern New England. Three groups of women were followed [60]: 20 women receiving placebo calcium, 20 women consuming four glasses of milk/day, and 20 women getting 1000 mg/day of calcium supplements. Baseline calcium intake was approximately 600 mg/day for each group. Mean summer serum 25OHD levels were statistically different than winter levels for each group at P < 0.01. (B) In the same three groups of women, the winter and summer urine NTX values were averaged for the 2 years of the study; bone turnover was greatest in the winter for the placebo group (mean 45% increase over baseline) consuming the lowest amount of calcium. During the summer months, bone turnover was negligible. A similar effect was noted for greater trochanteric BMD; winter loss for placebo women was –2.1% vs. summer loss which was negligible (–0.2%). (C) Serial changes in osteocalcin from baseline in three groups of healthy elderly women. Women consuming the least amount of calcium had the greatest increase in osteocalcin, all of which occurred during winter months.

months [68]. Based on these data, it is likely that this high rate of bone loss over a relatively short period of time in otherwise healthy elders has to be related to uncoupling of the bone remodeling cycle as a result of increased bone resorption and decreased bone formation.

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Although vitamin D deficiency results in bone loss, it also has other effects, which may contribute to increased bone fragility. Low vitamin D reduces mineralization of new bone and in some cases can lead to osteomalacia [72]. This may contribute to reduced bone strength. The percent of elderly women who sustain hip fractures and have osteomalacia is likely to be about 10%, although this is only an estimate since few patients have bone biopsies to verify overt osteomalacia. In several studies upwards of 50% of osteoporotic women with hip fractures were vitamin D deficient (<12 ng/ml) or insufficient [46, 73]. This proportion is highly dependent on the geographic site of the study, the origin of the patient (nursing home, hospital or community), and the general nutritional status of the individual. In addition to reduced bone mineralization, osteomalacia is associated with proximal muscle weakness, a risk factor for falls and subsequent fracture. A recent large, randomized, placebo-controlled study confirmed the importance of vitamin D in preventing fractures [74]. In that trial, high-dose vitamin D prevented non-vertebral fractures even when administered intermittently. Thus, it seems likely that vitamin D may play a critical role in several pathophysiologic pathways leading to enhanced skeletal fragility. 5. Vitamin K Deficiency and Age-related Osteoporosis

Vitamin K is a co-factor for enzymes involved in the post-translational modification of osteocalcin and other bone matrix proteins, which bind to hydroxyapatite. In some elderly patients with osteoporosis, undercarboxylated osteocalcin can be detected in the circulation, and these levels are suppressed by supplementary vitamin K [75]. Also, decreased serum levels of vitamin K (phyloquinone and the menaquinones) have been found in elders who have sustained a hip fracture [76–78]. In a recent prospective study of elderly institutionalized women, those women with high levels of undercarboxylated osteocalcin at baseline had a greater likelihood of hip fracture compared to any other biochemical marker or carboxylated osteocalcin [79]. However, the level of undercarboxylated osteocalcin also correlated with vitamin D status, suggesting that this may be a non-specific marker of undernutrition. It is not clear precisely how skeletal or serum vitamin K affect bone remodeling. In one study urinary calcium loss in some patients with osteoporosis was reduced [80]. In another study, vitamin K supplementation reduced bone resorption [71]. On the other hand, it is possible that vitamin K may modulate bone formation via matrix biosynthesis. The only prospective fracture study with vitamin K supplementation involved a small cohort of renal failure patients. That study demonstrated a relationship between vitamin K1 levels and fracture risk [81].

Longitudinal studies are currently underway to further assess the importance of vitamin K in age-related osteoporosis. In summary, age-related osteoporotic fractures are closely linked to overall nutrition as well as calcium/ vitamin D and vitamin K sufficiency. Lack of these dietary factors is associated with increased bone resorption and decreased bone formation. The end result of changes in nutrient intake is uncoupling of the bone remodeling process leading to rapid bone loss and increased skeletal fragility. Seasonal changes in 25OH vitamin D and PTH accelerate bone resorption and may suppress bone formation. From these perturbations, it is easy to see how accelerated bone loss in the elderly may lead to altered skeletal architecture and subsequent fractures. 6. Hormonal Deficiency States and Bone Turnover in the Elderly

a. Overview of Aging and Hormonal Factors Aging is accompanied by major changes in every hormonal system. Decreased synthesis, altered feedback inhibition, enhanced catabolism, reduced receptor half-life, and post receptor perturbations can lead to hormonal deficiency or resistance [82]. Such hormonal changes in older individuals are certain to affect bone remodeling. For example, diminution in gonadal steroid production in both males and females can lead to higher rates of bone resorption. Similarly, impaired growth hormone release can lower serum and skeletal IGF-I, thereby affecting bone formation. Age-related increases in PTH have already been examined in relation to calcium and vitamin D availability (see above). Weak adrenal androgens such as DHEA and DHEA-S decline with age, are associated with low serum IGF-I, and may affect both bone resorption and formation [83]. The major hormonal activators (or suppressors) of bone remodeling are discussed below. b. Testosterone Deficiency and Turnover In men there is a modest decline in total and free testosterone with age [84]. Although this drop is similar to the age-associated reduction in BMD noted among males, cause and effect have never been firmly established. Several cross-sectional studies have indirectly linked testosterone to BMD [14, 85, 86]. Certainly, males with frank hypogonadism have low bone mineral density and high rates of bone resorption. In response to parenteral testosterone replacement these men have an increase in BMD and decreased bone resorption [87, 88]. However, in the aging male without hypogonadism, the evidence that testosterone, or its metabolite, dihydrotestosterone (DHT), is critical for the maintenance of bone mass is controversial. Indeed, there is now emerging evidence that estrogen, rather than

Chapter 40 Age-related Osteoporosis and Skeletal Markers of Bone Turnover

androgen, is critical in preventing age-related bone loss due to increased bone resorption [89]. Theoretically, androgens can affect bone turnover in several ways. First, it has been shown that testosterone supplementation blocks cytokine release from osteoblasts, whereas androgen deficiency leads to up-regulation of several cytokines including IL-6 [90]. These perturbations are associated with a profound alteration in the rate of bone resorption. Second, there are androgen receptors on osteoblasts, and there is some evidence that androgens can increase osteoblastic activity, independent of their effects on resorption [91]. However, in vivo studies with androgen replacement suggest that inhibition of bone resorption is the predominant mechanism of action [92]. Third, the absence of androgens also reduces the circulating concentration of estradiol, a critical hormone for preserving BMD in men [93, 94]. Despite these lines of experimental evidence, data from the relatively few prospective studies in men reveal only a very weak relationship between serum testosterone (or free testosterone) and bone mineral density or future fracture [95, 96]. This is also true for postmenopausal women who produce some androgens, including testosterone. Several prospective studies have failed to demonstrate a significant relationship between testosterone and bone density [97]. Yet there is preliminary evidence that levels of sex hormone-binding globulin may be indirectly related to risk of a hip fracture [98]. c. Estrogen Deficiency and the Aging Skeleton For more than half a century the role of estrogen in controlling bone remodeling has been appreciated [99]. However, the effects of estrogens on the aging skeleton has only recently been examined. In previous decades, the conventional approach to the elderly individual with osteoporosis consisted exclusively of non-hormonal therapy. However, trials with HRT in older postmenopausal osteoporotic women, including the WHI, have established that estrogen and progesterone can prevent fractures, and that the bone mineral density response to HRT in elderly women is vigorous, exceeding bone mineral density increments in younger women [100–102]. This had led many investigators to postulate that estrogen deficiency contributes to the pathogenesis of age-related osteoporosis. In part, an exaggerated skeletal response to estrogen can be traced to baseline indices of bone turnover. In two longitudinal trials, one with transdermal and one with oral estrogens, the women undergoing treatment were frankly osteoporotic [100, 101]. Although not measured in the original studies, subsequent evaluations of older osteoporotic women have demonstrated a very high rate of bone turnover accompanied by accelerated bone loss. These studies employed several markers of resorption to demonstrate an age-associated rise in bone turnover [28, 103].

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Indeed, high turnover is the most likely explanation for the marked increase in bone density following antiresorptive therapy with estrogens. However, this does not mean that chronic estrogen deprivation causes age-related osteoporotic fractures. As noted previously, this disease is multifactorial and declining estrogen levels may be but one of several permissive factors that increase bone turnover. This hypothesis is reinforced by findings from several prospective studies using newer more sensitive assays for estradiol, in which endogenous estradiol in postmenopausal women is directly correlated with femoral bone mineral density [104]. In sum, older postmenopausal women who are osteoporotic are likely to have relatively high rates of bone turnover and are more responsive to antiresorptive therapy such as estrogen replacement. As noted, estrogen concentrations may play a crucial role in the pathogenesis of age-related bone loss in men. The report of a young male with osteoporosis lacking the alpha estrogen receptor provided the impetus to further examine the relationship between endogenous estrogen production and bone mass in men [105]. The phenotype of that seminal case was a tall man with severe osteopenia who had not fused his epiphyses and had high normal testosterone levels. Biochemical markers of bone turnover suggested a markedly increased rate of bone resorption although histomorphometric analysis was not conclusive [105]. Two recent cases of selective aromatase deficiency in men with a similar phenotype as the estrogen receptordeficient male further supports the hypothesis that absence of endogenous estrogen, or its receptor, is associated with low bone density and high bone turnover [106, 107]. In one of those cases, estrogen replacement led to a 25% increase in bone density after 1 year of treatment [108]. That response would be consistent with the antiresorptive properties of estrogen in patients with high turnover. These lines of evidence have pushed investigators to examine the role of endogenous estrogen in healthy men with and without low bone mass. One cross-sectional study of men has confirmed a very strong correlation between serum estradiol and BMD and more recently with bone turnover [89, 109]. In sum, estrogen production may play an important role in restraining bone resorption in males. Heritable or acquired disorders of estrogen synthesis or defects in the estrogen receptor are associated with very low bone mass and heightened bone turnover in men. Whether the gonadal steroids serve a dual function to maintain bone integrity (i.e. restrict bone resorption, but stimulate bone formation) will still need to be proved. However, this concept is relevant for age-related osteoporosis since estrogen levels in men decline with aging. Also, the advent of second- and third-generation selective estrogen receptor modulators

678 raises the possibility of treating high turnover bone loss in the elderly male with an agent such as raloxifene without causing other undue side effects. d. The Growth Hormone/IGF-I Axis and Age-related Osteoporosis Advanced age is associated with a decline in the magnitude and frequency of growth hormone surges from the pituitary [110]. This, in part, may be related to waning production of endogenous GH secretagogs, or enhanced synthesis of somatostatin. Irrespective of the etiology, less growth hormone secretion means lower concentrations of serum and tissue IGF-I [111]. Although attempts have been made to link age-associated declines in IGF-I with bone loss due to aging, the cause and effect have never been established. Several lines of evidence, however, suggest that relative growth hormone deficiency could impact bone turnover, thereby affecting the overall rate of bone loss. First, it is now recognized that adult growth hormone deficiency (GHD) is associated with low bone mass, and replacement with rhGH can increase bone density by upwards of 5% after 3 years [112]. Second, several cross-sectional studies have identified a correlation between serum IGF-I and bone density [113, 114]. In two large cohort studies there is further evidence of an association between IGF-I and age-related osteoporosis. In the Study of Osteoporotic Fractures with over 9000 postmenopausal women, the lowest quartile of IGF-I was associated with the greatest risk of hip fracture [23]. Similarly, in a large French cohort of elderly postmenopausal women, the lowest tertile of serum IGF-I was associated with the greatest risk of hip fracture [24]. Furthermore, in a subset of men and women from the Framingham cohort, there was a strong association between quartiles of serum IGF-I and BMD at the spine, hip, and wrist [115]. Third, other components of the IGF regulatory system that are synthesized in the skeleton have been linked to age-related changes in growth hormone secretion. As noted previously, serum levels of IGFBP-4 are increased in older individuals and may be even higher among hip fracture patients [50]. Since this is an inhibitory binding protein, which blocks IGF-I action, and there is an age-related decline in bone formation, this protein may have pathophysiologic significance. Two IGF binding proteins, which are agonists for IGF-I and are synthesized by osteoblasts, IGFBP-3, and IGFBP-5, are also affected by the aging process. IGFBP-3 is the major IGF carrier protein in the circulation and is modulated principally by growth hormone. Elderly individuals show a marked decrease in serum IGFBP-3 [112]. Since several cross-sectional studies have shown a relationship between serum IGFBP-3 and bone mineral density, changes in this binding protein, either locally or systemically, may be important [116, 117].

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Similarly, IGFBP-5, an osteoblast-derived IGFBP which binds IGF-I to hydroxyapatite, also shows a decline with age in both the serum and the skeleton [49]. Like IGFBP-3, IGFBP-5 enhances IGF bioactivity, is related to circulating GH, and increases during peak bone mass acquisition. Hence a combination of changes in several IGFBPs, as a result of a relative GH deficiency, could alter bone remodeling. Finally, studies in patients who survive a hip fracture reveal a marked decline in lean body mass, bone mineral content, and serum IGF-I in the immediate postoperative period [118]. These changes reflect not only the catabolic state, which follows injury, but also the predisposing characteristics of the individuals who fracture. And, as noted previously, protein supplementation to hip fracture patients in a randomized controlled trial demonstrated improved outcomes and higher IGF-I serum levels [25]. More recently, the use of IGF-I/IGFBP-3 to a small group of hip fracture women also revealed an improvement in BMD and greater grip strength [119]. One small study in elderly frail women used a GH-releasing analog in combination with alendronate and demonstrated a greater increase in femoral BMD than alendronate alone [120]. However, it is unclear whether these differences translate into changes in overall life quality, particularly after a hip fracture [121]. In sum, declining skeletal levels of IGF-I, IGFBP-3, and IGFBP-5 due to growth hormone deficiency, and an increase in IGFBP-4 secondary to high PTH levels, may contribute to the process of age-related bone loss and the subsequent fractures which accompany low bone mass. However, as noted previously, most if not all the components of the IGF regulatory system are under multiple layers of control, thereby making it difficult to assess how changes in a single regulatory component (e.g. growth hormone) perturb the system enough to cause a fracture. For example, nutritional deficiencies can affect not only IGF-I but also synthesis of IGFBP-1, -3, -4, and -5 as well as IGF receptor expression [22]. Thus the bioactivity of IGF-I could be affected at several different levels from the ligand all the way to the receptor. Clearly, changes in the hypothalamic-pituitary axis, which modulate GH secretion, could ultimately affect several IGF-rich tissues such as bone. These hormonal alterations may ultimately turn out to be major etiologic factors in the pathogenesis of age-related osteoporosis. 7. Genetic Determinants of Age-related Osteoporosis Maternal history of a hip fracture is a major and independent risk factor for a future fracture [122]. This would suggest that there might be genetic determinants of fracture that are independent of bone mineral density “genes”. Indeed, previous fracture and a maternal history of hip fracture are powerful predictors of subsequent fracture for

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Chapter 40 Age-related Osteoporosis and Skeletal Markers of Bone Turnover

elderly individuals [122]. However, assessing the genetic determinants of the hip fracture phenotype is daunting for several reasons. First, the onset of hip fractures is almost always after the seventh decade. This makes phenotyping extremely difficult. It also makes family and twin studies almost impossible to conduct. Second, hip fractures are multifactorial in origin. As noted above, several different structural, environmental, and hormonal factors contribute to fracture risk. These may include hip axis length, dietary intake of calcium, length of estrogen exposure, relative growth hormone production, serum IGF-I, and a host of other factors. Third, it is unclear that hip fracture represents a continuous variable like bone mineral density. Progress in defining the genetic aspects of osteoporosis has been successful to date, in part because the phenotype (i.e. bone mineral density) is so easy to characterize. These measurements have opened the door for more sophisticated twin, cohort, and population studies. Notwithstanding the difficulties inherent in studying the genetic regulation of bone loss in the elderly, most investigators believe there are genetic programs which regulate the onset and severity of age-related osteoporosis. For example, it is possible that with advanced age, GH regulation at the pituitary or hypothalamus level is under the control of genetic determinants. Similarly, it is conceivable that certain factors which control bone strength and thereby contribute to overall skeletal fragility, are programmed for alterations during the aging process. Several studies have demonstrated that polymorphisms in specific candidate genes can modestly predict non-vertebral fractures. Examples from the literature include the collagen 1A1 gene, the IGF-I gene, the vitamin D receptor gene, and the estrogen receptor gene [123–125]. In mice, there are clear-cut differences in peak bone mass acquisition, which are maintained throughout a lifetime, in healthy inbred strains. These differences have allowed investigators to search for bone-density “genes” which contribute as much as 80% to the variance in BMD for male and female mice. Independent of estrogen status or environmental factors, some strains of inbred mice lose bone after their first year of life, while others maintain a constant density [126]. These differences are reflected in patterns of bone turnover in that the high-density mouse strain has low bone turnover and the low-density strain has markedly increased rates of bone resorption. Mouse models may become particularly useful for studying the genetic regulation of age-related bone loss for several reasons. First, inbred strains of mice produce innumerable sets of identical “twins”. Second, the mouse genome has more than 9000 markers which facilitate genotyping and permit tracking of specific loci after breeding. Third, acquisition and maintenance of bone mass in mice

parallels the same processes in humans. Fourth, the finite life span of a mouse (i.e. 2 years) makes it amenable to studies on genetic regulation of bone loss. Finally, there is a 65% sequence homology between mouse and man supporting the notion that finding mouse ‘‘osteoporosis genes’’ will facilitate the discovery of similar genes in humans.

III. MARKERS OF BONE TURNOVER AND AGE-RELATED OSTEOPOROSIS – CLINICAL IMPLICATIONS A. Overview Bone turnover is altered in patients with age-related osteoporosis. As noted previously, osteoporotic women have higher rates of bone turnover than age-matched women without osteoporosis. Similarly, elderly women with osteoporotic fractures have greater rates of bone turnover than elders without fracture, based on studies of biochemical markers such as urinary N-telopeptide, C-telopeptide, or deoxypyridinoline [28]. Such changes have been confirmed by bone histomorphometry. Although there is strong evidence that aging is associated with greater bone resorption than previously appreciated, the difference in bone formation markers between osteoporotic and nonosteoporotic elders is much less clear. Bone formation by histomorphometry is reduced in some elderly osteoporotic individuals while biochemical markers of formation are variable. Osteocalcin levels are increased in most elderly women with fractures, but this marker may reflect turnover as much as expression of osteoblast activity. Bone-specific alkaline phosphatase and procollagen peptide concentrations can be high normal or low in most patients with age-related osteoporosis. Serum IGF-I, IGFBP-3, and IGFBP-5 are synthesized by osteoblasts but serum levels of these peptides cannot be used to extrapolate activity in the bone remodeling unit. Several other non-specific skeletal parameters are altered in elderly women who sustain fractures, but these cannot be used to determine how remodeling is altered. In addition, more longitudinal studies will be needed to assess the predictive value for fracture of these markers. Table I summarizes several variables that are altered in age-related osteoporosis and their relationship to bone loss and fracture prediction.

B. Clinical Application of Biochemical Markers in the Elderly Biochemical markers are potentially very useful in understanding the pathophysiology of age-related

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CLIFFORD J. ROSEN Table I.

Age-related Osteoporosis: Markers of Bone Turnover and Calciotropic Hormones

Skeletal marker

Effects on remodeling

Bone loss

Fracture prediction

dSerum 25OHD dSerum Vit K DSerum PTH dSerum albumin DUrine NTX DUrine DpD DUrine CTX DSerum OC DBSAP DPICP dIGF-I DIGFBP-4 dIGFBP-3,-5

dformation; Dresorption dformation dformation; Dresorption No effect Dresorption Dresorption Dresorption Dturnover Dturnover Dturnover ?formation dformation dformation

++++ +1/2 ++++ – ++++ ++++ ++++ +++ + + – – –

± – ± ± ± ++ ++ – – – ?+ ?+ ??

+, the strength of evidence that a marker can predict bone loss; –, no evidence; ±, possible; ?, conceivable although there are no data. 25OHD, 25 hydroxyvitamin D; PTH, parathyroid hormone; NTX, N-telopeptide; DpD, free deoxypyridinoline; CTx, C-telopeptide; OC, osteocalcin; BSAP, bone specific alkaline phosphatase; PICP, procollagen peptide; IGF-I, insulin-like growth factor-I; IGFBP-4, insulin-like growth factor binding protein-4; IGFBP-3, -5, insulin-like growth factor binding protein -3, 5.

osteoporosis, as well as diagnosing and treating older individuals. In particular, there are three clinically relevant indications for utilizing bone markers in the older adult population: (1) to monitor bone loss in the postmenopausal period; (2) to assess overall fracture risk; and (3) to monitor response to therapy.

of three biochemical markers accounted for more than 50% of the variance in BMD noted in elderly individuals (see Figs. 2 and 3) [28]. Similarly, in one of the largest ongoing prospective studies of elderly postmenopausal women in Europe (EPIDOS), the bone markers osteocalcin, CTX, and BSAP, all were significantly greater in older

1. Monitoring Bone Loss with Biochemical Markers

Bone loss can be measured by repetitive bone density determinations. However, this does not provide immediate feedback to the patient and can be expensive as well as inconvenient. In general, women during and after menopause lose approximately 1% of their spine bone density per year. However, as many as one-third of postmenopausal women lose bone at a rate which exceeds 1% a year during the immediate menopausal period [127]. Biochemical markers can detect those women who are considered “rapid losers”, i.e. those losing upwards of 3–5% of bone per year and can predict those women most likely to respond to HRT [128]. Similar data are now available regarding bone turnover in elderly individuals. Rapid bone loss is common in many older men and women. Ensrud et al. demonstrated in the Study of Osteoporotic Fractures (SOF) that bone loss worsens with advancing age [129]. Rates of bone loss from the spine and hip can exceed 1%/year after age 70 and these changes can be detected by markers of bone turnover. In a cross-sectional study of 653 healthy European women, including many who were more than 20 years beyond the menopause, the highest levels of OC, NTX, CTX, and B-SAP were found in those individuals who were in the lowest tertile of BMD (Fig. 2) [28]. In fact, a combination

Figure 2

Contribution of the bone turnover rate to bone mass as a function of menopausal status and skeletal site. To explain the variation of BBC (total body) and BMD (total hip, distal radius, and lumbar spine), stepwise regression analyses including height, age, years since menopause, and bone markers (serum OC, serum B-ALP, serum PICP, urinary CTX, and urinary NTX) as independent variables were performed in premenopausal women (pre-MP) and in three postmenopausal groups: women within 20 years of menopause (<20 years pMP; mean age, 81 years). Each bar represents the square of the multiple correlation coefficient between bone markers that significantly contributed to BMD variance and BMD measured at several skeletal sites. Thus, each bar represents the percentage of BMC and BMD variation that is explained by the bone turnover rate.

Chapter 40 Age-related Osteoporosis and Skeletal Markers of Bone Turnover

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Figure 3 Bone turnover in elderly women as a function of quartiles of total body BMC. Whole-body BMC measured with DXA was divided into quartiles with increasing value from quartile I to quartile IV. Bars represent the mean and 1 SEM of bone marker levels in each quartile of BMC. NTX cross-linked N-telopeptide of type I collagen; CTX, type I collagen C-telopeptide breakdown products. P values apply to the comparison of trend in bone markers levels for increasing total body BMC, assessed by analysis of the variance. The dashed line represents the mean +1 SD of premenopausal women.

than younger women. Furthermore those marker values were inversely related to hip BMD [32]. More recently, Garnero et al. noted that in early postmenopausal women, those subjects with normal turnover markers lost <1% of their BMD over 4 years, whereas those in the high turnover group (i.e. more than 2 SD above the mean premenopausal range) lost three to five times that amount [27]. Interpretation of biochemical markers of bone resorption have always been plagued by the relatively low signalto-noise ratio. This analytical variability has made single determinants of turnover to assess risk of bone loss in elderly subjects somewhat problematic. For resorption markers, the coefficient of variation for free deoxypyridinoline (fDPD) is somewhat less than 15% and similarly N-telopeptide measurements in an established laboratory have a coefficient of variation of approximately 20% [67, 128]. Recently, Bollen et al. demonstrated that in elderly men and women (57 men, 69 women; mean age 73 years), urinary N-telopeptide values were consistent over time, as assessed by coefficient of concordance, which was 0.74 for women and 0.79 for men [129]. In particular, among those subjects in the highest quartile for NTX at baseline, 82% of women and 64% of men remained within that quartile in the second year and 71% of both groups in the third year. In that same longitudinal study, smoking, physical activity, race, and BMI did not affect urine NTX values [129]. Still, there are several important variables which can affect bone turnover and relate to biologic variation in the test, independent of urinary collection or creatinine

excretion. For example, osteocalcin and urinary crosslinks exhibit diurnal variation with amplitudes of up to 30% [130]. Moreover, these changes are accentuated in the elderly [28]. But, probably one of the most important variables for people living in the northern hemisphere is the effect of seasonal variation. Several prospective studies have noted that turnover markers in the elderly increase during winter months [57, 60, 65–67]. This, as noted previously, can be attributed to a decline in serum 25OH vitamin D levels as a result of reduced sunlight exposure. In turn, this leads to secondary hyperparathyroidism, which may accelerate bone turnover [41, 57, 60, 65–67, 131]. In a cross-sectional study of nearly 600 European adults (50–81 years of age), Woitge et al. noted that by multivariate regression analyses, seasonal variations accounted for more of the variability in biomarkers (up to 12%) than any of the other anthropometric or lifestyle factors except age [67]. Very recently, the same group showed that the bone loss observed in winter can be prevented by supplementation with vitamin D and calcium during the winter season [41]. Thus, age-associated bone loss is related to increased bone turnover, which can be detected by several biochemical markers. Variability in bone turnover among elderly individuals is dependent on several factors besides age; these include calcium intake, seasonal effects, timing of urine and serum collection, and coexistence of other metabolic diseases. Still, use of one or more biochemical markers to predict subsequent bone loss remains both practical and acceptable.

682 2. Assessing Fracture Risk from Biochemical Markers

For the same bone mineral density, older individuals have a much greater risk of fracture. In part this may be due to the higher rates of bone turnover which lead to bone loss, disruption of trabecular networks, and reduced connectivity. Hence, a relationship between biochemical markers of bone turnover and fracture risk has been investigated. As noted above, there is significant variation in biochemical markers among individual osteoporotic subjects. This limits, for the present time, definitive conclusions about relative risk based on a single biochemical marker, however, there are several provocative studies, which are important. For example, in hip fracture patients sampled immediately after the injury, bone resorption markers were found to be markedly increased [132]. Although this could be a function of other factors which affect hip fracture risk, including decreased mobility, weight loss, protein undernutrition, or acute trauma, there is certainly a suggestion that turnover might be increased in patients sustaining a hip fracture. Stronger data are available from prospective studies of older women in large cohorts [32, 133, 134]. For example, in a 12-year followup study Hanson et al. reported that vertebral fractures occurred most commonly in those subjects with the highest rate of bone turnover as measured by osteocalcin [133]. In one of the largest prospective studies to date, EPIDOS, which includes more than 7500 women, baseline urinary C-Telopeptide (CTX) and free deoxypyridinoline (fDPD) in subsequent hip fracture patients were noted to be significantly greater than age-matched controls [32]. Moreover, for CTX values above the premenopausal range, there was a two-fold greater risk of hip fracture which was independent of femoral bone mineral density and mobility status. When the two risk factors were combined (i.e. low femoral BMD and high C-Tx) the relative risk of a hip fracture was nearly five-fold. On the other hand, in this cohort markers of bone formation were not predictive of hip fracture risk. However, in a Swedish population study, a decrease in the carboxy-terminal propeptide of collagen (PICP) was associated with a greater risk of future fracture [135]. A similar finding was recently reported by Gerdhem et al., who followed 1040 women over the age of 75 [136]. One hundred and seventy-eight women sustained at least one fracture, urinary osteocalcin fragments and serum Trap 5B, a measure of bone resorption, strongly predicted the occurrence of any fracture [136]. In sum, elderly people, especially those who are calcium deficient, can suffer rapid bone loss. This is principally related to an increase in bone resorption. Increased bone turnover may be associated with a greater risk of vertebral and femoral fractures independent of bone density although

CLIFFORD J. ROSEN

the mechanisms responsible for the structural changes are likely more related to changes in bone quality rather than bone quantity. These data would suggest that one mechanism whereby antiresorptive therapy may reduce fracture risk is through reduction in bone turnover. Whether this effect is more important than a secondary rise in BMD remains to be determined. 3. Monitoring Antiosteoporosis Therapy with Markers

The gold standard for assessing the effects of treatment on bone mass is follow-up bone density measurements. The timing and the frequency of repeat determinations remain major issues not only scientifically but from a cost-effectiveness standpoint. Since current therapy for osteoporosis centers on inhibiting bone turnover with agents such as calcium, calcitonin, estrogens, or bisphosphonates, baseline and follow-up biochemical markers of turnover could be particularly useful. In fact, almost all the antiosteoporotic agents cause a marked decrease in bone resorption, which is detectable by conventional assays. In healthy early postmenopausal women treated with hormone replacement or calcium supplementation, Rosen et al. reported that the percent change in NTX from baseline to 6 months was the strongest predictor of subsequent spine BMD [128]. Similarly, Riis et al. reported that bone turnover markers decline by 50–100% in postmenopausal women treated with HRT for at least 6 months [137]. For bisphosphonate-treated older postmenopausal women, urinary markers of resorption declined by at least 50% within 3 months of alendronate therapy. A marked decline in bone resorption indices has also been noted in women treated with nasal calcitonin [138]. Like resorption indices, formation markers are also suppressed by antiresorptive therapy. Garnero et al. reported very significant correlations between the percent change in formation markers (BSAP, OC, and PICP) after 3 months of alendronate and spinal BMD at 24 months [139]. It should be noted that changes in biochemical markers in response to antiosteoporosis therapies may differ by treatment regimen and vary considerably even in the same individual. For example, bisphosphonates strongly decrease the urinary excretion of peptide-bound collagen products (NTX, CTX), whilst this effect is less pronounced for urinary free cross-link [140]. On the other hand, HRT suppresses both free and peptide-bound cross-links to the same extent. The problem of biologic variation for markers was alluded to earlier and remains a major concern for clinicians who want to use sequential markers as an indicator of therapeutic efficacy (see also Chapters 32, 33, 34 and 35). Signal-to-noise ratios can be enhanced by reducing intrinsic variability in measurements. For example,

Chapter 40 Age-related Osteoporosis and Skeletal Markers of Bone Turnover

either 24-hour sampling or collecting second-void 2-hour morning samples (i.e. the time of peak excretion) reduces inter-sample variation. Bone turnover changes due to seasonal variation should also be considered [41, 60, 67, 68]. Still, most people exhibit more than a 50% decline in bone resorption markers during therapy with calcitonin, estrogen, or the bisphosphonates. Hence, some level of confidence can be attained in most situations if there is consistent suppression of bone resorption and the patient is compliant with the medication.

IV. SUMMARY Bone turnover is increased in elderly individuals. In part this can be related to reduced calcium and vitamin D intake, as well as other environmental variables. Age is an independent predictor of osteoporotic fracture and there are now data to suggest that accelerated bone turnover itself may predispose an individual to future fractures, most likely as a function of changes in bone quality as well as quantity. Still, it is not clear yet that a single marker of bone turnover, or a calciotropic hormone will be able to predict which elderly individual will sustain an osteoporotic fracture. Newer serum markers such as RANKL, OPG, and Trap 5B hold some promise as future indices of fracture risk and overall bone loss. These markers will have to be studied in large randomized trials, some of which are currently ongoing. Since the osteoporotic disorder is multifactorial in origin, there are likely to be a host of parameters which can be used to identify those patients at greatest risk. Some of these indices also will help provide insight into the pathogenesis of this disease and could be useful in following patients undergoing treatment with antiresorptive drugs.

References 1. Hui, S. L., Slemenda, C. W., and Johnston, C. C., Jr. (1988). Age and bone mass as predictors of fracture in a prospective study. J. Clin. Invest. 81, 1804–1809. 2. Kiel, D. P. (1996). The approach to osteoporosis in the elderly patient. In: Osteoporosis: Diagnostic and Therapeutic Principles, C. J. Rosen (ed.), vol. 1, pp. 225–238. Humana Press, Totowa, NJ. 3. Melton, L. J., 3rd (1993). Hip fractures: a worldwide problem today and tomorrow. Bone 14 Suppl 1: S1–8. 4. Miller, P. D., Bonnick, S. L., and Rosen, C. J. (1996). Consensus of an international panel on the clinical utility of bone mass measurements in the detection of low bone mass in the adult population. Calcif. Tissue Int. 58, 207–214. 5. Cummings, S. R., Kelsey, J. L., Nevitt, M. C., and O’Dowd, K. J. (1985). Epidemiology of osteoporosis and osteoporotic fractures. Epidemiol. Rev. 7, 178–208.

683

6. Looker, A. C., Johnston. C. C. Jr., Wahner, H. W., Dunn, W. L., Calvo, M. S., Harris, T. B., Heyse, S. P., and Lindsay, R. L. (1995). Prevalence of low femoral bone density in older U.S. women from NHANES III. J. Bone Miner. Res. 10, 796–802. 7. Parker, M. J., and Palmer, C. R. (1995). Prediction of rehabilitation after hip fracture. Age Ageing 24, 96–98. 8. Keene, G. S., Parker, M. J., and Pryor, G. A. (1993). Mortality and morbidity after hip fractures. BMJ 307, 1248–1250. 9. Campbell, A. J., Borrie, M. J., and Spears, G. F. (1989). Risk factors for falls in a community-based prospective study of people 70 years and older. J. Gerontol. 44, M112–117. 10. Tinetti, M. E., Speechley, M., and Ginter, S. F. (1988). Risk factors for falls among elderly persons living in the community. N. Engl. J. Med. 319, 1701–1707. 11. Nevitt, M. C., Cummings, S. R., and Hudes, E. S. (1991). Risk factors for injurious falls: a prospective study. J. Gerontol. 46, M164–170. 12. Greenspan, S. L., Myers, E. R., Maitland, L. A., Resnick, N. M., and Hayes, W. C. (1994). Fall severity and bone mineral density as risk factors for hip fracture in ambulatory elderly. JAMA 271, 128–133. 13. Chesnut, C. H., 3rd, and Rose, C. J. (2001). Reconsidering the effects of antiresorptive therapies in reducing osteoporotic fracture. J. Bone Miner. Res. 16, 2163–2172. 14. Rucker, D., Ezzat, S., Diamandi, A., Khosravi, J., and Hanley, D. A. (2004). IGF-I and testosterone levels as predictors of bone mineral density in healthy, community-dwelling men. Clin. Endocrinol. (Oxf) 60, 491–499. 15. Chesnut, C., Majumdar, S., Gardner, J., Shields, A., Newitt, D. C., Erickson, E., Glott, M., Kriegman, A., and Mindeholm, L. (2001). Assessment of bone quality, quantity, and turnover with multiple methodologies at multiple skeletal sites. Adv. Exp. Med. Biol. 496, 95–97. 16. Vellas, B., Baumgartner, R. N., Wayne, S. J., Conceicao, J., Lafont, C., Albarede, J. L., and Garry, P. J. (1992). Relationship between malnutrition and falls in the elderly. Nutrition 8, 105–108. 17. Heaney, R. P. (1993). Nutritional factors in osteoporosis. Annu. Rev. Nutr. 13, 287–316. 18. Bastow, M. D., Rawlings, J., and Allison, S. P. (1983). Benefits of supplementary tube feeding after fractured neck of femur: a randomised controlled trial. BMJ (Clin. Res. Ed.) 287, 1589–1592. 19. Ceder, L., Thorngren, K. G., and Wallden, B. (1980). Prognostic indicators and early home rehabilitation in elderly patients with hip fractures. Clin. Orthop. 152, 173–184. 20. Rosen, C. J., Donahue, L. R., and Hunter, S. J. (1994). Insulin-like growth factors and bone: the osteoporosis connection. Proc. Soc. Exp. Biol. Med. 206, 83–102. 21. Donahue, L. R., Hunter, S. J., Sherblom, A. P., and Rosen, C. (1990). Age-related changes in serum insulin-like growth factor-binding proteins in women. J. Clin. Endocrinol. Metab. 71, 575–579. 22. Estivariz, C. F., and Ziegler, T. R. (1997). Nutrition and the insulin-like growth factor system. Endocrine 7, 65–71. 23. Bauer, D. C., Rosen, C. J., Cauley, J., and Cummings, S. R. (1997). Low serum IGF-I but not IGFBP-3 predicts hip and spine fractures. Bone Sup:561. 24. Gamero, P., Sornay-Rendu, E., and Delmas, P. D. (2000). Low serum IGF-1 and occurrence of osteoporotic fractures in postmenopausal women. Lancet 355, 898–899. 25. Schurch, M.A., Rizzoli, R., Slosman, D., Vadas, L., Vergnaud, P., and Bonjour, J. P. (1998). Protein supplements increase serum insulin-like growth factor-I levels and attenuate proximal femur bone loss in patients with recent hip fracture. A randomized, double-blind, placebocontrolled trial. Ann. Intern. Med. 128, 801–809. 26. Grinspoon, S., Miller, K., Herzog, D., Clemmons, D., and Klibanski, A. (2003). Effects of recombinant human insulin-like growth factor (IGF)-I and estrogen administration on IGF-I, IGF binding protein

684

27.

28.

29. 30.

31. 32.

33.

34.

35.

36.

37.

38.

39.

40.

41.

42.

43.

(IGFBP)-2, and IGFBP-3 in anorexia nervosa: a randomizedcontrolled study. J. Clin. Endocrinol. Metab. 88, 1142–1149. Delmas, P., Eastell, R., Garnero, P., Seibel, M., and Stepan, J. (2000). The use of biochemical markers of bone turnover in osteoporosis. Osteoporos. Int. 6 Suppl, S2–S17. Garnero, P., Sornay-Rendu, E., Chapuy, M. C., and Delmas, P. D. (1996). Increased bone turnover in late postmenopausal women is a major determinant of osteoporosis. J. Bone Miner. Res. 11, 337–349. Conference, N.C. (1995). Optimal calcium intake. JAMA 272, 1942–1948. McKane, W. R., Khosla, S., Egan, K. S., Robins, S. P., Burritt, M. F., and Riggs, B. L. (1996). Role of calcium intake in modulating agerelated increases in parathyroid function and bone resorption. J. Clin. Endocrinol. Metab. 81, 1699–1703. Heaney, R. P. (1991). Effect of calcium on skeletal development, bone loss, and risk of fractures. Am. J. Med. 91, 23S–28S. Garnero, P., Hausherr, E., Chapuy, M. C., Marcelli, C., Grandjean, H., Muller, C., Cormier, C., Breart, G., Meunier, P. J., and Delmas, P. D. (1996). Markers of bone resorption predict hip fracture in elderly women: the EPIDOS Prospective Study. J. Bone Miner. Res. 11, 1531–1538. Schett, G., Kiechl, S., Redlich, K., Oberhollenzer, F., Weger, S., Egger, G., Mayr, A., Jocher, J., Xu, Q., Pietschmann, P., Teitelbaum, S., Smolen, J., and Willeit, J. (2004). Soluble RANKL and risk of nontraumatic fracture. JAMA 291, 1108–1113. Sarkar, S., Reginster, J. Y., Crans, G. G., Diez-Perez, A., Pinette, K. V., and Delmas, P. D. (2004). Relationship between changes in biochemical markers of bone turnover and BMD to predict vertebral fracture risk. J. Bone Miner. Res. 19, 394–401. Dresner-Pollak, R., Parker, R. A., Poku, M., Thompson, J., Seibel, M. J., and Greenspan, S. L. (1996). Biochemical markers of bone turnover reflect femoral bone loss in elderly women. Calcif. Tissue Int. 59, 328–333. Orwoll, E. S., and Meier, D. E. (1986). Alterations in calcium, vitamin D, and parathyroid hormone physiology in normal men with aging: relationship to the development of senile osteopenia. J. Clin. Endocrinol. Metab. 63, 1262–1269. Sahota, O., Mundey, M. K., San, P., Godber, I. M., Lawson, N., and Hosking, D. J. (2004). The relationship between vitamin D and parathyroid hormone: calcium homeostasis, bone turnover, and bone mineral density in postmenopausal women with established osteoporosis. Bone 35, 312–319. Dawson-Hughes, B., Harris, S. S., Rasmussen, H., Song, L., and Dallal, G. E. (2004). Effect of dietary protein supplements on calcium excretion in healthy older men and women. J. Clin. Endocrinol. Metab. 89, 1169–1173. Heaney, R. P., McCarron, D. A., Dawson-Hughes, B., Oparil, S., Berga, S. L., Stern, J. S., Barr, S. I., and Rosen, C. J. (1999). Dietary changes favorably affect bone remodeling in older adults. J. Am. Diet. Assoc. 99, 1228–1233. Ledger, G. A., Burritt, M. F., Kao, P. C., O’Fallon, W. M., Riggs, B. L., and Khosla, S. (1994). Abnormalities of parathyroid hormone secretion in elderly women that are reversible by short term therapy with 1,25-dihydroxyvitamin D3. J. Clin. Endocrinol. Metab. 79, 211–216. Meier, C., Woitge, H. W., Witte, K., Lemmer, B., and Seibel, M. J. (2004). Supplementation with oral vitamin D3 and calcium during winter prevents seasonal bone loss: a randomized controlled open-label prospective trial. J. Bone Miner. Res. 19, 1221–1230. Sherman, S. S., Hollis, B. W., and Tobin, J. D. (1990). Vitamin D status and related parameters in a healthy population: the effects of age, sex, and season. J. Clin. Endocrinol. Metab. 71, 405–413. Compston, J. E., Silver, A. C., Croucher, P. I., Brown, R. C., and Woodhead, J. S. (1989). Elevated serum intact parathyroid hormone levels in elderly patients with hip fracture. Clin. Endocrinol. (Oxf) 31, 667–672.

CLIFFORD J. ROSEN 44. Chapuy, M. C., Arlot, M. E., Duboeuf, F., Brun, J., Crouzet, B., Arnaud, S., Delmas, P. D., and Meunier, P. J. (1992). Vitamin D3 and calcium to prevent hip fractures in the elderly women. N. Engl. J. Med. 327, 1637–1642. 45. Kamel, S., Brazier, M., Picard, C., Boitte, F., Samson, L., Desmet, G., and Sebert, J. L. (1994). Urinary excretion of pyridinolines crosslinks measured by immunoassay and HPLC techniques in normal subjects and in elderly patients with vitamin D deficiency. Bone Miner. 26, 197–208. 46. LeBoff, M. S., Kohlmeier, L., Hurwitz, S., Franklin, J., Wright, J., and Glowacki, J. (1999). Occult vitamin D deficiency in postmenopausal US women with acute hip fracture. JAMA 281, 1505–1511. 47. Nuti, R., Martini, G., Valenti, R., Gambera, D., Gennari, L., Salvadori, S., and Avanzati, A. (2004). Vitamin D status and bone turnover in women with acute hip fracture. Clin. Orthop. 422, 208–213. 48. Mohan, S., Farley, J. R., and Baylink, D. J. (1995). Age-related changes in IGFBP-4 and IGFBP-5 levels in human serum and bone: implications for bone loss with aging. Prog. Growth Factor Res. 6, 465–473. 49. Mohan, S., and Baylink, D. J. (1997). Serum insulin-like growth factor binding protein (IGFBP)-4 and IGFBP-5 levels in aging and ageassociated diseases. Endocrine 7, 87–91. 50. Rosen, C., Donahue, L. R., Hunter, S., Holick, M., Kavookjian, H., Kirschenbaum, A., Mohan, S., and Baylink, D. J. (1992). The 24/25-kDa serum insulin-like growth factor-binding protein is increased in elderly women with hip and spine fractures. J. Clin. Endocrinol. Metab. 74, 24–27. 51. Karasik, D., Rosen, C. J., Hannan, M. T., Broe, K. E., Dawson-Hughes, B., Gagnon, D. R., Wilson, P. W., Visser, M., Langlois, J. A., Mohan, S., and Kiel, D. P. (2002). Insulin-like growth factor binding proteins 4 and 5 and bone mineral density in elderly men and women. Calcif. Tissue Int. 71, 323–328. 52. Holbrook, T. L., Barrett-Connor, E., and Wingard, D. L. (1988). Dietary calcium and risk of hip fracture: 14-year prospective population study. Lancet 2, 1046–1049. 53. Wickham, C. A., Walsh, K., Cooper, C., Barker, D. J., Margetts, B. M., Morris, J., and Bruce, S. A. (1989). Dietary calcium, physical activity, and risk of hip fracture: a prospective study. BMJ 299, 889–892. 54. Devine, A., Dick, I. M., Dhaliwal, S. S., Naheed, R., Beilby, J., and Prince, R. L. (2005). Prediction of incident osteoporotic fractures in elderly women using the free estradiol index. Osteoporos. Int. 16, 216–221. 55. Dawson-Hughes, B., Dallal, G. E., Krall, E. A., Sadowski, L., Sahyoun, N., and Tannenbaum, S. (1990). A controlled trial of the effect of calcium supplementation on bone density in postmenopausal women. N. Engl. J. Med. 323, 878–883. 56. Recker, R. R., Hinders, S., Davies, K. M., Heaney, R. P., Stegman, M. R., Lappe, J. M., and Kimmel, D. B. (1996). Correcting calcium nutritional deficiency prevents spine fractures in elderly women. J. Bone Miner. Res 11, 1961–1966. 57. Dawson-Hughes, B., Harris, S. S., Krall, E. A., and Dallal, G. E. (1997). Effect of calcium and vitamin D supplementation on bone density in men and women 65 years of age or older. N. Engl. J. Med. 337, 670–676. 58. Ju, H. S., Leung, S., Brown, B., Stringer, M. A., Leigh, S., Scherrer, C., Shepard, K., Jenkins, D., Knudsen, J., and Cannon, R. (1997). Comparison of analytical performance and biological variability of three bone resorption assays. Clin. Chem. 43, 1570–1576. 59. Lo, C. W., Paris, P. W., Clemens, T. L., Nolan, J., and Holick, M. F. (1985). Vitamin D absorption in healthy subjects and in patients with intestinal malabsorption syndromes. Am. J. Clin. Nutr. 42, 644–649. 60. Rosen, C. J., Morrison, A., Zhou, H., Storm, D., Hunter, S. J., Musgrave, K., Chen, T., Wei, W., and Holick, M. F. (1994). Elderly women in northern New England exhibit seasonal changes in bone mineral density and calciotropic hormones. Bone Miner. 25, 83–92.

Chapter 40 Age-related Osteoporosis and Skeletal Markers of Bone Turnover 61. Prince, R. L. (1997). Diet and the prevention of osteoporotic fractures. N. Engl. J. Med. 337, 701–702. 62. Holick, M. F. (1994). McCollum Award Lecture, 1994: vitamin D—new horizons for the 21st century. Am. J. Clin. Nutr. 60, 619–630. 63. Reichel, H., Koeffler, H. P., and Norman, A. W. (1989). The role of the vitamin D endocrine system in health and disease. N. Engl. J. Med. 320, 980–991. 64. Gallagher, J. C., Riggs, B. L., Eisman, J., Hamstra, A., Arnaud, S. B., and DeLuca, H. F. (1979). Intestinal calcium absorption and serum vitamin D metabolites in normal subjects and osteoporotic patients: effect of age and dietary calcium. J. Clin. Invest. 64, 729–736. 65. Webb, A. R., Pilbeam, C., Hanafin, N., and Holick, M. F. (1990). An evaluation of the relative contributions of exposure to sunlight and of diet to the circulating concentrations of 25-hydroxyvitamin D in an elderly nursing home population in Boston. Am. J. Clin. Nutr. 51, 1075–1081. 66. Holick, M. F., Matsuoka, L. Y., and Wortsman, J. (1989). Age, vitamin D, and solar ultraviolet. Lancet 2, 1104–1105. 67. Woitge, H. W., Scheidt-Nave, C., Kissling, C., Leidig-Bruckner, G., Meyer, K., Grauer, A., Scharla, S. H., Ziegler, R., and Seibel, M. J. (1998). Seasonal variation of biochemical indexes of bone turnover: results of a population-based study. J. Clin. Endocrinol. Metab. 83, 68–75. 68. Storm, D., Eslin, R., Porter, E. S., Musgrave, K., Vereault, D., Patton, C., Kessenich, C., Mohan, S., Chen, T., Holick, M. F., and Rosen, C. J. (1998). Calcium supplementation prevents seasonal bone loss and changes in biochemical markers of bone turnover in elderly New England women: a randomized placebo-controlled trial. J. Clin. Endocrinol. Metab. 83, 3817–3825. 69. Sambrook, P. N., Chen, J. S., March, L. M., Cameron, I. D., Cumming, R. G., Lord, S. R., Zochling, J., Sitoh, Y. Y., Lau, T. C., Schwarz, J., and Seibel, M. J. (2004). Serum parathyroid hormone predicts time to fall independent of vitamin D status in a frail elderly population. J. Clin. Endocrinol. Metab. 89, 1572–1576. 70. Sambrook, P. N., Chen, J. S., March, L. M., Cameron, I. D., Cumming, R. G., Lord, S. R., Zochling, J., Sitoh, Y. Y., Lau, T. C., Schwarz, J., and Seibel, M. J. (2004). Serum PTH predicts mortality in the frail elderly. J. Clin. Endoc. Metab. (in press). 71. Hara, K., Akiyama, Y., Nakamura, T., Murota, S., and Morita, I. (1995). The inhibitory effect of vitamin K2 (menatetrenone) on bone resorption may be related to its side chain. Bone 16, 179–184. 72. Doppelt, S. H., Neer, R. M., Daly, M., Bourret, L., Schiller, A. L., and Holick, M. F. (1983). Vitamin D deficiency and ostemalacia in patients with hip fractures. Orthop. Trans. 512–513. 73. Holick, M. F. (1995). Vitamin D in health and prevention of metabolic bone disease. In: Osteoporosis: Diagnostic and Therapeutic Principles. C. J. Rosen (ed.), pp 29–35. Humana Press, Totowa, NJ. 74. Trivedi, D. P., Doll, R., and Khaw, K. T. (2003). Effect of four monthly oral vitamin D3 (cholecalciferol) supplementation on fractures and mortality in men and women living in the community: randomised double blind controlled trial. BMJ 326, 469. 75. Szulc, P., Arlot, M., Chapuy, M. C., Duboeuf, F., Meunier, P. J., and Delmas, P. D. (1994). Serum undercarboxylated osteocalcin correlates with hip bone mineral density in elderly women. J. Bone Miner. Res. 9, 1591–1595. 76. Hodges, S. J., Akesson, K., Vergnaud, P., Obrant, K., and Delmas, P. D. (1993). Circulating levels of vitamins K1 and K2 decreased in elderly women with hip fracture. J. Bone Miner. Res. 8, 1241–1245. 77. Booth, S., Tucker, K., Chen, H., Hannan, M., Gagnon, D., Cupples, L., Wilson, P., Ordovas, J., Schaefer, E., Dawson-Hughes, B., and Kiel, D. (2000). Dietary vitamin K intakes are associated with hip fracture but not with bone mineral density in elderly men and women. Am. J. Clin. Nutr. 71, 1201–1208. 78. McKeown, N. M., Jacques, P. F., Gundberg, C. M., Peterson, J. W., Tucker, K. L., Kiel, D. P., Wilson, P. W., and Booth, S. L. (2002).

79.

80.

81.

82.

83.

84. 85.

86. 87.

88.

89.

90.

91.

92.

93.

94. 95.

96.

97.

98.

685

Dietary and nondietary determinants of vitamin K biochemical measures in men and women. J. Nutr. 132, 1329–1334. Szulc, P., Chapuy, M. C., Meunier, P. J., and Delmas, P. D. (1993). Serum undercarboxylated osteocalcin is a marker of the risk of hip fracture in elderly women. J. Clin. Invest. 91, 1769–1774. Knapen, M. H., Jie, K. S., Hamulyak, K., and Vermeer, C. (1993). Vitamin K-induced changes in markers for osteoblast activity and urinary calcium loss. Calcif. Tissue Int. 53, 81–85. Kohlmier, M., Saupe, J., Shearer, M. J., Shaefer, I. K., and Asmus, G. (1996). Bone fracture risk in hemodialysis patients is related to apolipoprotein E genotype, a modulator of vitamin K status. Osteoporos. Int. 6(S1): 89. Rosen, C. J. (1994). Growth hormone, insulin-like growth factors, and the senescent skeleton: Ponce de Leon’s Fountain revisited? J. Cell. Biochem. 56, 348–356. Yen, S. S., Morales, A. J., and Khorram, O. (1995). Replacement of DHEA in aging men and women. Potential remedial effects. Ann. N Y Acad. Sci. 774, 128–142. Vanderschueren, D. (1991). Androgens in the aging male. J. Clin. Endocrinol. Metab. 73, 221–224. Jackson, J. J., Riggs, B., and Spiekerman, A. (1992). Testosterone deficiency as a risk factor for hip fractures in men: a case control study. Am. J. Med. Sci. 304, 4–8. Orwoll, E. S., and Klein, R. F. (1995). Osteoporosis in men. Endocr. Rev. 16, 87–116. Finkelstein, J. S., Klibanski, A., Neer, R. M., Greenspan, S. L., Rosenthal, D. I., and Crowley, W. F., Jr. (1987). Osteoporosis in men with idiopathic hypogonadotropic hypogonadism. Ann. Intern. Med. 106, 354–361. Snyder, P. J., Peachey, H., Hannoush, P., Berlin, J. A., Loh, L., Lenrow, D. A., Holmes, J. H., Dlewati, A., Santanna, J., Rosen, C. J., and Strom, B. L. (1999). Effect of testosterone treatment on body composition and muscle strength in men over 65 years of age. J. Clin. Endocrinol. Metab. 84, 2647–2653. Lormeau, C., Soudan, B., d’Herbomez, M., Pigny, P., Duquesnoy, B., and Cortet, B. (2004). Sex hormone-binding globulin, estradiol, and bone turnover markers in male osteoporosis. Bone 34, 933–939. Manolagas, S. C., and Jilka, R. L. (1995). Bone marrow, cytokines, and bone remodeling. Emerging insights into the pathophysiology of osteoporosis. N. Engl. J. Med. 332, 305–311. Wakley, G. K., Schutte, H. D., Jr., Hannon, K. S., and Turner, R. T. (1991). Androgen treatment prevents loss of cancellous bone in the orchidectomized rat. J. Bone Miner. Res. 6, 325–330. Devogelaer, J. P., De Cooman, S., and Nagant de Deuxchaisnes, C. (1992). Low bone mass in hypogonadal males. Effect of testosterone substitution therapy, a densitometric study. Maturitas 15, 17–23. Khosla, S., Melton, L. J., 3rd, Atkinson, E. J., and O’Fallon, W. M. (2001). Relationship of serum sex steroid levels to longitudinal changes in bone density in young versus elderly men. J. Clin. Endocrinol. Metab. 86, 3555–3561. Khosla, S., Melton, L. J., 3rd, and Riggs, B. L. (2001). Estrogens and bone health in men. Calcif. Tissue Int. 69, 189–192. Meier, D. E., Orwoll, E. S., Keenan, E. J., and Fagerstrom, R. M. (1987). Marked decline in trabecular bone mineral content in healthy men with age: lack of association with sex steroid levels. J. Am. Geriatr. Soc. 35, 189–197. Gray, A., Feldman, H. A., McKinlay, J. B., and Longcope, C. (1991). Age, disease, and changing sex hormone levels in middle-aged men: results of the Massachusetts Male Aging Study. J. Clin. Endocrinol. Metab. 73, 1016–1025. Greendale, G. A., Edelstein, S., and Barrett-Connor, E. (1997). Endogenous sex steroids and bone mineral density in older women and men: the Rancho Bernardo Study. J. Bone Miner. Res. 12, 1833–1843. Davidson, B. J., Ross, R. K., Paganini-Hill, A., Hammond, G. D., Siiteri, P. K., and Judd, H. L. (1982). Total and free estrogens and

686

99. 100.

101.

102.

103.

104.

105.

106.

107.

108. 109.

110.

111. 112. 113.

114.

115.

116.

CLIFFORD J. ROSEN androgens in postmenopausal women with hip fractures. J. Clin. Endocrinol. Metab. 54, 115–120. Albright, F., Smith, P. H., and Richardson, A. M. (1941). Postmenopausal Osteoporosis. JAMA 116, 2465–2474. Lufkin, E. G., Wahner, H. W., O’Fallon, W. M., Hodgson, S. F., Kotowicz, M. A., Lane, A. W., Judd, H. L., Caplan, R. H., and Riggs, B. L. (1992). Treatment of postmenopausal osteoporosis with transdermal estrogen. Ann. Intern. Med. 117, 1–9. Lindsay, R., and Tohme, J. F. (1990). Estrogen treatment of patients with established postmenopausal osteoporosis. Obstet. Gynecol. 76, 290–295. Rossouw, J. E., Anderson, G. L., Prentice, R. L., LaCroix, A. Z., Kooperberg, C., Stefanick, M. L., Jackson, R. D., Beresford, S. A., Howard, B. V., Johnson, K. C., Kotchen, J. M., and Ockene, J. (2002). Risks and benefits of estrogen plus progestin in healthy postmenopausal women: principal results from the Women’s Health Initiative randomized controlled trial. JAMA 288, 321–333. Uebelhart, D., Schlemmer, A., Johansen, J. S., Gineyts, E., Christiansen, C., and Delmas, P. D. (1991). Effect of menopause and hormone replacement therapy on the urinary excretion of pyridinium cross-links. J. Clin. Endocrinol. Metab. 72, 367–373. Slemenda, C., Longcope, C., Peacock, M., Hui, S., and Johnston, C. C. (1996). Sex steroids, bone mass, and bone loss. A prospective study of pre-, peri-, and postmenopausal women. J. Clin. Invest. 97, 14–21. Smith, E. P., Boyd, J., Frank, G. R., Takahashi, H., Cohen, R. M., Specker, B., Williams, T. C., Lubahn, D. B., and Korach, K. S. (1994). Estrogen resistance caused by a mutation in the estrogen-receptor gene in a man. N. Engl. J. Med. 331, 1056–1061. Morishima, A., Grumbach, M. M., Simpson, E. R., Fisher, C., and Qin, K. (1995). Aromatase deficiency in male and female siblings caused by a novel mutation and the physiological role of estrogens. J. Clin. Endocrinol. Metab. 80, 3689–3698. Carani, C., Qin, K., Simoni, M., Faustini-Fustini, M., Serpente, S., Boyd, J., Korach, K. S., and Simpson, E. R. (1997). Effect of testosterone and estradiol in a man with aromatase deficiency. N. Engl. J. Med. 337, 91–95. Bilezikian, J. P., Kurland, E. S., and Rosen, C. J. (1999). Male skeletal health and osteoporosis. Trends. Endocrinol. Metab. 10, 244–250. Riggs, B. L., Khosla, S., and Melton, L. J., 3rd (1998). A unitary model for involutional osteoporosis: estrogen deficiency causes both type I and type II osteoporosis in postmenopausal women and contributes to bone loss in aging men. J. Bone Miner. Res. 13, 763–773. Veldhuis, J. D., Iranmanesh, A., and Weltman, A. (1997). Elements in the pathophysiology of diminished growth hormone (GH) secretion in aging humans. Endocrine 7, 41–48. Jones, J. I., and Clemmons, D. R. (1995). Insulin-like growth factors and their binding proteins: biological actions. Endocr. Rev. 16, 3–34. Marcus, R. (1997). Skeletal effects of growth hormone and IGF-I in adults. Endocrine 7, 53–55. Boonen, S., Aerssens, J., Dequeker, J., Nicholson, P., Cheng, X., Lowet, G., Verbeke, G., and Bouillon, R. (1997). Age-associated decline in human femoral neck cortical and trabecular content of insulin-like growth factor I: potential implications for age-related (type II) osteoporotic fracture occurrence. Calcif. Tissue Int. 61, 173–178. Boonen, S., Lesaffre, E., Dequeker, J., Aerssens, J., Nijs, J., Pelemans, W., and Bouillon, R. (1996). Relationship between baseline insulin-like growth factor-I (IGF-I) and femoral bone density in women aged over 70 years: potential implications for the prevention of age-related bone loss. J. Am. Geriatr. Soc. 44, 1301–1306. Gelato, M. C., and Frost, R. A. (1997). IGFBP-3. Functional and structural implications in aging and wasting syndromes. Endocrine 7, 81–85. Johansson, A. G., Forslund, A., Hambraeus, L., Blum, W. F., and Ljunghall, S. (1994). Growth hormone-dependent insulin-like growth

117.

118.

119.

120.

121.

122.

123.

124.

125.

126.

127. 128.

127.

129.

factor binding protein is a major determinant of bone mineral density in healthy men. J. Bone Miner. Res. 9, 915–921. Benedict, M. R., Ayers, D. C., Calore, J. D., and Richman, R. A. (1994). Differential distribution of insulin-like growth factors and their binding proteins within bone: relationship to bone mineral density. J. Bone Miner. Res. 9, 1803–1811. Cook, F., Rosen, C. J., Vereault, D., Steffens, C., Kessenich, C. R., Greenspan, S., Ziegler, T. R., Watts, N. B., Mohan, S., and Baylink, D. J. (1996). Major changes in the circulatory IGF regulatory system after hip fracture surgery. J. Bone Min. Res. 11(S): 327. Boonen, S., Rosen, C., Bouillon, R., Sommer, A., McKay, M., Rosen, D., Adams, S., Broos, P., Lenaerts, J., Raus, J., Vanderschueren, D., and Geusens, P. (2002). Musculoskeletal effects of the recombinant human IGF-I/IGF binding protein-3 complex in osteoporotic patients with proximal femoral fracture: a double-blind, placebo-controlled pilot study. J. Clin. Endocrinol. Metab. 87, 1593–1599. Murphy, M. G., Bach, M. A., Plotkin, D., Bolognese, J., Ng, J., Krupa, D., Cerchio, K., and Gertz, B. J. (1999). Oral administration of the growth hormone secretagogue MK-677 increases markers of bone turnover in healthy and functionally impaired elderly adults. The MK-677 Study Group. J. Bone Miner. Res. 14, 1182–1188. Bach, M. A., Rockwood, K., Zetterberg, C., Thamsborg, G., Hebert, R., Devogelaer, J. P., Christiansen, J. S., Rizzoli, R., Ochsner, J. L., Beisaw, N., Gluck, O., Yu, L., Schwab, T., Farrington, J., Taylor, A. M., Ng, J., and Fuh, V. (2004). The effects of MK-0677, an oral growth hormone secretagogue, in patients with hip fracture. J. Am. Geriatr. Soc. 52, 516–523. Slemenda, C. W., Christian, J. C., Williams, C. J., Norton, J. A., and Johnston, C. C., Jr. (1991). Genetic determinants of bone mass in adult women: a reevaluation of the twin model and the potential importance of gene interaction on heritability estimates. J. Bone Miner. Res. 6, 561–567. Rivadeneira, F., Houwing-Duistermaat, J. J., Beck, T. J., Janssen, J. A., Hofman, A., Pols, H. A., Van Duijn, C. M., and Uitterlinden, A. G. (2004). The influence of an insulin-like growth factor I gene promoter polymorphism on hip bone geometry and the risk of nonvertebral fracture in the elderly: The Rotterdam Study. J. Bone Miner. Res. 19, 1280–1290. Riggs, B. L., Nguyen, T. V., Melton, L. J., 3rd, Morrison, N. A., O’Fallon, W. M., Kelly, P. J., Egan, K. S., Sambrook, P. N., Muhs, J. M., and Eisman, J. A. (1995). The contribution of vitamin D receptor gene alleles to the determination of bone mineral density in normal and osteoporotic women. J. Bone Miner. Res. 10, 991–996. Rogers, J., Mahaney, M. C., Beamer, W. G., Donahue, L. R., and Rosen, C. J. (1997). Beyond one gene-one disease: alternative strategies for deciphering genetic determinants of osteoporosis. Calcif. Tissue Int. 60, 225–228. Beamer, W. G., Donahue, L. R., Rosen, C. J., and Baylink, D. J. (1996). Genetic variability in adult bone density among inbred strains of mice. Bone 18, 397–403. Christiansen, C., Riis, B. J., and Rodbro, P. (1987). Prediction of rapid bone loss in postmenopausal women. Lancet 1, 1105–1108. Rosen, C. J., Chesnut, C. H., 3rd, and Mallinak, N. J. (1997). The predictive value of biochemical markers of bone turnover for bone mineral density in early postmenopausal women treated with hormone replacement or calcium supplementation. J. Clin. Endocrinol. Metab. 82, 1904–1910. Ensrud, K. E., Palermo, L., Black, D. M., Cauley, J., Jergas, M., Orwoll, E. S., Nevitt, M.C., Fox, K. M., and Cummings, S. R. (1995). Hip and calcaneal bone loss increase with advancing age: longitudinal results from the study of osteoporotic fractures. J. Bone Miner. Res. 10, 1778–1787. Bollen, A. M., Kiyak, H. A., and Eyre, D. R. (1997). Longitudinal evaluation of a bone resorption marker in elderly subjects. Osteoporos. Int. 7, 544–549.

Chapter 40 Age-related Osteoporosis and Skeletal Markers of Bone Turnover 130. Nielsen, H. K., Brixen, K., and Mosekilde, L. (1990). Diurnal rhythm and 24-hour integrated concentrations of serum osteocalcin in normals: influence of age, sex, season, and smoking habits. Calcif. Tissue Int. 47, 284–290. 131. Chapuy, M. C., Schott, A. M., Garnero, P., Hans, D., Delmas, P. D., and Meunier, P. J. (1996). Healthy elderly French women living at home have secondary hyperparathyroidism and high bone turnover in winter. EPIDOS Study Group. J. Clin. Endocrinol. Metab. 81, 1129–1133. 132. Cheung, C. K., Panesar, N. S., Lau, E., Woo, J., and Swaminathan, R. (1995). Increased bone resorption and decreased bone formation in Chinese patients with hip fracture. Calcif. Tissue Int. 56, 347–349. 133. Hansen, M. A., Overgaard, K., Riis, B. J., and Christiansen, C. (1991). Role of peak bone mass and bone loss in postmenopausal osteoporosis: 12 year study. BMJ 303, 961–964. 134. van Daele, P. L., Seibel, M. J., Burger, H., Hofman, A., Grobbee, D. E., van Leeuwen, J. P., Birkenhäger, J. C., and Pols, H. A. P. (1996). Case control analysis of bone resorption markers, disability and hip fracture risk: The Rotterdam Study. BMJ 312, 482–483. 135. Akesson, K., Vergnaud, P., Gineyts, E., Delmas, P. D., and Obrant, K. J. (1993). Impairment of bone turnover in elderly women with hip fracture. Calcif. Tissue Int. 53, 162–169.

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136. Gerdhem, P., Ivaska, K. K., Alatalo, S. L., Halleen, J. M., Hellman, J., Isaksson, A., Pettersson, K., Vaananen, H. K., Akesson, K., and Obrant, K. J. (2004). Biochemical markers of bone metabolism and prediction of fracture in elderly women. J. Bone Miner. Res. 19, 386–393. 137. Riis, B. J., Overgaard, K., and Christiansen, C. (1995). Biochemical markers of bone turnover to monitor the bone response to postmenopausal hormone replacement therapy. Osteoporos. Int. 5, 276–280. 138. Kraenzlin, M. E., Seibel, M. J., Trechsel, U., Boerlin, V., Azria, M., Kraenzlin, C. A., and Haas, H. G. (1996). The effect of intranasal salmon calcitonin on postmenopausal bone turnover as assessed by biochemical markers: evidence of maximal effect after 8 weeks of continuous treatment. Calcif. Tissue Int. 58, 216–220. 139. Garnero, P., Shih, W. J., Gineyts, E., Karpf, D. B., and Delmas, P. D. (1994). Comparison of new biochemical markers of bone turnover in late postmenopausal osteoporotic women in response to alendronate treatment. J. Clin. Endocrinol. Metab. 79, 1693–1700. 140. Garnero, P., Gineyts, E., Arbault, P., Christiansen, C., and Delmas, P. D. (1995). Different effects of bisphosphonate and estrogen therapy on free and peptide-bound bone cross-links excretion. J. Bone Miner. Res. 10, 641–649.

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Chapter 41

Steroid-Induced Osteoporosis Ian R. Reid MD

Department of Medicine, University of Auckland, Private Bag 92019, Auckland, New Zealand

VI. Effect of Glucocorticoids on Markers of Bone Turnover VII. Evaluation of Steroid-treated Patients VIII. Treatment and Follow-up References

I. Summary II. Introduction III. Epidemiology IV. Pathogenesis V. Effects of Glucocorticoids on Histological Indices of Bone Turnover

I. SUMMARY

II. INTRODUCTION

Glucocorticoids in supraphysiological doses rapidly lead to a reduction in bone mass, up to 40% of trabecular bone being lost after several years of continuous therapy. This results in fractures in about one-third of patients treated long-term with these drugs. Histologically the most consistent changes are a reduction in bone formation, and this is probably mediated by direct glucocorticoid receptor effects on the transcription of genes for bone matrix proteins, as well as by a reduction in osteoblast proliferation and an increase in osteoblast and osteocyte apoptosis. Circulating concentrations of osteocalcin, a marker of bone formation, are very sensitive to glucocorticoid effects, being substantially reduced within hours of glucocorticoid exposure, and recovering within a day or two of its discontinuation. Changes in other specific osteoblast markers are somewhat less, and there is no clear evidence of glucocorticoid effects on biochemical indices of bone resorption. Markers have proven valuable in determining the mechanisms by which glucocorticoids impact on bone and in assessing the likely bone effects of local glucocorticoid preparations, but they are not, at the present time, established as having a role in the evaluation and management of individual patients requiring glucocorticoid therapy.

Glucocorticoid drugs, which were introduced into clinical practice 50 years ago, have proved life-saving in conditions such as asthma, and are able to substantially modify the course of many other conditions, such as rheumatoid arthritis and inflammatory bowel disease. Their more widespread use is prevented by their side-effects, including the development of osteoporosis. This chapter will review the mechanisms by which glucocorticoids produce this dramatic bone loss and summarize the changes that have been reported in biochemical markers of bone turnover. The potential role of markers in the management of steroid-treated subjects will then be discussed.

Dynamics of Bone and Cartilage Metabolism

III. EPIDEMIOLOGY Bone loss occurs rapidly following the introduction of these drugs, barely supraphysiological doses (e.g. prednisone 7.5 mg/day) reducing trabecular bone density by 8% over a 20-week period [1]. Cross-sectional studies in patients receiving treatment for several years, typically show reductions of integral bone density in the spine and proximal femur of 10–20% [2] and of approximately 689

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twice this magnitude when predominantly trabecular bone is assessed [3, 4]. Because glucocorticoids have their greatest effect on trabecular bone, fractures are most common in regions of the skeleton, which are predominantly trabecular, such as the vertebral bodies and ribs. Hip and forearm fractures are also more common [5, 6]. The increase in fracture risk is seen within a few months of the initiation of glucocorticoid therapy, and this has been confirmed in at least one randomized controlled trial [7]. In the placebo arms of two recent trials, approximately 20% of older men and postmenopausal women had vertebral fractures in the first year of steroid treatment [8, 9]. About one-third of patients have vertebral fractures after 5–10 years of glucocorticoid treatment [10], though this can be very much higher in older patients [11]. Fracture risk is related to the steroid dose, duration of use, age, body weight (inversely), and to female sex [5, 12].

IV. PATHOGENESIS Because of the widespread distribution of the glucocorticoid receptor, these agents are able to impact on bone and calcium metabolism at many levels.

A. Osteoblasts The most consistently demonstrated effects of glucocorticoids on bone are in the osteoblast. In general, glucocorticoids can be characterized as increasing osteoblast differentiation and decreasing the proliferation of these cells [13, 14], though these findings are dependent on the experimental conditions used [15]. The former effect may be partly attributable to increased production of bone morphogenic protein-6 [16], and the latter to reduced expression of cyclin-dependent kinases and cyclin-D3 together with enhanced transcription of inhibitors of cyclindependent kinases [17]. There is also evidence for glucocorticoid regulation of a number of important osteoblast genes including those for type I collagen, osteocalcin, osteopontin, fibronectin, β-1 integrin, bone sialoprotein, alkaline phosphatase, collagenase, the nuclear protooncogenes c-myc, c-fos, c-jun, and dickkopf-1 [18]. The induction of dickkopf-1 provides an antagonist to Wnt signaling, which is a crucial regulator for bone formation, so is likely to contribute to the reduced osteoblast anabolism of glucocorticoid excess. The effects on type I collagen mRNA are reflected in reduced intracellular immunoreactivity for type I procollagen in osteoblasts

treated with dexamethasone, as well as reduced release of the amino-terminal propeptide of type I pro-collagen (PINP) and the carboxy-terminal propeptide of type I pro-collagen (PICP) into their culture medium [19]. In osteoblast precursor cells, gene expression is modulated to produce a more differentiated osteoblastic phenotype, whereas in the mature osteoblast, cell proliferation and matrix synthesis are reduced with glucocorticoids. These effects may be biphasic with respect to both dose and time, inhibition of osteoblast proliferation and activity being evident at high hormone concentrations and with long exposure periods [20]. Osteoblasts produce factors that act in an autocrine manner to regulate their own activity. Insulin-like growth factors (IGFs)-1 and -2 act in this way (see also Chapter ..), and their local synthesis is inhibited by glucocorticoids. Their local activity is modulated by the interplay of specific binding proteins and there is now evidence for a reduction in the levels of the stimulatory binding proteins, IGFBP-3 and IGFBP-5, and for increased production of IGFBP-6, an inhibitor of IGF-2 activity. However, blocking the effects of endogenous IGFs does not abrogate the effect of glucocorticoids on osteoblast proliferation and collagen synthesis. Transforming growth factor-β is a further important autocrine factor modulated by glucocorticoids. Osteoblast factors play important paracrine roles in regulating the differentiation and activity of osteoclasts, through the production of osteoprotegerin and receptor activator of NF-κB ligand (RANK-L). Hofbauer et al. [21] have shown that glucocorticoids promote osteoclastogenesis by inhibiting osteoprotegerin production and by stimulating RANK-L production in a variety of osteoblastic cells models. These effects on osteoprotegerin have been confirmed by others [14]. Von Tirpitz et al. have found the same pattern of changes in circulating levels of these two proteins in patients with Crohn’s disease treated with glucocorticoids [22], and Sasaki also found that steroid treatment reduced circulating osteoprotegerin [23, 24], though this index appears to be increased in patients with Cushing’s syndrome [25]. Following heart transplantation, serum osteoprotegerin falls by almost 50%, accounts for 67% of the variance in bone loss over the first 6 months, and is related to fracture risk [26]. A major contributor to the negative effects of glucocorticoids on osteoblasts is their promotion of apoptosis, both in these cells and in osteocytes [27]. The latter may be a key contributor to the loss of skeletal strength caused by glucocorticoids, since transgenic mouse models of steroid treatment in which there is bone loss but not osteoblast/osteocyte apoptosis, do not show a loss of vertebral compressive strength [28].

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B. Osteoclasts The direct effects of glucocorticoids on osteoclasts are contradictory. There is evidence that glucocorticoids increase rodent osteoclast formation from precursor cells in bone marrow [29, 30] but also that they lead to apoptosis of mature rodent osteoclasts [31, 32]. In human cell cultures, glucocorticoids stimulate the proliferation and differentiation of osteoclast precursors but inhibit the bone-resorbing activity of mature osteoclasts [33]. These opposing effects may account for the findings in organ culture that glucocorticoids can either increase or decrease bone resorption, depending on the culture conditions [34–36]. In organ culture, glucocorticoid effects may also be contributed to by their effects on osteoprotegerin and RANK-L, as discussed above, and by inhibition of local production of cytokines such as interleukin-1 and the tumor necrosis factors, which are important regulators of bone resorption (see Chapter ..).

C. Intestinal and Renal Handling of Calcium and Phosphate Clinical studies have consistently demonstrated an inhibition of calcium absorption associated with glucocorticoid treatment. This is not mediated by changes in vitamin D metabolites and is therefore likely to represent a direct effect on the calcium transport system in the small intestine. Within weeks of glucocorticoid treatment there is a substantial rise in urine calcium excretion [37], which is not accounted for by changes in the serum ionized calcium or the glomerular filtration rate. This suggests that glucocorticoids directly regulate tubular resorption of calcium [38]. There is also evidence for malabsorption of phosphate in both the gut and renal tubule associated with glucocorticoid use [39].

glucocorticoid effects on concentrations of vitamin D binding protein [40].

E. Parathyroid Hormone A number of groups have found increases in circulating concentrations of parathyroid hormone within minutes to weeks of the initiation of steroid therapy [41], though this has not been universal. In cross-sectional studies of patients receiving chronic glucocorticoid therapy, elevations of parathyroid hormone levels of 50–100% above those of control subjects have been reported [41]. In vitro studies of parathyroid tissue from rats, cattle, and humans suggest that hyperparathyroidism may result directly from the action of glucocorticoids on the parathyroid cell, though it is also possible that calcium malabsorption in both the gut and the renal tubule contributes. In addition, there is a substantial literature suggesting an increase in osteoblast sensitivity to parathyroid hormone after glucocorticoid treatment, possibly as a result of increased G-protein expression.

F. Sex Hormones Sex hormones are important regulators of bone metabolism, and hypogonadism in either sex is associated with the development of osteoporosis. Glucocorticoids acutely depress plasma levels of testosterone in men [42], and their chronic use is associated with a dose-dependent reduction in free testosterone concentrations of approximately 50% [43]. These changes appear to result from inhibition of gonadotropin secretion and a reduction in numbers of gonadotropin-binding sites in the testis. Circulating estradiol and adrenal androgen concentrations are also reduced in men on glucocorticoids [44]. High-dose steroid therapy is associated with oligomenorrhea in women, suggesting a similar effect on the pituitary-gonadal axis.

D. Vitamin D There is little evidence to support the contention that changes in vitamin D metabolism contribute significantly to the development of steroid osteoporosis. Prospective studies of patients or normal subjects beginning steroid therapy have shown no changes in 25-hydroxyvitamin D or 24,25-dihydroxyvitamin D but significant increases in 1,25-dihydroxyvitamin D have been observed 2–15 days after initiation of therapy [39]. These are likely to be secondary to changes in parathyroid hormone and/or serum phosphate concentrations. There is no evidence for

V. EFFECTS OF GLUCOCORTICOIDS ON HISTOLOGICAL INDICES OF BONE TURNOVER The morphometric effects of glucocorticoids on bone have been reviewed by Dempster [45]. Such studies have sampled bone at different sites and studied patients with a variety of underlying conditions – both these factors may account for the heterogeneity of findings. In general,

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osteoid thickness is reduced, as is the mineral apposition rate [46–49]. The rate of formation of mineralized bone per unit of osteoid surface is usually found to be reduced [47, 50]. Consistent with this is the finding that the amount of bone replaced within each osteoclast lacuna (i.e. the wall thickness) is reduced [50, 51]. Thus, there is clear histomorphometric evidence of reduced osteoblast activity during steroid treatment. This is likely to be contributed to by the increase in cell apoptosis, referred to above. The effects on histological assessments of bone resorption are less clear-cut. Eroded surfaces are increased [46, 52]. However, much of this surface does not contain active osteoclasts, and some studies have actually shown osteoclast numbers to be decreased [53]. This has been interpreted as reflecting an inhibition of recruitment of osteoclasts to the eroded surface (leaving eroded surfaces unfilled for a greater time than normal) rather than an acceleration of bone resorption itself. This would be consistent with the data from biochemical markers of bone resorption, to be reviewed subsequently. Prospective studies in rabbits, assessed using magnetic resonance imaging, show uniform trabecular thinning without changes in network architecture [54]. Paralleling the loss in bone volume is a conversion of hematopoietic to yellow marrow in the femoral metaphysis, and atrophy of the femoral epiphyseal growth plate [55]. These effects on growth plate chondrocytes result in marked growth retardation in children, which may not be reversible after prolonged use [56]. A further skeletal effect of glucocorticoids is the development of ischemic necrosis of the femoral head. The pathogenesis of this is debated. Some believe this is indeed an ischemic phenomenon, possibly caused by fat accumulation in bone, which increases the intraosseous pressure, thus reducing perfusion [57]. Stimulation of marrow adipocyte development is a key action of glucocorticoids [58]. Others have argued that the problem is not one of necrosis, but of steroid-induced apoptosis, particularly of osteocytes [59]. The nature and cause of this condition remain unresolved at present but it is distinct from the osteoporosis these agents cause.

VI. EFFECT OF GLUCOCORTICOIDS ON MARKERS OF BONE TURNOVER A. Osteocalcin Shortly after this bone matrix protein was shown to be measurable in the circulation and to reflect bone formation rates, it was noted to be reduced by approximately 50%

in subjects treated with glucocorticoids [60]. Circulating levels of osteocalcin are inversely related to the current glucocorticoid dose and are positively related to circulating levels of 1,25-dihydroxyvitamin D. Circulating osteocalcin concentrations decline within 24 hours of the first steroid dose and rapidly return to normal following the discontinuation of steroid therapy [61]. The molecular basis of these observations has been provided by the work of Morrison et al. [62], who demonstrated that transcription of the osteocalcin gene is negatively regulated by the glucocorticoid receptor and positively by the calcitriol receptor (see also Chapter 18). There appear to be multiple glucocorticoid receptor binding sites in the osteocalcin gene promoter [63]. The physiological relevance of this is suggested by the dependence of the diurnal variation in serum osteocalcin on that of cortisol [64]. Detailed studies in sheep have indicated that glucocorticoids reduce the plasma production rate of osteocalcin and that osteocalcin plasma clearance rates are unaffected by prednisone, though they are increased by high doses of deflazacort [65]. Similar changes are seen in Cushing’s syndrome, where urine free cortisol is inversely correlated with serum osteocalcin concentrations (r = −0.68) [66]. Indeed it has been suggested that reduced osteocalcin levels are present before clinical stigmata of glucocorticoid excess are apparent [67]. The sensitivity of circulating osteocalcin concentrations to glucocorticoids has been of great value in assessing the systemic bioavailability of topically administered steroids. This has been extensively used in studies of the various inhaled glucocorticoids [68] as well as in demonstrating bone effects from glucocorticoids administered rectally [69, 70] and by intra-articular injection [71]. However, studies of osteocalcin in steroid-treated patients need to be interpreted with caution since the patient’s underlying condition, in some cases, will also influence circulating concentrations of this peptide. In rheumatoid arthritis, for example, osteocalcin levels are lower than normal even in those not receiving glucocorticoids [72].

B. Other Markers of Bone Formation The sensitivity of osteocalcin to glucocorticoids is probably a reflection of the direct regulation of the osteocalcin gene by the glucocorticoid receptor, since other osteoblast markers show smaller responses (see Table I). Thus, total alkaline phosphatase has been found to be no different from control values when studied crosssectionally [73], though in most prospective studies of glucocorticoid administration some depression of this index is detectable. Similarly, the bone iso-enzyme of alkaline

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Chapter 41 Steroid-induced Osteoporosis Table I.

Prospective Studies of the Effects of Glucocorticoids on Biochemical Markers of Bone Turnover

Reference

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Duration Formation markers Osteocalcin Total ALP Bone ALP PICP PINP Resorption markers ICTP Pyridinoline Deoxypyridinoline Hydroxyproline TRAP sCTX uCTX Urine calcium

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ALP, alkaline phosphatase; TRAP, tartrate-resistant acid phosphatase. *Changes inversely related to age – mean change, nil. Copyright IR Reid, used with permission.

phosphatase is usually depressed in prospective assessments. The carboxy-terminal propeptide of type I procollagen (PICP) is a circulating marker of type I collagen synthesis (see Chapter 32). This has been found to decrease following the introduction of glucocorticoids [74–76]. The amino-terminal propeptide of type I pro-collagen (PINP) is also a circulating marker of type I collagen synthesis and it declines with glucocorticoid therapy, though less dramatically than osteocalcin [77]. Following cure of Cushing’s syndrome, there is a prompt increase in markers of bone formation, associated with substantial increases in bone mass [78, 79].

C. Resorption Markers The short-term prospective studies reviewed above have also assessed the effects of glucocorticoids on biochemical markers of bone resorption (Table I). Across the studies, there is no consistent evidence of a change in bone resorption following glucocorticoid treatment. In a prospective study of the administration of inhaled glucocorticoids, there was evidence of increased hydroxyproline excretion [80]. Since this change is not seen with the more specific indices of bone resorption, it is possible that it represents an effect of glucocorticoids on release of hydroxyproline from other sources (such as skin or complement) or an alteration in its metabolism or excretion. Tartrate-resistant acid phosphatase activity was found to increase in two studies, but remained unchanged in another. The increases

could be interpreted as indicating a change in osteoclast function not resulting in increased bone resorption. These mixed responses parallel the mixed effects of glucocorticoids on osteoclast development and function, in vitro. Cross-sectional studies in disease states are further complicated by disease effects on markers – for instance, serum C-telopeptide of type I collagen (sCTX) has been reported to be increased in polymyalgia rheumatica [81]. Studies in Cushing’s syndrome provide mixed results with respect to resorption markers, there being evidence of both increased [66, 82] and decreased [83, 84] urinary deoxypyridinoline, sCTX, and ICTP. Following cure of Cushing’s syndrome, there is an increase in markers of bone resorption (carboxy-terminal cross-linked telopeptide of type I collagen [ICTP] and hydroxyproline) as well as those of bone formation referred to above [78, 79]. This may, to some extent, reflect the coupling of resorption and formation, and may also result from a decline in the rate of osteoclast apoptosis. It does imply that direct stimulation of bone resorption is not a feature of the severe hypercortisolism seen in these patients prior to cure. Fasting urine calcium excretion has, in the past, been used as a measure of bone resorption. In several studies, this index has been shown to rise following glucocorticoid use. However, urine calcium excretion will only reflect bone resorption when all other variables affecting it remain constant. There is clear evidence that glucocorticoids inhibit the tubular reabsorption of calcium [38]. This finding coupled to the absence of any change in the specific markers of bone matrix breakdown suggest that

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changes in urine calcium do not reliably reflect changes in bone resorption in subjects taking steroid treatment. The biochemical data are thus fairly consistent in indicating that bone resorption is not accelerated during the first few months of steroid therapy, a time during which bone loss is greatest. These data have helped resolve some of the inconsistencies in the histomorphometric studies, discussed above. Implicit in these conclusions is the assumption that glucocorticoids have no effect on the metabolism or clearance of the collagen fragments being studied. While this cannot be absolutely proven, the consistent lack of change over a range of different fragments suggests that changes in bone resorption rate are not a major contributor to steroid-induced bone loss.

VII. EVALUATION OF STEROIDTREATED PATIENTS The prediction of fractures from clinical characteristics or bone densitometry has only been studied extensively in postmenopausal women, though there is evidence in steroid-treated subjects that fracture risk is related to the steroid dose, duration of use, age, body weight (inversely), and to female sex [5, 12, 85]. In the absence of comprehensive data relating to steroid osteoporosis, most clinicians extrapolate from the data that are available in postmenopausal women to conclude that previous fractures after minimal trauma, and low bone density are both substantial risk factors for future fracture. A clinical history and directed examination are important to determine past fracture history and the extent of trauma associated with each fracture, and also to assess other risk factors, such as body weight, cigarette smoking, frequency of falls, alcohol intake, and the presence of other medical conditions. In the absence of previous low-trauma fractures, assessment of future fracture risk is based substantially on the current bone density. Bone loss is initially most marked in the spine, and assessment of spinal bone mass by quantitative computed tomography or dual-energy X-ray absorptiometry (DXA) is appropriate. There is preliminary evidence that ultrasound assessment of bone is as satisfactory as conventional bone densitometry.

A. Bone Markers The development of biochemical markers of bone turnover has been substantially driven by the hope that these would be useful in assessing an individual’s fracture risk. There is little convincing evidence that this is so

in steroid-treated subjects, though the available data are few. Baseline levels of osteocalcin and other markers, or their changes in the early months of therapy, have generally not been found to relate to subsequent bone loss in either adults [4, 72, 86, 87] or children [88]. In a group of patients with rheumatoid arthritis some of whom began hormone replacement therapy, there was a correlation (−0.37) between the change in the CTX at 3 months and the change in bone density at 2 years [72]. This finding indicates that both techniques can detect the effects of hormone replacement therapy and it probably has little relevance for predicting outcomes in individuals. In a study of patients undergoing heart transplantation, Shane and co-workers [89] showed some correlations of osteocalcin, urinary deoxypyridinoline, urinary hydroxyproline, and urinary calcium with the subsequent rates of bone loss. However, these relationships varied from one bone density site to the other, and from time to time over the period of study. In general, they accounted for less than 10% of the variance in rates of bone loss, suggesting that they are unlikely to be of clinical utility in assessing individuals. This study also suggested that low levels of either serum 25-hydroxyvitamin D or serum testosterone 3 months after transplantation were related to more rapid bone loss, though the predictive value was, as for the turnover markers, relatively low. Sambrook et al. have reported similar findings [90] in their cardiac transplant patients in whom change in bone mass was related to concentrations of both osteocalcin (directly) and testosterone (inversely). It is of interest in both of these studies that bone loss was positively related to osteocalcin concentrations whereas the effect of glucocorticoids on osteocalcin is to depress it. However, in cardiac transplant patients cyclosporin is administered as well as steroids, and this increases bone turnover. Thus, this relationship may be specific to this particular drug combination and is not necessarily generalizable to monotherapy with glucocorticoids.

VIII. TREATMENT AND FOLLOW-UP The principal treatments available for steroid osteoporosis are calcium supplementation, replacement of sex hormones in those with demonstrable deficiency, bisphosphonates, vitamin D and its metabolites, calcitonin, and fluoride. There is little evidence that measurements other than sex hormones and 25-hydroxyvitamin D are useful in selecting which therapy is most appropriate in a specific patient. The need for calcium supplementation is usually based on the baseline dietary intake, aiming for a total intake of 1000–1500 mg/day. There is some evidence that high

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intakes reduce bone resorption [91] but the effect of this on bone loss is probably relatively small. Sex hormone replacement is only appropriate in those who are hypogonadal. In premenopausal women the presence of regular menstruation is presumptive evidence of adequate sex hormone levels. In the amenorrheic, premenopausal women and in men, measurement of estradiol or testosterone, respectively, is necessary to assess gonadal function. Hormone replacement increases densities in both sexes [92–94]. Bisphosphonates are currently the treatment of choice in steroid osteoporosis. There is a number of studies showing that cyclic etidronate is effective, and this treatment has high patient acceptability since medication is only taken for 2 weeks every 3 months. A large randomized controlled trial of etidronate has now been reported [8] and demonstrated prevention of bone loss in both the lumbar spine and proximal femur in patients recently started on steroid treatment. The beneficial effects of etidronate persisted beyond the 12-month administration period. Furthermore, the study suggested that etidronate reduced fracture rates by 50% and that height loss was also diminished. A similar trial of alendronate has recently been reported [95]. This study included patients already established on steroid treatment as well as those just commencing. Beneficial effects on bone density throughout the skeleton were found with both 5 mg /day and 10 mg/day of alendronate, though the higher dose tended to be more effective. Gains in bone mass occurred independently of the duration of previous steroid use. At 2 years, there were four morphometric vertebral fractures in 59 subjects on placebo, compared with one in 143 subjects taking alendronate (P = 0.026). All fractures occurred in women. Further studies have demonstrated the effectiveness of alendronate in steroid-treated patients with sarcoidosis, in patients with Cushing’s syndrome, and in Chinese women receiving inhaled steroids. Risedronate has been evaluated in both the prevention and treatment of steroid osteoporosis [9, 96]. The changes in bone density (with respect to placebo) were comparable to those seen with etidronate and alendronate, and data from pooling the two studies [97] indicated a more than 50% reduction in fractures in the first year of treatment. Intravenous bisphosphonates are increasingly coming into use. Pamidronate has been used in this way for some years, and there is some evidence for fracture prevention by it [98]. Ibandronate has the advantage of being administered as bolus injections rather than infusions, and it produces large increases in bone density, and a >50% reduction in vertebral fractures in steroid-treated subjects [99].

Vitamin D (calciferol) supplementation is appropriate in all individuals with demonstrable deficiency – i.e. those with serum levels of 25-hydroxyvitamin D <50 nmol/L. The use of the metabolites of vitamin D has been associated with inconsistent results from clinical trials, but a recent meta-analysis suggests beneficial effects on both bone density and fracture risk, though the benefits are less than those seen with bisphosphonates. There is also evidence that vitamin K is of benefit in steroid osteoporosis [100]. Monitoring of efficacy is based on bone density measurement and fracture occurrence. The latter is an uncommon event, and thus the absence of a new fracture does not necessarily reflect the success of a management regimen. Bone density is a much more sensitive end-point, and after 12 months therapy with a bisphosphonate there is usually clear evidence of increased spinal bone density in individual patients. Conversely, if a patient is continuing to suffer multiple fractures, this is a clear indication for review of the management of that individual’s osteoporosis, whatever the bone densitometry indicates. The availability of effective interventions in this condition places a responsibility on any prescriber of glucocorticoids to assess fracture risk in patients, and to provide prophylaxis against bone loss. The widespread adoption of this strategy will result in many fewer glucocorticoid-treated patients having to accept the morbidity of multiple fractures in addition to that of their other medical conditions.

References 1. Laan, R. F. J. M., Vanriel, P. L. C. M., Vandeputte, L. B. A., Vanerning, L. J. T. O., Vanthof, M. A., and Lemmens, J. A. M. (1993). Low-dose prednisone induces rapid reversible axial bone loss in patients with rheumatoid arthritis – a randomized, controlled study. Ann. Intern. Med. 119, 963–968. 2. Reid, I. R., Evans, M. C., Wattie, D. J., Ames, R., and Cundy, T. F. (1992). Bone mineral density of the proximal femur and lumbar spine in glucocorticoid-treated asthmatic patients. Osteoporos. Int. 2, 103–105. 3. Reid, I. R., Evans, M. C., and Stapleton, J. (1992). Lateral spine densitometry is a more sensitive indicator of glucocorticoid-induced bone loss. J. Bone Mineral Res. 7, 1221–1225. 4. Reid, I. R., and Heap, S. W. (1990). Determinants of vertebral mineral density in patients receiving chronic glucocorticoid therapy. Arch. Int. Med. 150, 2545–2548. 5. van Staa, T. P., Leufkens, H. G. M., and Cooper, C. (2002). The epidemiology of corticosteroid-induced osteoporosis: A meta-analysis. Osteoporos. Int. 13, 777–787. 6. Baltzan, M. A., Suissa, S., Bauer, D. C., and Cummings, S. R. (1999). Hip fractures attributable to corticosteroid use. Study of Osteoporotic Fractures Group [letter]. Lancet 353, 1327. 7. van Everdingen, A. A., Jacobs, J. W. G., van Reesema, D. R. S., and Bijlsma, J. W. J. (2002). Low-dose prednisone therapy for patients with early active rheumatoid arthritis: Clinical efficacy, disease-modifying properties, and side effects – A randomized, double-blind, placebocontrolled clinical trial. Ann. Int. Med. 136, 1–12.

696 8. Adachi, J. D., Bensen, W. G., Brown, J., Hanley, D., Hodsman, A., Josse, R., Kendler, D. L., Lentle, B., Olszynski, W., Stemarie, L. G., Tenenhouse, A., and Chines, A. A. (1997). Intermittent etidronate therapy to prevent corticosteroid-induced osteoporosis. N. Engl. J. Med. 337, 382–387. 9. Cohen, S., Levy, R. M., Keller, M., Boling, E., Emkey, R. D., Greenwald, M., Zizic, T. M., Wallach, S., Sewell, K. L., Lukert, B. P., Axelrod, D. W., and Chines, A. A. (1999). Risedronate therapy prevents corticosteroid-induced bone loss – A twelve-month, multicenter, randomized, double-blind, placebo-controlled, parallel-group study. Arthritis Rheum. 42, 2309–2318. 10. Walsh, L. J., Wong, C. A., Oborne, J., Cooper, S., Lewis, S. A., Pringle, M., Hubbard, R., and Tattersfield, A. E. (2001). Adverse effects of oral corticosteroids in relation to dose in patients with lung disease. Thorax 56, 279–284. 11. Walsh, L. J., Lewis, S. A., Wong, C. A., Cooper, S., Oborne, J., Cawte, S. A., Harrison, T., Green, D. J., Pringle, M., Hubbard, R., and Tattersfield, A. E. (2002). The impact of oral corticosteroid use on bone mineral density and vertebral fracture. Am. J. Resp. Crit. Care Med. 166, 691–695. 12. Michel, B. A., Bloch, D. A., Wolfe, F., and Fries, J. F. (1993). Fractures in rheumatoid arthritis – an evaluation of associated risk factors. J. Rheumatol. 20, 1666–1669. 13. Walsh, S., Jordan, G. R., Jefferiss, C., Stewart, K., and Beresford, J. N. (2001). High concentrations of dexamethasone suppress the proliferation but not the differentiation or further maturation of human osteoblast precursors in vitro: relevance to glucocorticoid-induced osteoporosis. Rheumatology 40, 74–83. 14. Atkins, G. J., Kostakis, P., Pan, B. Q., Farrugia, A., Gronthos, S., Evdokiou, A., Harrison, K., Findlay, D. M., and Zannettino, A. C. W. (2003). RANKL expression is related to the differentiation state of human osteoblasts. J. Bone Min. Res. 18, 1088–1098. 15. Pereira, R. M. R., Delany, A. M., and Canalis, E. (2001). Cortisol inhibits the differentiation and apoptosis of osteoblasts in culture. Bone 28, 484–490. 16. Boden, S. D., Hair, G., Titus, L., Racine, M., McCuaig, K., Wozney, J. M., and Nanes, M. S. (1997). Glucocorticoid-induced differentiation of fetal rat calvarial osteoblasts is mediated by bone morphogenetic protein-6. Endocrinology 138, 2820–2828. 17. Rogatsky, I., Trowbridge, J. M., and Garabedian, M. J. (1997). Glucocorticoid receptor-mediated cell cycle arrest is achieved through distinct cell-specific transcriptional regulatory mechanisms. Mol. Cell. Biol. 17, 3181–3193. 18. Ohnaka, K., Taniguchi, H., Kawate, H., Nawata, H., and Takayanagi, R. (2004). Glucocorticoid enhances the expression of dickkopf-1 in human osteoblasts: novel mechanism of glucocorticoid-induced osteoporosis. Biochem. Biophys. Res. Commun. 318, 259–264. 19. Hernandez, M. V., Guanabens, N., Alvarez, L., Monegal, A., Peris, P., Riba, J., Ercilla, G., de Osaba, M. J. M., and Munoz-Gomez, J. (2004). Immunocytochemical evidence on the effects of glucocorticoids on type I collagen synthesis in human osteoblastic cells. Calcif. Tissue Int. 74, 284–293. 20. Dietrich, J. W., Canalis, E. M., Maina, D. M., and Raisz, L. G. (1979). Effects of glucocorticoids on fetal rat bone collagen synthesis in vitro. Endocrinology 104, 715–721. 21. Hofbauer, L. C., Gori, F., Riggs, B. L., Lacey, D. L., Dunstan, C. R., Spelsberg, T. C., and Khosla, S. (1999). Stimulation of osteoprotegerin ligand and inhibition of osteoprotegerin production by glucocorticoids in human osteoblastic lineage cells: Potential paracrine mechanisms of glucocorticoid-induced osteoporosis. Endocrinology 140, 4382–4389. 22. von Tirpitz, C., Epp, S., Klaus, J., Mason, R., Hawa, G., BrinskelleSchmal, N., Hofbauer, L. C., Adler, G., Kratzer, W., and Reinshagen, M. (2003). Effect of systemic glucocorticoid therapy on bone metabolism

IAN R. REID

23.

24.

25.

26.

27.

28.

29.

30.

31.

32.

33.

34.

35.

36.

37.

38.

and the osteoprotegerin system in patients with active Crohn’s disease. Eur. J. Gastroenterol. Hepatol. 15, 1165–1170. Sasaki, N., Kusano, E., Ando, Y., Yano, K., Tsuda, E., and Asano, Y. (2001). Glucocorticoid decreases circulating osteoprotegerin (OPG): possible mechanism for glucocorticoid induced osteoporosis. Nephrol. Dialysis Transplant. 16, 479–482. Sasaki, N., Kusano, E., Ando, Y., Nemoto, J., Iimura, O., Ito, C., Takeda, S., Yano, K., Tsuda, E., and Asano, Y. (2002). Changes in osteoprotegerin and markers of bone metabolism during glucocorticoid treatment in patients with chronic glomerulonephritis. Bone 30, 853–858. Ueland, T., Bollerslev, J., Godang, K., Muller, F., Froland, S. S., and Aukrust, P. (2001). Increased serum osteoprotegerin in disorders characterized by persistent immune activation or glucocorticoid excess – possible role in bone homeostasis. Eur. J. Endocrinology 145, 685–690. Fahrleitner, A., Prenner, G., Leb, G., Tscheliessnigg, K. H., PiswangerSolkner, C., Obermayer-Pietsch, B., Portugaller, H. R., Berghold, A., and Dobnig, H. (2003). Serum osteoprotegerin is a major determinant of bone density development and prevalent vertebral fracture status following cardiac transplantation. Bone 32, 96–106. Weinstein, R. S., Jilka, R. L., Parfitt, A. M., and Manolagas, S. C. (1998). Inhibition of osteoblastogenesis and promotion of apoptosis of osteoblasts and osteocytes by glucocorticoids – potential mechanisms of their deleterious effects on bone. J. Clin. Invest. 102, 274–282. O’Brien, C. A., Jia, D., Plotkin, L. I., Bellido, T., Powers, C. C., Stewart, S. A., Manolagas, S. C., and Weinstein, R. S. (2004). Glucocorticoids act directly on osteoblasts and osteocytes to induce their apoptosis and reduce bone formation and strength. Endocrinology 145, 1835–1841. Shuto, T., Kukita, T., Hirata, M., Jimi, E., and Koga, T. (1994). Dexamethasone stimulates osteoclast-like cell formation by inhibiting granulocyte-macrophage colony-stimulating factor production in mouse bone marrow cultures. Endocrinology 134, 1121–1126. Kaji, H., Sugimoto, T., Kanatani, M., Nishiyama, K., and Chihara, K. (1997). Dexamethasone stimulates osteoclast-like cell formation by directly acting on hemopoietic blast cells and enhances osteoclast-like cell formation stimulated by parathyroid hormone and prostaglandin e-2. J. Bone Min. Res. 12, 734–741. Tobias, J., and Chambers, T. J. (1989). Glucocorticoids impair bone resorptive activity and viability of osteoclasts disaggregated from neonatal rat long bones. Endocrinology 125, 1290–1295. Dempster, D. W., Moonga, B. S., Stein, L. S., Horbert, W. R., and Antakly, T. (1997). Glucocorticoids inhibit bone resorption by isolated rat osteoclasts by enhancing apoptosis. J. Endocrinol. 154, 397–406. Hirayama, T., Sabokbar, A., and Athanasou, N. A. (2002). Effect of corticosteroids on human osteoclast formation and activity. J. Endocrinol. 175, 155–163. Caputo, C. B., Meadows, D., and Raisz, L. G. (1976). Failure of estrogens and androgens to inhibit bone resorption in tissue culture. Endocrinology 98, 1065–1068. Lowe, C., Gray, D. H., and Reid, I. R. (1992). Serum blocks the osteolytic effect of cortisol in neonatal mouse calvaria. Calcif. Tissue Int. 50, 189–192. Gronowicz, G., McCarthy, M. B., and Raisz, L. G. (1990). Glucocorticoids stimulate resorption in fetal rat parietal bones in vitro. J. Bone Mineral Res. 5, 1223–1230. Gray, R. E., Doherty, S. M., Galloway, J., Coulton, L., de Broe, M., and Kanis, J. A. (1991). A double-blind study of deflazacort and prednisone in patients with chronic inflammatory disorders. Arthritis Rheum. 34, 287–295. Reid, I. R., and Ibbertson, H. K. (1987). Evidence for decreased tubular reabsorption of calcium in glucocorticoid-treated asthmatics. Hormone Res. 27, 200–204.

697

Chapter 41 Steroid-induced Osteoporosis 39. Cosman, F., Nieves, J., Herbert, J., Shen, V., and Lindsay, R. (1994). High-dose glucocorticoids in multiple sclerosis patients exert direct effects on the kidney and skeleton. J. Bone Mineral Res. 9, 1097–1105. 40. Braun, J. J., Juttman, J. R., Visser, T. J., and Birkenhager, J. C. (1982). Short-term effect of prednisone on serum 1,25-hydroxy-vitamin D in normal individuals an in hyper- and hypoparathyroidism. Clin. Endocrinol. 17, 21–28. 41. Fucik, R. F., Kukreja, S. C., Hargis, G. K., Bowser, E. N., Henderson, W. J., and Williams, G. A. (1975). Effect of glucocorticoids on function of the parathyroid glands in man. J. Clin. Endocrinol. Metab. 40, 152–185. 42. Doerr, P., and Pirke, K. M. (1976). Cortisol-induced suppression of plasma testosterone in normal adult males. J. Clin. Endocrinol. Metab. 43, 622–629. 43. Reid, I. R., France, J. T., Pybus, J., and Ibbertson, H. K. (1985). Low plasma testosterone levels in glucocorticoid-treated male asthmatics. BMJ 291, 574. 44. Hampson, G., Bhargava, N., Cheung, J., Vaja, S., Seed, P. T., and Fogelman, I. (2002). Low circulating estradiol and adrenal androgens concentrations in men on glucocorticoids: A potential contributory factor in steroid-induced osteoporosis. Metab. Clin. Exper. 51, 1458–1462. 45. Dempster, D. W. (1989). Bone histomorphometry in glucocorticoidinduced osteoporosis. J. Bone Min. Res. 4, 137–141. 46. Bressot, C., Meunier, P. J., Chapuy, M. C., Lejeune, E., Edouard, C., and Darby, A. J. (1979). Histomorphometric profile, pathophysiology and reversibility of corticosteroid-induced osteoporosis. Met. Bone Dis. Rel. Res. 1, 303–311. 47. Lund, B., Storm, T. L., Melsen, F., Mosekilde, L., Andersen, R. B., Egmose, C., and Sorensen, O. H. (1985). Bone mineral loss, bone histomorphometry and vitamin D metabolism in patients with rheumatoid arthritis on long-term glucocorticoid treatment. Clin. Rheumatol. 4, 143–149. 48. Aaron, J. E., Francis, R. M., Peacock, M., and Makins, N. B. (1989). Contrasting microanatomy of idiopathic and corticosteroid-induced osteoporosis. Clin. Orthop. 243, 294–305. 49. Chavassieux, P., Pastoureau, P., Chapuy, M. C., Delmas, P. D., and Meunier, P. J. (1993). Glucocorticoid-induced inhibition of osteoblastic bone formation in ewes: a biochemical and histomorphometric study. Osteoporos. Int. 3, 97–102. 50. Stellon, A. J., Webb, A., and Compston, J. E. (1988). Bone histomorphometry and structure in corticosteroid treated chronic active hepatitis. Gut 29, 378–384. 51. Dempster, D. W., Arlot, M. A., and Meunier, P. J. (1983). Mean wall thickness and formation periods of trabecular bone packets in corticosteroid-induced osteoporosis. Calcif. Tissue Int. 35, 410–417. 52. Gallagher, J. C., Aaron, J., Horsman, A., Wilkinson, R., and Nordin, B. E. (1973). Corticosteroid osteoporosis. Clin. Endocrinol. Metab. 2, 355–368. 53. Chavassieux, P., Buffet, A., Vergnaud, P., Garnero, P., and Meunier, P. J. (1997). Short-term effects of corticosteroids on trabecular bone remodeling in old ewes. Bone 20, 451–455. 54. Takahashi, M., Wehrli, F. W., Hilaire, L., Zemel, B. S., and Hwang, S. N. (2002). In vivo NMR microscopy allows short-term serial assessment of multiple skeletal implications of corticosteroid exposure. Proc. Nat. Acad. Scie. USA 99, 4574–4579. 55. Chrysis, D., Ritzen, E. M., and Savendahl, L. (2003). Growth retardation induced by dexamethasone is associated with increased apoptosis of the growth plate chondrocytes. J. Endocrinol. 176, 331–337. 56. Abad, V., Chrousos, G. P., Reynolds, J. C., Nieman, L. K., Hill, S. C., Weinstein, R. S., and Leong, G. M. (2001). Glucocorticoid excess during adolescence leads to a major persistent deficit in bone mass and an increase in central body fat. J. Bone Min. Res. 16, 1879–1885. 57. Miyanishi, K., Yamamoto, T., Irisa, T., Yamashita, A., Jingushi, S., Noguchi, Y., and Iwamoto, Y. (2002). Bone marrow fat cell enlargement

58.

59.

60.

61.

62.

63.

64.

65.

66.

67.

68.

69.

70.

71.

72.

and a rise in intraosseous pressure in steroid-treated rabbits with osteonecrosis. Bone 30, 185–190. Justesen, J., Mosekilde, L., Holmes, M., Stenderup, K., Gasser, J., Mullins, J. J., Seckl, J. R., and Kassem, M. (2004). Mice deficient in 11 beta-hydroxysteroid dehydrogenase type 1 lack bone marrow adipocytes, but maintain normal bone formation. Endocrinology 145, 1916–1925. Weinstein, R. S., Nicholas, R. W., and Manolagas, S. C. (2000). Apoptosis of osteocytes in glucocorticoid-induced osteonecrosis of the hip. J. Clin. Endocrinol. Metab. 85, 2907–2912. Reid, I. R., Chapman, G. E., Fraser, T. R., Davies, A. D., Surus, A. S., Meyer, J., Huq, N. L., and Ibbertson, H. K. (1986). Low serum osteocalcin levels in glucocorticoid-treated asthmatics. J. Clin. Endocrinol. Metab. 62, 379–383. Godschalk, M. F., and Downs, R. W. (1988). Effect of short-term glucocorticoids on serum osteocalcin in healthy young men. J. Bone Mineral Res. 3, 113–115. Morrison, N. A., Shine, J., Fragonas, J. C., Verkest, V., McMenemy, M. L., and Eisman, J. A. (1989). 1,25-dihydroxyvitamin D-responsive element and glucocorticoid repression in the osteocalcin gene. Science 246, 1158–1161. Heinrichs, A. A. J., Bortell, R., Rahman, S., Stein, J. L., Alnemri, E. S., Litwack, G., Lian, J. B., and Stein, G. S. (1993). Identification of multiple glucocorticoid receptor binding sites in the rat osteocalcin gene promoter. Biochemistry 32, 11436–11444. Heshmati, H. M., Riggs, B. L., Burritt, M. F., McAlister, C. A., Wollan, P., and Khosla, S. (1998). Effects of the circadian variation in serum cortisol on markers of bone turnover and calcium homeostasis in normal postmenopausal women. J. Clin. Endocrinol. Metab. 83, 751–756. O’Connell, S. L., Tresham, J., Fortune, C. L., Farrugia, W., McDougall, J. G., Scoggins, B. A., and Wark, J. D. (1993). Effects of prednisolone and deflazacort on osteocalcin metabolism in sheep. Calcif. Tissue Int. 53, 117–121. Scillitani, A., Dicembrino, F., Chiodini, I., Minisola, S., Fusilli, S., Di Giorgio, A., Garrubba, M., D’Aloiso, L., Frusciante, V., Torlontano, M., Modomi, S., Trischitta, V., and Carnevale, V. (2002). Global skeletal uptake of Tc-99m-methylene diphosphonate (GSU) in patients affected by endocrine diseases: Comparison with biochemical markers of bone turnover. Osteoporos. Int. 13, 829–834. Sartorio, A., Conti, A., Ferrero, S., Giambona, S., Re, T., Passini, E., and Ambrosi, B. (1998). Evaluation of markers of bone and collagen turnover in patients with active and preclinical cushings-syndrome and in patients with adrenal incidentaloma. Euro. J. Endocrinol. 138, 146–152. Richy, F., Bousquet, J., Ehrlich, G. E., Meunier, P. J., Israel, E., Morii, H., Devogelaer, J. P., Peel, N., Haim, M., Bruyere, O., and Reginster, J. Y. (2003). Inhaled corticosteroids effects on bone in asthmatic and COPD patients: a quantitative systematic review. Osteoporos. Int. 14, 179–190. Robinson, R. J., Iqbal, S. J., Wolfe, R., Patel, K., Abrams, K., and Mayberry, J. F. (1998). The effect of rectally administered steroids on bone turnover – a comparative study. Aliment. Pharmacol. Therapeut. 12, 213–217. Dhaens, G., Verstraete, A., Cheyns, K., Aerden, I., Bouillon, R., and Rutgeerts, P. (1998). Bone turnover during short-term therapy with methylprednisolone or budesonide in crohns-disease. Aliment. Pharmacol. Therapeut. 12, 419–424. Emkey, R. D., Lindsay, R., Lyssy, J., Weisberg, J. S., Dempster, D. W., and Shen, V. (1996). The systemic effect of intraarticular administration of corticosteroid on markers of bone formation and bone resorption in patients with rheumatoid arthritis. Arthritis Rheum. 39, 277–282. Hall, G. M., Spector, T. D., and Delmas, P. D. (1995). Markers of bone metabolism in postmenopausal women with rheumatoid arthritis – Effects

698

73.

74.

75.

76.

77.

78.

79.

80. 81.

82.

83.

84.

85.

86.

87.

88.

of corticosteroids and hormone replacement therapy. Arthritis Rheum. 38, 902–906. Reid, I. R., Chapman, G. E., Fraser, T. R. C., Davies, A. D., Surus, A. S., Meyer, J., Huq, N. L., and Ibbertson, H. K. (1986). Low serum osteocalcin levels in glucocorticoid-treated asthmatics. J. Clin. Endocrinol. Metab. 62, 379–383. Lems, W. F., Gerrits, M. I., Jacobs, J. W. G., Vanvugt, R. M., Vanrijn, H. J. M., and Bijlsma, J. W. J. (1996). Changes in (markers of) bone metabolism during high dose corticosteroid pulse treatment in patients with rheumatoid arthritis. Ann. Rheum. Dis. 55, 288–293. Wolthers, O. D., Heuck, C., Hansen, M., and Kollerup, G. (1997). Serum and urinary markers of types I and III collagen turnover during short-term prednisolone treatment in healthy adults. Scand. J. Clin. Lab. Invest. 57, 133–139. Bornefalk, E., Dahlen, I., Michaelsson, K., Ljunggren, O., and Ljunghall, S. (1998). Age-dependent effect of oral glucocorticoids on markers of bone resorption in patients with acute asthma. Calcif. Tissue Int. 63, 9–13. Cooper, M. S., Blumsohn, A., Goddard, P. E., Bartlett, W. A., Shackleton, C. H., Eastell, R., Hewison, M., and Stewart, P. M. (2003). 11 beta-hydroxysteroid dehydrogenase type 1 activity predicts the effects of glucocorticoids on bone. J. Clin. Endocrinol. Metab. 88, 3874–3877. Hermus, A. R., Smals, A. G., Swinkels, L. M., Huysmans, D. A., Pieters, G. F., Sweep, C. F., Corstens, F. H., and Kloppenborg, P. W. (1995). Bone mineral density and bone turnover before and after surgical cure of Cushing’s syndrome. J. Clin. Endocrinol. Metab. 80, 2859–2865. Sartorio, A., Conti, A., Ferrario, S., Passini, E., Re, T., and Ambrosi, B. (1996). Serum bone gla protein and carboxyterminal cross-linked telopeptide of type I collagen in patients with Cushings syndrome. Postgrad. Med. J. 72, 419–422. Ali, N. J., Capewell, S., and Ward, M. J. (1991). Bone turnover during high dose inhaled corticosteroid treatment. Thorax 46, 160–164. Pearce, G., Ryan, P. F. J., Delmas, P. D., Tabensky, D. A., and Seeman, E. (1998). The deleterious effects of low-dose corticosteroids on bone density in patients with polymyalgia rheumatica. Br. J. Rheumatol. 37, 292–299. Godang, K., Ueland, T., and Bollerslev, J. (1999). Decreased bone area, bone mineral content, formative markers, and increased bone resorptive markers in endogenous Cushing’s syndrome. Eur. J. Endocrinol. 141, 126–131. Francucci, C. M., Pantanetti, P., Garrapa, G. G., Massi, F., Arnaldi, G., and Mantero, F. (2002). Bone metabolism and mass in women with Cushing’s syndrome and adrenal incidentaloma. Clin. Endocrinol. 57, 587–593. Cortet, B., Cortet, C., Blanckaert, F., d’Herbomez, M., Marchandise, X., Wemeau, J. L., Decoulx, M., and Dewailly, D. (2001). Quantitative ultrasound of bone and markers of bone turnover in Cushing’s syndrome. Osteoporos. Int. 12, 117–123. Goemaere, S., Liberman, U. A., Adachi, J. D., Hawkins, F., Lane, N., Saag, K. G., Schnitzer, T., Kaufman, J. M., Malice, M. P., Carofano, W., and Daifotis, A. (2003). Incidence of nonvertebral fractures in relation to time on treatment and bone density in glucocorticoid-treated patients – A retrospective approach. J. Clin. Rheumatol. 9, 170–175. Boulet, L. P., Milot, J., Gagnon, L., Poubelle, P. E., and Brown, J. (1999). Long-term influence of inhaled corticosteroids on bone metabolism and density – Are biological markers predictors of bone loss? Am. J. Respir. Crit. Care. Med. 159, 838–844. Toogood, J. H., Hodsman, A. B., Fraher, L. J., Markov, A. E., and Baskerville, J. C. (1999). Serum osteocalcin and procollagen as markers for the risk of osteoporotic fracture in corticosteroid-treated asthmatic adults. J. Allergy Clin. Immunol. 104, 769–774. Reeve, J., Loftus, J., Hesp, R., Ansell, B. M., Wright, D. J., and Woo, P. M. M. (1993). Biochemical prediction of changes in spinal bone mass in juvenile chronic (or rheumatoid) arthritis treated with glucocorticoids. J. Rheumatol. 20, 1189–1195.

IAN R. REID 89. Shane, E., Rivas, M., McMahon, D. J., Staron, R. B., Silverberg, S. J., Seibel, M. J., Mancini, D., Michler, R. E., Aaronson, K., Addesso, V., and Lo, S. H. (1997). Bone loss and turnover after cardiac transplantation. J. Clin. Endocrinol. Metab. 82, 1497–1506. 90. Sambrook, P. N., Kelly, P. J., Fontana, D., Nguyen, T., Keogh, A., Macdonald, P., Spratt, P., Freund, J., and Eisman, J. A. (1994). Mechanisms of rapid bone loss following cardiac transplantation. Osteoporos. Int. 4, 273–276. 91. Reid, I. R., and Ibbertson, H. K. (1986). Calcium supplements in the prevention of steroid-induced osteoporosis. Am. J. Clin. Nutr. 44, 287–290. 92. Hall, G. M., Daniels, M., Doyle, D. V., and Spector, T. D. (1994). Effect of hormone replacement therapy on bone mass in rheumatoid arthritis patients treated with and without steroids. Arthritis Rheum. 37, 1499–1505. 93. Reid, I. R., Wattie, D. J., Evans, M. C., and Stapleton, J. P. (1996). Testosterone therapy in glucocorticoid-treated men. Arch. Int. Med. 156, 1173–1177. 94. Crawford, B. A. L., Liu, P. Y., Kean, M. T., Bleasel, J. F., and Handelsman, D. J. (2003). Randomized placebo-controlled trial of androgen effects on muscle and bone in men requiring long-term systemic glucocorticoid treatment. J. Clin. Endocrinol. Metab. 88, 3167–3176. 95. Adachi, J. D., Saag, K. G., Delmas, P. D., Liberman, U. A., Emkey, R. D., Seeman, E., Lane, N. E., Kaufman, J. M., Poubelle, P. E. E., Hawkins, F., Correa-Rotter, R., Menkes, C. J., Rodriguez-Portales, J. A., Schnitzer, T. J., Block, J. A., Wing, J., McIlwain, H. H., Westhovens, R., Brown, J., Melo-Gomes, J. A., Gruber, B. L., Yanover, M. J., Leite, M. O. R., Siminoski, K. G., Nevitt, M. C., Sharp, J. T., Malice, M., Dumortier, T., Czachur, M., Carofano, W., Daifotis, A. (2001). Two-year effects of alendronate on bone mineral density and vertebral fracture in patients receiving glucocorticoids – A randomized, double-blind, placebo-controlled extension trial. Arthritis Rheumat. 44, 202–211. 96. Reid, D. M., Adami, S., Emkey, R., Chines, A., Jencen, D., and Cohen, S. (1999). Risedronate increases bone mass regardless of gender or underlying condition in patients taking corticosteroids. J. Bone Min. Res. 14 suppl 1, s209. 97. Wallach, S., Cohen, S., Reid, D. M., Hughes, R. A., Hosking, D. J., Laan, R. F., Doherty, S. M., Maricic, M., Rosen, C., Brown, J., Barton, I., and Chines, A. A. (2000). Effects of risedronate treatment on bone density and vertebral fracture in patients on corticosteroid therapy. Calcif. Tissue Int. 67, 277–285. 98. Kim, S. H., Lim, S. K., and Hahn, J. S. (2004). Effect of pamidronate on new vertebral fractures and bone mineral density in patients with malignant lymphoma receiving chemotherapy. Am. J. Med. 116, 524–528. 99. Ringe, J. D., Dorst, A., Faber, H., Ibach, K., and Sorenson, F. (2003). Intermittent intravenous ibandronate injections reduce vertebral fracture risk in corticosteroid-induced osteoporosis : results from a long-term comparative study. Osteoporosis Int. 14, 801–807. 100. Yonemura, K., Fukasawa, H., Fujigaki, Y., and Hishida, A. (2004). Protective effect of vitamins K2 and D3 on prednisolone-induced loss of bone mineral density in the lumbar spine. Am. J. Kidney Diseases 43, 53–60. 101. Prummel, M. F., Wiersinga, W. M., and Lips, P. (1991). The course of biochemical parameters of bone turnover during treatment with corticosteroids. J. Clin. Endocrinol. Metab. 72, 382–386. 102. Morrison, D., Ali, N. J., Routledge, P. A., and Capewell, S. (1992). Bone turnover during short course prednisolone treatment in patients with chronic obstructive airways disease. Thorax 47, 418–420. 103. Lane, S. J., Vaja, S., Swaminathan, R., and Lee, T. H. (1996). Effects of prednisolone on bone turnover in patients with corticosteroid resistant asthma. Clin. Experiment. Allergy. 26, 1197–1201.

Chapter 41 Steroid-induced Osteoporosis 104. Lems, W. F., Vanveen, G. J. M., Gerrits, M. I., Jacobs, J. W. G., Houben, H. H. M. L., Vanrijn, H. J. M., and Bijlsma, J. W. J. (1998). Effect of low-dose prednisone (with calcium and calcitriol supplementation) on calcium and bone metabolism in healthy volunteers. Br. J. Rheumatol. 37, 27–33. 105. Pearce, G., Tabensky, D. A., Delmas, P. D., Baker, H. W. G., and Seeman, E. (1998). Corticosteroid-induced bone loss in men. J. Clin. Endocrinol. Metab. 83, 801–806.

699 106. van Everdingen, A. A., van Reesema, D. R. S., Jacobs, J. W. G., and Bijlsma, J. W. J. (2003). Low-dose glucocorticoids in early rheumatoid arthritis: Discordant effects on bone mineral density and fractures? Clin. Exp. Rheumatol. 21, 155–160.

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Chapter 42

Transplantation Osteoporosis: Biochemical Correlates of Pathogenesis and Treatment Carolina A. Moreira Kulak, M.D. and Elizabeth Shane, M.D.

Department of Medicine, College of Physicians and Surgeons, Columbia University, New York, New York 10032

VII. Mechanisms of Bone Loss after Transplantation VIII. Prevention and Management of Transplantation Osteoporosis IX. Conclusions References

I. Introduction II. Kidney Transplantation III. Cardiac Transplantation IV. Liver Transplantation V. Lung Transplantation VI. Bone Marrow Transplantation

I. INTRODUCTION

propensity to fracture, which has a negative impact upon their quality of life [2–4]. The pathogenesis of transplantation osteoporosis is complex and incompletely understood. Risk factors for osteoporosis (Caucasian race, older age, postmenopausal status, dietary calcium deficiency, vitamin D deficiency, physical inactivity, excessive use of tobacco, and alcohol) and exposure to drugs associated with bone loss (loop diuretics, anticoagulants, glucocorticoids) are common among patients who are candidates for transplantation. Moreover, bone and mineral homeostasis is also influenced by the diseased organ before transplantation, a particularly

Within the past two decades, organ transplantation has become established as effective therapy for endstage renal, hepatic, cardiac, and pulmonary disease. The kidney is the most commonly transplanted organ, followed by liver, heart, pancreas, lung, and heart–lung. Overall 1-year patient survival is excellent, ranging from 97% for living donor kidney recipients to 63% for heart–lung recipients [1]. Five-year survival is also quite good and many patients live more than 10 years after transplantation. Unfortunately, however, many transplant recipients demonstrate a Dynamics of Bone and Cartilage Metabolism

Department of Endocrinology, Federal University of Parana, Hospital de Clinicas, Curitiba, Brazil

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Copyright © 2006 by Academic Press. All rights of reproduction in any form reserved.

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common circumstance in the case of renal or hepatic failure. However, it is generally considered that posttransplantation immunosuppression plays the pivotal role in the evolution of bone loss and incidence of fracture in organ transplant recipients [4]. All transplant recipients require immunosuppressive therapy to prevent rejection of the transplanted organ. For the past decade, the majority of patients have been managed with “triple” immunosuppressive therapy. Such regimens generally include glucocorticoids, cyclosporine A (CsA), or tacrolimus (FK506) which suppress the immune response by inhibition of T cell function, and azathioprine or mycophenolate mofetil. Glucocorticoids, notorious for causing osteoporosis (see Chapter 45), are commonly prescribed in high doses (up to 60 mg of prednisone or its equivalent daily) immediately after transplantation and during episodes of severe rejection, with gradual reduction over several weeks. In addition, Epstein and colleagues have repeatedly demonstrated that administration of CsA to rats in doses comparable to and higher than those used after transplantation, causes severe trabecular bone loss [5–7]. In this model, bone turnover is accelerated, with histological evidence of increased bone resorption and bone formation. The bone loss is largely prevented by agents that inhibit bone resorption [8–10]. FK506 causes bone loss in the rat of even greater magnitude than that observed with CsA [11]. Whether CsA and FK506 contribute to the pathogenesis of transplantation osteoporosis remains controversial. Since both drugs are used in conjunction with glucocorticoids, it has been difficult to determine whether CsA has specific effects on bone and mineral metabolism in humans. However, it is generally considered that concomitant use of glucocorticoids and calcineurin inhibitors has more deleterious effects on bone and mineral metabolism than exclusive administration of either class of drug [4]. In this chapter, we will summarize the natural history of bone loss and fracture associated with kidney, heart, lung, liver, and bone marrow transplantation. The changes in calciotropic hormones and biochemical markers of bone turnover that follow various types of organ transplants will also be reviewed.

II. KIDNEY TRANSPLANTATION Most patients undergoing kidney transplantation already have at least some evidence of renal osteodystrophy. This may include hyperparathyroidism (with or without osteitis fibrosa), osteomalacia, osteosclerosis, adynamic or aplastic bone disease [12]. In some patients, more than one of these conditions may be present.

In addition, patients with end-stage renal disease are commonly hypogonadal and many have received previous therapy with medications (glucocorticoids and/or CsA for immune complex disease, loop diuretics, or aluminum-containing phosphate binders), which can affect bone and mineral metabolism. Restoration of renal function after kidney transplantation rectifies many of the disturbances that lead to renal osteodystrophy [12]. There is resolution of hyperphosphatemia, an increase in serum 1,25(OH)2D levels and a rapid decline in the high levels of PTH often observed in patients with end-stage renal disease. However, PTH levels frequently remain elevated during the immediate post-transplant period and may never completely normalize. Not surprisingly, PTH-dependent hypercalcemia occurs commonly during the first few months after kidney transplantation and hypophosphatemia is also common. In the majority of patients, hypophosphatemia is related to persistent parathyroid hyperplasia. However, in many patients, it occurs in the absence of hyperparathyroidism and may persist for years. While glucocorticoids and also rapamycin [13] increase renal phosphate losses and may contribute to the hypophosphatemia, kidney transplant recipients may also have a primary defect in renal phosphate handling, perhaps related to circulating factors (phosphatonins) that cause renal phosphate wasting [14]. Mild hypophosphatemia (>1.0 mg/dl) is generally well-tolerated and should not be treated with oral phosphate replacement, which may prolong the resolution of hyperparathyroidism and may not improve phosphate balance [15]. Immunosuppressive therapy constitutes an additional insult to a skeleton, often already compromised with respect to bone mass (see Chapter 45). Prospective studies indicate that lumbar spine bone loss during the first 3 to 18 months following renal transplantation varies from 4% to 9% at the lumbar spine and 5% to 9% at the hip (Table I). Bone loss is greatest during the first 6 months after transplantation and predominantly affects cancellous bone [16–18]. Some studies report gender differences at the site of bone loss, with men losing more bone at the hip [16, 19]. Virtually all cross-sectional studies of renal transplant recipients demonstrate that bone mass is lower than in age- and sex-matched normal populations [20–26]. One study of 20 young adults who received a renal transplant during childhood revealed that 70% had lumbar spine BMD T scores below −1.0, of whom six were below −2.0 [25]. Bone loss has not been consistently related to gender, patient age, glucocorticoid dose, rejection episodes, activity level, or PTH levels. There is also evidence for significant ongoing bone loss in long-term kidney transplant recipients, albeit less rapid

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Chapter 42 Transplantation Osteoporosis: Biochemical Correlates of Pathogenesis and Treatment Table I.

Osteoporosis after Solid Organ Transplantation

Prevalence after transplantation Type of transplant

Osteoporosis*

Fractures

Kidney+

11–56%

Heart

25–50%

Vertebral: 3–29% Peripheral: 11–43% Vertebral: 22–35%

Liver

16–46%

Vertebral: 29–47%

Lung

57–73%

42%

Bone loss: first post-transplant year

Fracture incidence

Spine: 4–9% Hip: 8% Spine: 3–8% Hip: 6–11% Spine: 0–24% Hip: 2–4% Spine: 1–5% Hip: 2–5%

Vertebral: 3–10% Peripheral: 10–50% Vertebral and non-vertebral: 10–36% Vertebral: 24–65% 18–37%

*Accepted definitions included BMD of spine and/or hip (by dual X-ray absorptiometry) > 2 SD below age- and sex-matched control or > 2.5 SD below young normal controls. +Definition of osteoporosis also included BMD of predominantly cortical sites such as the femoral shaft or proximal radius that are adversely affected by excessive PTH secretion. Adapted from Cohen, A., Ebeling, P., Sprague, S., Shane, E. (2003) Transplantation osteoporosis. In (M. J. Favus, ed), Primer on the Metabolic Bone Diseases and Disorders of Mineral Metabolism, 5th edn. American Society for Bone and Mineral Research, Washington, D.C., pp. 370–379.

(approximately 2%/year) than in the first post-transplant year [24, 27]. Those with elevated bone turnover markers lost more bone than those with normal or low markers [27]. Glucocorticoid withdrawal has been associated with significant increases in BMD and bone formation markers [28]. In contrast, other studies have not shown ongoing bone loss or have demonstrated site-specific increases and decreases [29–32]. Fracture prevalence (Table I) ranges from 7 to 11% in non-diabetic renal transplant recipients [33, 34], but is considerably higher in patients transplanted because of diabetic nephropathy and in those who receive kidney–pancreas transplants [34–37]. Fractures more commonly involve the long bones or metatarsals than the vertebral bodies or ribs [38] and occur relatively late in the post-transplant period, usually more than 3 years after transplantation. This may be due, in part, to the detrimental effects of pretransplant hyperparathyroidism or persistent post-transplant hyperparathyroidism on cortical bone, rendering it thin, porous, and vulnerable to further insult. A recent cohort study of 101039 patients with ESRD found that kidney transplantation was associated with a 34% greater risk of hip fracture than continued dialysis [39]. Markers of bone formation have been frequently assessed after renal transplantation. Julian et al. [17] observed that total serum alkaline phosphatase activity declined by 60% during the first 18 months after renal transplantation. In a cross-sectional study, Bagni and colleagues [20] measured serum osteocalcin in 44 renal transplant recipients 6–144 months after transplantation. Levels were markedly increased, on average, and directly correlated with intact PTH concentrations, which were also mildly elevated. Osteocalcin levels did not differ

between patients taking and not taking CsA. Renal function was mildly compromised (mean creatinine clearance, 66 ml/min and serum creatinine 1.5 mg/dl). Since serum osteocalcin levels have been shown to increase only after the creatinine clearance falls below 30 ml/min [40], it is unlikely that the increase in osteocalcin could be attributed to impaired clearance of this small molecule. Briner et al. [41] measured biochemical indices of mineral metabolism, including serum PTH, osteocalcin and urinary hydroxyproline excretion in 34 patients before and at 6 month intervals for 2 years after transplantation. They also performed bone biopsies in 20 patients, before and during the second post-transplant year. Serum total alkaline phosphatase activity declined in patients with elevated pretransplant levels and rose in those with normal or low pretransplant levels. Serum osteocalcin was markedly elevated before transplantation, fell by approximately 50% at 6 months and remained stable for the next 18 months. However, despite the marked decrease in serum osteocalcin after transplantation, levels remained almost twice the upper limit of normal throughout the 2 years of observation. PTH also remained elevated and correlated with serum osteocalcin and creatinine concentrations. Static parameters of bone formation and resorption were increased before and 1 year after transplantation. Unfortunately, no data were given regarding correlation of histologic and biochemical bone formation parameters. Approximately one-third of the patients were managed with CsA monotherapy. However, there was no comparison of biochemical or histomorphometric parameters between patients on CsA monotherapy and those who required prednisone in addition. In another cross-sectional study, Boiskin et al. measured serum alkaline phosphatase activity, osteocalcin, and PTH (in a midmolecule assay)

704 at various intervals and in varying numbers of patients before and during an 8-month period after transplantation [42]. All were observed to be markedly elevated before transplantation and all declined after transplantation. Serum alkaline phosphatase and osteocalcin fell to slightly above the upper end of the normal range and remained there. Values of these markers were directly correlated with each other and with PTH. Pietschmann et al. compared serum osteocalcin and intact PTH concentrations in 30 patients measured an average of 2 years after transplantation and 30 age- and sex-matched controls [43]. All patients were receiving prednisone (7.5 mg/day on average) and CsA. PTH and osteocalcin were significantly higher in the renal transplant recipients, even those with creatinine clearances above 95 ml/min. Again, serum osteocalcin correlated with total alkaline phosphatase activity and PTH, but bore no relationship to serum creatinine, prednisolone dose, or CsA levels. Bone-specific alkaline phosphatase (BSAP) is an enzyme located in the membrane of the osteoblastic cells and is considered to be a more sensitive and specific marker of bone formation than total alkaline phosphatase activity (see Chapter 9). BSAP is a large molecule and is not renally excreted. Thus when glomerular filtration is impaired, as it may frequently be after renal transplantation, serum BSAP may more closely reflect bone formation/ remodeling activity than serum osteocalcin, a small molecule that is cleared by the kidney. Withold et al. observed that total alkaline phosphatase and BSAP were normal and highly correlated prior to transplantation in 23 patients with ESRD [44]. Although there was a significant decrease in BSAP during the first week after transplantation, perhaps reflecting the high doses of glucocorticoids given during this period, the next 6 months were characterized by an approximate doubling of BSAP. Levels correlated highly with intact PTH both before and after transplantation. The authors concluded that BSAP is a useful marker of bone formation after renal transplantation. These same investigators also evaluated the carboxyterminal extension peptide of type 1 procollagen (PICP) in 13 renal transplant recipients [45]. The concentration of circulating PICP is thought to be directly associated with type 1 collagen synthesis and, hence, bone formation and PICP has been shown to correlate with histomorphometric parameters of bone formation in patients on dialysis with renal osteodystrophy [46, 47]. BSAP and PICP were highly correlated both before and after transplantation [45]. There was an approximate 50% increase in both BSAP and PICP by 3 months after transplantation. BSAP and PICP correlated with PTH before, but not after, transplantation. Gonzalez et al. compared osteocalcin, BSAP, and PICP in 23 patients after renal

Carolina A. Moreira Kulak and Elizabeth Shane

transplantation [48]. Despite a concomitant decline in intact PTH concentrations, there was a significant rise in all three markers of bone formation, which became apparent by the third post-transplant month, with a further increase by the sixth month. In general, glucocorticoids suppress bone formation, inhibiting the transformation of preosteoblasts to osteoblasts, the synthesis of type 1 collagen and osteocalcin (Chapter 45). Since virtually all of the studies summarized above have been conducted in patients who were taking glucocorticoids, the mechanism of the increase in the biochemical and histomorphometric parameters of bone formation is not clear. One possibility is that serum PTH concentrations, while usually lower than before transplantation, remain elevated in many renal transplant recipients for a significant period of time, sometimes years. PTH is a known stimulator of bone formation and could conceivably overcome the suppressive effects of glucocorticoids on bone formation. In this regard, intermittent injections of PTH significantly increase bone mass in patients on chronic glucocorticoid therapy [49]. In support of this theory, most of the studies reviewed report significant correlations between biochemical markers of bone formation and PTH. The degree of posttransplant renal impairment may influence the situation by delaying the resolution of hyperparathyroidism, hence keeping bone turnover high for longer. Also osteocalcin [40], although not BSAP or PICP excretion, may be reduced when renal function is impaired. Another possible reason for the increase in bone formation markers is the use of CsA, which has been shown to increase both bone formation and resorption when administered to rats [5–7]. Some data suggest that CsA may stimulate bone formation, an effect opposite to that usually observed with glucocorticoids. Raumbausek et al. reported that total alkaline phosphatase activity was significantly higher in renal transplant recipients managed with prednisone and CsA than those managed with prednisone and azathioprine [50]. In a histomorphometric study, Aubia and colleagues reported that indices of both bone resorption and formation were significantly higher in patients receiving CsA and prednisone than those managed with prednisone and azathioprine [51]. Wilmink et al. compared three groups of patients: one managed with CsA alone, the second with CsA and prednisone, and the third managed with azathioprine and prednisone [52]. Patients managed with CsA were significantly more likely to have an elevated alkaline phosphatase, and alkaline phosphatase activity decreased markedly after patients were switched from CsA to prednisone and azathioprine. However, Dumoulin et al. [53] compared two groups of long-term renal transplant recipients, one group receiving CsA and

Chapter 42 Transplantation Osteoporosis: Biochemical Correlates of Pathogenesis and Treatment

the other not. The groups were matched with respect to gender, age, time since transplantation (4–5 years), doses of other immunosuppressive agents, and renal function, which was well preserved. Intact PTH was at the upper end of the normal range. Osteocalcin levels, at the upper end of the normal range on average, did not differ. Finally, no study has yet reported any relationship between CsA level or dose and any marker of bone formation. Thus the balance of the evidence seems to implicate hyperparathyroidism rather than CsA as the cause of increased bone turnover in renal transplant recipients, as suggested by Amado et al. [54].

III. CARDIAC TRANSPLANTATION In contrast to end-stage renal disease, congestive heart failure (CHF) has not been associated with a well-defined disorder of bone and mineral metabolism. Mean bone mineral density (BMD) has been reported to be lower in patients before heart transplantation than in age- and sexmatched normal individuals in some [55–57] but not all studies [58]. A study of 101 patients with advanced CHF [57] found very low serum (<10 ng/ml) concentrations of 25-hydroxyvitamin D (25-OHD) in 18% of the patients. Moreover, vitamin D deficiency was significantly more common in the patients with more severe heart failure. Secondary hyperparathyroidism, related primarily to prerenal azotemia, was observed in 30% of the patients. Markers of bone resorption (hydroxyproline, pyridinoline and deoxypyridinoline; see Chapter 35) were elevated in the group as a whole and were significantly higher in patients with low 25-OHD and 1,25(OH)2D concentrations. Despite mild prerenal azotemia, the bone formation marker measured (serum osteocalcin) was normal in men and women. In contrast, Lee et al. found both serum PTH and osteocalcin concentrations to be elevated in a smaller group of patients with CHF [55]. Osteoporosis and fractures are common in cardiac transplant recipients. Cross-sectional studies have documented that vertebral fracture prevalence ranges from 22% to 50% [55–57, 59]. Longitudinal studies [58, 60–62] have reported that fracture incidence ranges between 18% and 36%. Most fractures involve the spine and occur during the first 6–12 months following transplantation. Although no pretransplant BMD or biochemical parameter has been shown to predict fracture after transplantation in the individual patient, lower pretransplant BMD (T score < −1.0) and female gender is associated with increased fracture risk. Several longitudinal studies describing the natural history and pattern of bone loss after cardiac transplantation

705

have been published, virtually all of them during the early to mid 1990s [61, 63–66]. In these studies, lumbar spine BMD fell by approximately 6–10% during the first 6 months after transplantation, after which there was little further deterioration [63]. In contrast, hip BMD declined throughout the first year, reaching 10–15% below pretransplant levels [63]. Bone loss slows or stops in the majority of patients after the first year and lumbar spine BMD has been shown to increase slightly in the third year [63, 66]. A more recent study found that rates of bone loss at the spine and hip in subjects who did not receive specific preventive therapy are approximately 50% less than observed previously [58]. Biochemical changes after cardiac transplantation include sustained decreases in serum 1,25(OH)2D and transient decreases in markers of bone formation (osteocalcin) and testosterone (in men), with return to pretransplant concentrations by 6 months [63, 67]. Higher rates of bone loss were associated with greater exposure to prednisone, lower serum concentrations of vitamin D metabolites, higher levels of bone resorption markers [59], and in men, lower serum testosterone concentrations [63, 67]. Mean intact PTH concentrations were at the upper end of the normal range before transplantation and remained there throughout the 3 years of observation [63]. Sambrook et al. reported that serum osteocalcin, suppressed immediately after transplantation, increased markedly over the ensuing year [67]. They observed that the change in osteocalcin correlated significantly with the daily doses of CsA and prednisolone and that the 6-month osteocalcin level correlated inversely with lumbar spine bone loss. Thiebaud et al. observed progressive increases in serum osteocalcin and intact PTH concentrations during 18 months following cardiac transplantation [68], a pattern that differs both from our observations [63] and the suppression of osteocalcin concentrations observed in patients on glucocorticoids. Osteocalcin concentrations were directly correlated with PTH, but unrelated to serum creatinine, doses of CsA, or glucocorticoids [68]. Urinary hydroxyproline excretion was elevated at all points after transplantation. Since many of these patients were withdrawn from glucocorticoids and maintained on CsA alone, this observation may reflect either an independent effect of CsA to increase bone formation or simply the withdrawal of steroids. Markers of bone resorption have been shown to increase transiently after cardiac transplantation [63, 67], a pattern which also differs from that reported in patients on glucocorticoids alone [69, 70]. In a cross-sectional study of 50 male cardiac transplant recipients [71], evaluated between 2 weeks and 47 months after transplantation, BMD was significantly decreased and markers of both formation and resorption were

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significantly increased. Bone turnover markers correlated with intact PTH, which in turn was related to the degree of renal impairment.

IV. LIVER TRANSPLANTATION Candidates for liver transplantation frequently have significant skeletal demineralization and fractures, which place them at increased risk for fracture after surgery. A study of 58 patients with cirrhotic end-stage liver disease referred for liver transplantation [72] found that 43% had osteoporosis (defined as a Z score < −2.0 or prevalent vertebral fractures). Serum 25-OHD, 1,25(OH)2D, intact PTH, and osteocalcin were lower and urinary hydroxyproline excretion was higher in cirrhotic patients than controls and male patients had lower serum testosterone levels than controls. Ninkovic et al. also noted a high prevalence of low BMD, which was associated with older age, lower body weight, and the presence of cholestatic liver disease [73]. Other investigators have observed low serum osteocalcin concentrations in patients with primary biliary cirrhosis [74, 75] and histomorphometric data have confirmed that bone formation is decreased in patients with primary biliary cirrhosis [76–78]. While the pathogenesis of the decrease in bone formation is not known, unconjugated hyperbilirubinemia is associated with reduced osteoblast activity in jaundiced patients, reversible after selective removal of bilirubin from plasma by photobleaching [79]. While serum osteocalcin appears to be a valid marker of bone formation in cholestatic liver disease, the utility of collagen-related markers of bone turnover has been called into question [75]. In fibrotic liver diseases, the synthesis of type I collagen is markedly increased. Guanabens et al. recently evaluated collagen-related markers of bone formation (osteocalcin, carboxyterminal and aminoterminal propeptides of type I collagen) and resorption (serum tartrate-resistant acid phosphatase, cross-linked carboxyterminal telopeptide of type I collagen, and urinary excretion of pyridinium crosslinks, hydroxyproline, N- and C-telopeptides) in 34 women with primary biliary cirrhosis [75]. Of these, serum osteocalcin was significantly lower than, and tartrate-resistant acid phosphatase (TRAP) did not differ from, controls. In contrast, all of the collagen-related markers of bone turnover were significantly higher in patients than controls and all correlated with indices of liver fibrogenesis and disease severity rather than with osteocalcin and TRAP. Thus, these collagen-related markers appear influenced by liver, rather than bone, collagen metabolism. The natural history of osteoporosis after liver transplantation is similar to that observed after cardiac

transplantation [80–85]. Lumbar spine BMD falls by 3.5–24%, primarily during the first 3–6 months, with gradual improvement thereafter. Fracture incidence is also highest in the first year [60]. In older studies, women with primary biliary cirrhosis appeared to be at greatest risk [84]. The vertebrae and ribs are the most common fracture site. Type of liver disease, glucocorticoid exposure, and markers of bone turnover do not reliably predict fracture risk. However, retransplanted patients, those with primary biliary cirrhosis and those with previous fragility fractures, are at increased risk [60, 86]. Older age and pretransplant BMD predicted post-transplantation fractures in one prospective study [60]. More recent studies suggest that rates of bone loss (particularly at the spine) and fracture are considerably lower now than reported in the early 1990s [87]. Studies of calciotropic hormone levels and bone turnover markers after liver transplantation are limited. Compston et al. reported a significant increase in serumintact PTH during the first 3 months after liver transplantation [88]. Others have found intact PTH levels to be within the normal range in liver transplant recipients [85, 89, 90]. In a cross-sectional study, markers of bone formation (osteocalcin and carboxyterminal peptide of type I collagen) were significantly higher in 120 liver transplant recipients than in a normal control population [90]. Similarly, serum osteocalcin was higher than age- and sex-matched controls in 18 men and nine postmenopausal women after liver transplantation and urinary hydroxyproline excretion was also elevated [85]. Significant increases in serum osteocalcin were not observed in one study in which levels were obtained before and 1 month after transplantation [91]. However, in studies of longer duration, serum osteocalcin levels were found to increase substantially during the first posttransplant year [92, 93]. The balance of the data thus suggests that low bone turnover observed in many patients with liver failure converts to a high turnover state that persists indefinitely after liver transplantation. The biochemical change from low to high bone turnover has been confirmed by quantitative histomorphometry [78]. The cause may be related to resolution of cholestasis and/or hypogonadism, increased PTH secretion, calcineurin inhibitors (cyclosporine A or tacrolimus), or the development of renal insufficiency.

V. LUNG TRANSPLANTATION Patients who undergo lung transplantation are highly likely to have osteoporosis even before surgery [94–96]. Decreased mobility, hypoxemia, malnutrition, vitamin D

Chapter 42 Transplantation Osteoporosis: Biochemical Correlates of Pathogenesis and Treatment

deficiency, tobacco use, low body weight, and prior glucocorticoid therapy are frequent attributes of candidates for lung transplantation and all of these factors may contribute to pretransplant osteopenia. Cystic fibrosis, a common disease for which patients undergo lung transplantation, is itself associated with osteoporosis and fractures [97–100] due to pancreatic insufficiency, vitamin D deficiency and calcium malabsorption and hypogonadism. One cross-sectional study revealed that 45% of 55 pretransplant patients and 73% of 45 post-transplant patients had BMD > 2 SD below age- and sex-matched controls [94]. Another study of 70 patients awaiting lung transplantation revealed that only 34% had normal spine BMD and 22% had normal hip BMD [95]. Fracture prevalence was 29% in patients with chronic obstructive pulmonary disease and 25% in patients with cystic fibrosis [95]. In both studies, glucocorticoid exposure was inversely related to BMD. Serum 25-hydroxyvitamin D levels were at the low end of the then normal range and 20% of the patients had markedly deficient values, particularly by today’s standards [95]. Several small studies have prospectively evaluated changes in bone mass and fracture incidence in patients who have received a lung transplant [101–105]. A study of 12 patients who underwent lung transplantation demonstrated an average 4% decrease in lumbar spine BMD during the first 6 months after transplantation, despite supplementation with calcium and 400 IU of vitamin D [102]. Eight sustained significant lumbar spine bone loss, seven sustained significant femoral neck bone loss and two men sustained multiple vertebral fractures. On average, BMD at the femoral neck, femoral shaft, and whole body did not change significantly. We have also prospectively followed spine and hip BMD and fracture incidence in a group of 30 lung-transplant recipients [103]. Despite the fact that all received antiresorptive therapy initiated shortly after transplantation, 37% of the patients suffered fragility fractures during the first post-transplant year and approximately 50% sustained significant bone loss at either the spine or the hip. Risk factors for fracture included low pretransplant lumbar spine bone mass and pretransplant glucocorticoid therapy. Biochemical indices of mineral metabolism (serum and urinary calcium, intact PTH, vitamin D metabolites) were similar in patients who did and did not fracture. However, bone turnover markers (both resorption and formation) were higher in fracture patients including 6-month serum bone alkaline phosphatase and 12-month urinary pyridinoline and deoxypyridinoline excretion. The 15 patients with significant bone loss at one or both sites did not differ from those with stable BMD with respect to any pretransplant demographic, densitometric or biochemical

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parameter, exposure to glucocorticoids, or tobacco. Nor did they differ with respect to post-transplant exposure to glucocorticoids, number of rejection episodes or type of osteoporosis therapy. At 12 months after transplantation, serum calcium was lower, serum PTH tended to be higher and urinary deoxypyridinoline excretion was significantly higher. Thus, increased bone resorption, possibly due to secondary hyperparathyroidism, may have contributed to bone loss in these patients. The pathogenesis of the secondary hyperparathyroidism may have been related to urinary calcium losses, which were greater in patients with significant bone loss and were directly related to intact PTH concentrations. In contrast, Spira et al. did find a significant relationship between bone loss and posttransplant glucocorticoid exposure [104].

VI. BONE MARROW TRANSPLANTATION Patients receiving allogeneic bone marrow transplantation (BMT) for various hematological conditions are exposed to many factors that may influence bone mass and bone metabolism. Induction and consolidation regimens (which may involve chemotherapeutic agents, glucocorticoids, and/or total body irradiation), hypogonadism, and immobility may all contribute. In patients studied prior to transplantation (after chemotherapy), normal bone density at the LS and FN has been documented in 72%, osteopenia in 24%, and osteoporosis in only 4% [106]. Over the first year after BMT, studies have found rates of bone loss of 2–9% at the LS and 6–11% at the FN [106–111]. Recovery of LS BMD after the first 6–12 months has also been documented [106, 110]. An uncoupling of bone turnover, with increased markers of resorption and decreased markers of formation, has been shown in the weeks just prior to and after transplantation [108–112]. Studies of bone marrow stromal cells suggest that osteoblastic differentiation may be affected by the BMT medication regimen, resulting in decreased bone formation [111, 113]. Bone loss has also been correlated to steroid exposure [108, 111] and amenorrhea (in the female patients) [108] in prospective studies. Vertebral and non-vertebral fractures incidence has ranged from 1–16% [106–108, 110].

VII. MECHANISMS OF BONE LOSS AFTER TRANSPLANTATION In the previous sections, we have summarized the now considerable body of research published over the past

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decade into the natural history and pathogenesis of bone loss and fracture after solid organ transplantation. This accumulating knowledge base has yielded fairly consistent data that now enable us to develop a unifying hypothesis of the mechanisms of post-transplantation bone loss. It seems very clear that the mechanisms differ according to the amount of time that has elapsed since transplantation. There appear to be two main phases of bone loss (Fig. 1). These phases can best be differentiated from each other by the presence (Fig. 1A) or absence (Fig. 1B) of high-dose corticosteroids in the immunosuppressive regimen.

During the first 6 months after transplantation, steroid doses are generally high enough to profoundly suppress bone formation by virtue of their effects to reduce osteoblast numbers, increase osteoblast apoptosis, and inhibit osteoblast synthetic function. Virtually every published study has found serum markers of bone formation, particularly serum osteocalcin, to be suppressed during this period. During this same period, there has also been consistency in reports of increased urinary markers of bone resorption. The pathogenesis of the increase in resorption markers is probably in part related to suppressive effects of glucocorticoids on osteoblast synthesis of OPG, on the

A

B

Figure 1

(A) Mechanisms of bone loss in the early post-transplantation period. (B) Mechaninsms of bone loss in the later post-transplantation period, after glucocorticoid doses are tapered.

Chapter 42 Transplantation Osteoporosis: Biochemical Correlates of Pathogenesis and Treatment

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hypothalamic-pituitary-gonadal axis and on calcium transport across the intestinal, renal tubular, and parathyroid cell membranes. In addition, the well-known nephrotoxic effects of CsA and tacrolimus administration result in measurable declines in renal function and decreased synthesis of 1,25(OH)2D that also inhibits calcium transport in the gut. Thus both calcineurin inhibitors and glucocorticoids have the potential to cause secondary increases in parathyroid hormone secretion, which in turn increases osteoclast-mediated bone resorption. In addition, there may be direct effects of both CsA and FK506 to increase bone resorption. The concomitant administration of high-dose glucocorticoids and cyclosporine (or FK506) is therefore associated with uncoupling of resorption and formation. During this phase of the posttransplant period, there is rapid bone loss and high fracture rates. As steroid doses are tapered to below 5 mg per day, osteoblast function recovers and the suppressive effects on bone formation are reversed. However, adverse effects of CsA and FK506 remain – both the direct effects on the skeleton and those indirect effects mediated by renal toxicity of these drugs that result in secondary hyperparathyroidism. Thus, resorption remains elevated. With the tapering of steroids, bone formation increases also, resulting in “recoupling” of bone turnover. Rates of bone loss slow and there may even be some recovery, particularly at the spine. Each time steroid doses are increased for treatment of rejection or stress, the pathophysiologic picture again resembles the first phase.

patients with end-stage renal disease. All patients should receive the recommended daily allowance for calcium (1000–1500 mg/day) and vitamin D (400–800 IU/day). Finally, specific attention should be given to the subsets of patients who are at higher risk for fractures after transplantation. Lower BMD before transplantation is not a consistent risk factor for fractures after transplantation. Moreover, it is unclear whether treatment of osteoporosis present before transplantation decreases the incidence of fractures after transplantation, as controlled studies regarding this issue have not been conducted. However, antiresorptive agents may help improve BMD in liver, heart, and lung [114] transplant candidates and reduce fractures in other populations and thus their use can be supported on these bases. In our view, therefore, BMD should be measured to detect osteoporosis that exists before the transplant, and can be helpful in selecting patients who would benefit from immediate initiation of antiresorptive or anabolic therapy. In patients who are found to have normal BMD, such therapy could be safely deferred until the time of transplantation. Interpretation of BMD measurements is difficult in the setting of renal bone disease, and the World Health Organization criteria for diagnosis of osteoporosis should not be used in these patients. Bisphosphonates may increase the risk for adynamic bone disease in ESRD patients and are not approved for the treatment of these patients [115].

VIII. PREVENTION AND MANAGEMENT OF TRANSPLANTATION OSTEOPOROSIS

Since rates of bone loss and fracture incidence are highest immediately following transplantation, preventive and therapeutic measures should be instituted at that time and without delay. In our opinion, the lack of reliable clinical predictors to identify individual patients who will experience osteoporotic fractures renders all transplant recipients candidates for preventive therapy. Most prevention trials in post-transplantation osteoporosis are limited by small sample size, the absence of fracture data, the lack of randomization, and conduct in single centers which limits generalizability.

A. Pretransplantation Measures All transplant candidates should be evaluated and treated prior to transplantation, as bone disease is common in patients awaiting organ transplantation. Since waiting periods on the transplant list can be as long as 2 years, there is often time to implement measures to improve skeletal health. Lifestyle factors such as immobilization, smoking, and alcohol abuse should be addressed. The use of medications that can negatively impact skeletal health (e.g. anticonvulsants, glucocorticoids, heparin, frusemide) should be assessed and minimized to the extent possible. Other factors that can be corrected include hypogonadism and negative calcium balance (due to malabsorption and/or vitamin D deficiency) leading to secondary hyperparathyroidism. Evaluation and treatment of renal osteodystrophy according to accepted guidelines is recommended for all

B. Prevention of Early Post-transplantation Bone Loss

1. Exercise

The importance of physical activity in restoring BMD was demonstrated in three prospective, randomized, albeit very small, studies conducted in heart [116, 117] and lung [118] transplant recipients. The type of training consisted of supervised exercises of the lumbar extensor muscles (1 day/week) and upper and lower body (2 days/week), started 2 months following transplantation

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and continued for 6 months. This resistance training restored BMD toward pretransplantation levels more rapidly [116, 118], and in conjunction with alendronate, was more efficacious than alendronate alone [117]. 2. Calcium and Vitamin D

Replacement doses of calcium and vitamin D (up to 1000 IU) fail to prevent clinically significant posttransplantation bone loss, as demonstrated by numerous observational studies and the control arms of several trials studying the effects of different medications on the prevention of transplantation osteoporosis. Nevertheless, it is our opinion that all transplant recipients should receive 1000 mg elemental calcium and at least 400 IU of vitamin D daily. 3. Calcitonin

Calcitonin is relatively ineffective in preventing bone loss after transplantation [119–121], although it is a safe alternative if other agents are contraindicated or poorly tolerated. 4. Active Vitamin D Metabolites

Calcium and calcidiol (25-OHD) therapy was associated with an increase in BMD levels or prevention of further bone loss in heart transplant recipients [122] and renal transplant recipients [123]. The efficacy of calcitriol is controversial. At doses averaging 0.25 µg daily, calcitriol did not significantly prevent lumbar spine bone loss in renal [124] and heart [121] transplant recipients. Calcitriol therapy at doses greater than 0.5 µg daily may be more effective. In a 2-year randomized double-blind study, calcitriol (0.5–0.75 µg daily) reduced proximal femur bone loss in heart and lung transplant recipients, and reduced the occurrence of vertebral fractures/deformities, although not significantly [125]. However, there was no reduction of lumbar spine bone loss. In another study, calcitriol (0.5 µg daily) reduced bone loss at the femoral neck and lumbar spine in a group of 75 heart transplant recipients compared to an untreated reference group [58]. At doses of 0.5 µg or higher, common side effects of calcitriol therapy include hypercalcemia and hypercalciuria (seen in over 50% of patients), which may develop at any time during therapy, and which require close monitoring [58, 125]. 5. Bisphosphonates

Bisphosphonates are effective for the prevention of glucocorticoid-induced osteoporosis, and their antiresorptive mechanism of action makes these drugs an obvious option in preventing the phase of rapid bone loss early after transplantation, which is associated with increased

bone resorption. Several clinical trials have confirmed this benefit. Cyclical etidronate has been studied extensively, but has fallen out of favor since the introduction of newer, more potent bisphosphonates. Compared to placebo, two intravenous infusions of pamidronate (0.5 mg/kg), given at the time of renal transplantation and 1 month later, prevented bone loss at the femoral neck and lumbar spine at 1 year [126], and the protection conferred at the hip was maintained 4 years after transplantation [127]. Similar results have been reported with repeated intravenous doses of pamidronate in a controlled randomized study of 34 patients with cystic fibrosis [128], and in non-randomized studies of heart [129, 130], lung [105], and liver transplant recipients [131]. However, in a recent study of liver transplant recipients, a single dose of 60 mg IV pamidronate had no significant effect on fracture rate or BMD change post-transplantation compared to placebo [87], possibly because pamidronate was administered as much as 3 months before transplantation in some patients. In a study of renal transplant recipients, repeated doses of intravenous pamidronate preserved vertebral BMD during treatment and 6 months after cessation of treatment. However, pamidronate treatment was associated with development of adynamic bone histology, and no significant difference in the incidence of fractures was noted between the two groups in the study [132]. Alendronate use (10 mg daily) significantly reduced bone loss at the femoral neck and lumbar spine in heart transplant recipients compared to an untreated reference group. However, alendronate was not significantly different from calcitriol (0.5 µg daily) [58]. In a non-randomized study of renal transplant recipients, alendronate reduced bone loss [133]. Two 4 mg IV doses of zoledronic acid given to 20 patients at 2 weeks and 3 months after renal transplantation led to higher bone mineral density values than placebo at 6 months but not at 3 years [134]. Intravenous ibandronate reduced bone loss in liver [135] and kidney transplant recipients [136] compared to calcium supplementation alone. In summary, oral and intravenous bisphosphonates, in conjunction with calcium and vitamin D, are effective in preventing post-transplantation bone loss when started shortly after grafting. The optimal dose, timing, and frequency, particularly of intravenous bisphosphonate administration, remain to be determined. No reports to date demonstrate unequivocal protection from fractures. Their use after pediatric transplantation and in patients with poor renal function should be considered carefully. Finally, the risk of inducing or prolonging adynamic bone disease in renal transplant recipients, especially with repeated dosing, must be kept in mind. Despite these concerns, however,

Chapter 42 Transplantation Osteoporosis: Biochemical Correlates of Pathogenesis and Treatment

we recommend their use after renal transplantation, at least during the first year when rates of bone loss are most rapid.

C. Treatment of Bone Loss in Long-Term Transplant Recipients Many transplant recipients have established osteoporosis and/or persistent ongoing bone loss. Several studies have examined treatment options for these patients.

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divided into two groups based on BMD and markers of bone turnover [27]. The group with normal markers of bone turnover was followed untreated, and did not lose any further bone mass at the lumbar spine or femoral neck at one year. On the other hand, in the group with high bone turnover and osteoporosis or osteopenia, alendronate therapy (10 mg daily) decreased the rate of bone loss and increased bone mass over a year of treatment [27]. Although uncontrolled, this study may be helpful in guiding the therapy of long-term transplant recipients, and in identifying those who may benefit from bisphosphonate therapy.

1. Vitamin D Metabolites

Calcitirol (0.5 µg daily) has been reported to be superior to calcium alone in preserving lumbar spine bone mass in renal transplant recipients who were started on treatment approximately 3 years after grafting [137]. In contrast, a lower dose of calcitriol (0.25 µg/day) did not significantly increase BMD in renal transplant recipients, who were on average 10 years post-transplant [32]. Similarly, lumbar spine BMD increased similarly in calcitriol- and calcium-treated cardiac transplant recipients enrolled at 6 months [138] or 35 months [139] after grafting, although the interpretation of these two studies is confounded by the use of concomitant hormone therapy replacement in hypogonadal patients. 2. Bisphosphonates

Pamidronate (30 mg IV every 3 months for 2 years) was studied in 13 cardiac and 21 liver transplant recipients with osteoporosis, approximately 2 years after transplantation. Significant gains in lumbar spine and femoral neck BMD were noted compared to historical controls [140]. Therapy with 1 year of oral clodronate (1600 mg daily), initiated 6 months after transplantation in 64 cardiac transplant recipients with low BMD, induced a significant increase in lumbar spine BMD at 1 year [141]. In renal transplant recipients with low bone mass, cyclical clodronate therapy (800 mg daily) did not result in significant changes in spine BMD compared to calcium alone [142]. Alendronate (10 mg daily), started approximately 5 years after renal transplantation, resulted in significant gains in BMD; again the number of patients included was too small to demonstrate fracture prevention [143]. In two other studies of long-term renal transplant recipients, alendronate resulted in significant increases in femoral neck and lumbar spine BMD [144, 145] . In neither did the increase in BMD differ between the alendronate and calcitriol groups. In another study of 58 long-term kidney transplant recipients more than 1 year post-grafting, patients were

IX. CONCLUSIONS Candidates for all types of transplantation have significant risk factors for osteoporosis and many have low bone mass, prevalent fractures or evidence of abnormal mineral metabolism before transplantation. In the majority of cases, markers of both bone formation and resorption are increased before transplantation. The exception to this is cholestatic liver disease in which bone turnover is low before transplantation. After transplantation, there is rapid bone loss in the majority of patients and fractures also occur commonly. These complications occur with the greatest frequency during the first 6–12 months. Early after transplantation, when glucocorticoid doses are highest, serum osteocalcin and bone alkaline phosphatase are suppressed and markers of resorption are elevated, biochemical evidence of uncoupled formation and resorption. However, by the middle of the first year after transplantation, bone formation markers recover. The balance of the evidence in organ transplant recipients evaluated after the first 6 months favors the notion that transplantation-related bone loss is a form of high turnover osteoporosis. As such, antiresorptive therapy is likely to have benefit in the prevention of bone loss and fractures after transplantation.

Acknowledgments This work was supported in part by Grants AR-41391 and RR-006645 from the National Institutes of Health.

References 1. United Network of Organ Sharing (UNOS). www.unos.org. 2. Cohen, A., and Shane, E. (2001). Transplantation osteoporosis. Curr. Opin. Endocrinol. Diab. 8, 283–290.

712 3. Cohen, A., and Shane, E. (2003). Osteoporosis after solid organ and bone marrow transplantation. Osteoporos. Int. 14, 617–630. 4. Cohen, A., Ebeling, P., Sprague, S., and Shane, E. (2003). Transplantation osteoporosis. In: “Primer on the Metabolic Bone Diseases and Disorders of Mineral Metabolism”. (M. J. Favus, ed.), pp. 370–379. American Society for Bone and Mineral Research, Washington, D.C. 5. Movsowitz, C., Epstein, S., Fallon, M., Ismail, F., and Thomas, S. (1988). Cyclosporin A in vivo produces severe osteopenia in the rat: Effect of dose and duration of administration. Endocrinol. 123, 2571–2577. 6. Movsowitz, C., Epstein, S., Ismail, F., Fallon, M., and Thomas, S. (1989). Cyclosporin A in the oophorectomized rat: Unexpected severe bone resorption. J. Bone Miner. Res. 4, 393–398. 7. Schlosberg, M., Movsowitz, C., Epstein, S., Ismail, F., Fallon, M. D., and Thomas, S. (1989). The effects of cyclosporin A administration and its withdrawal on bone mineral metabolism in the rat. Endocrinol. 124, 2179–2184. 8. Stein, B., Halloran, P., Reinhardt, T., Engstrom, G. W., Bales, C. W., Drezner, M. K., Currie, K. L., Takizawa, M., Adams, J. S., and Epstein, S. (1991). Cyclosporin A increases synthesis of 1,25 dihydroxyvitamin D3 in the rat and mouse. Endocrinol. 128, 1369–1373. 9. Stein, B., Takizawa, M., Katz, I., Berlin, J., Joffe, I., Fallon, M., and Epstein, S. (1991). Salmon calcitonin prevents cyclosporin A induced high turnover bone loss. Endocrinol. 129, 92–98. 10. Sass, D. A., Bowman, A. R., Marshall, I., Yuan, Z., Ma, Y. F., Jee, W. S. S., and Epstein, S. (1997). Alendronate prevents cyclosporin-induced osteopenia in the rat. Bone 21, 65–70. 11. Cvetkovic, M., Mann, G. N., Romero, D. F., Liang, X., Ma, F., Jee, W. S. S., and Epstein, S. (1994). The deleterious effects of long term cyclosporin A, cyclosporin G and FK506 on bone mineral metabolism in vivo. Transplantation 57, 1231–1237. 12. Goodman, W. G., Coburn, J. W., Slatopolsky, E., Salusky, I. B., and Quarles, L. D. (2003). Renal osteodystrophy in adults and children. In: “Primer on the Metabolic Bone Diseases and Disorders of Mineral Metabolism”. (M. Favus, ed), pp. 430–447. American Society for Bone and Mineral Research, Washington, DC. 13. Schwarz, C., Bohmig, G. A., Steininer, R., Mayer, G., and Oberbauer, R. (2001). Impaired phosphate handling of renal allografts is aggravated under rapamycin-based immunosuppression. Nephrol. Dial. Transplant. 16, 378–382. 14. Green, J., Debby, H., Lederer, E., Levi, M., Zajicek, H. K., and Bick, T. (2001). Evidence for a PTH-independent humoral mechanism in post-transplant hypophosphatemia and phosphaturia. Kidney Int. 60, 1182–1196. 15. Caravaca, F., Fernandez, M. A., Ruiz-Calero, R., Cubero, J., Aparicio, A., Jimenez, F., and Garcia, M. C. (1998). Effects of oral phosphorus supplementation on mineral metabolism of renal transplant recipients. Nephrol. Dial. Transplant. 13, 2605–2611. 16. Almond, M. K., Kwan, J. T. C., Evans, K., and Cunningham, J. (1994). Loss of regional bone mineral density in the first 12 months following renal transplantation. Nephron. 66, 52–57. 17. Julian, B. A., Laskow, D. A., Dubovsky, J., Dubovsky, E. V., Curtis, J. J., and Quarrles, L. D. (1991). Rapid loss of vertebral bone density after renal transplantation. N. Engl. J. Med. 325, 544–550. 18. Kwan, J. T. C., Almond, M. K., Evans, K., and Cunningham, J. (1992). Changes in total body bone mineral content and regional bone mineral density in renal patients following renal transplantation. Miner. Electrolyte Metab. 18, 166–168. 19. Horber, F. F., Casez, J. P., Steiger, U., Czerniack, A., Montandon, A., and Jaeger, P. H. (1994). Changes in bone mass early after kidney transplantation. J. Bone Miner. Res. 9, 1–9. 20. Bagni, B., Gilli, P., Cavallini, A., Bagni, I., Marzola, M. C., Orzincolo, C., and Wahner, H. W. (1994). Continuing loss of vertebral mineral density in renal transplant recipients. Eur. J. Nucl. Med. 21, 108–112.

Carolina A. Moreira Kulak and Elizabeth Shane 21. Massari, P. U., Garay, G., and Ulla, M. R. (1994). Bone mineral content in cyclosporine-treated renal transplant patients. Transplant Proc. 26, 2646–2648. 22. Kalef-Ezra, J. A., Karantanas, A. H., Hatzikonstantnou, I., Sferopoulos, G., Glaros, D. K., and Siampoulos, K. C. (1994). Bone mineral status after renal transplantation. Invest. Radiol. 29, 127–133. 23. Grotz, W. H., Mundinger, F. A., Gugel, B., Exner, V. M., Kirste, G., and Schollmeyer, P. J. (1995). Bone mineral density after kidney transplantation. Transplantation 59, 982–986. 24. Pichette, V., Bonnardeaux, A., Prudhomme, L., Gagne, M., Cardinal, J., and Ouimet, D. (1996). Long-term bone loss in kidney transplant recipients: A cross-sectional and longitudinal study. Am. J. Kidney Dis. 28, 105–114. 25. Boot, A. M., Nauta, J., Hokken-Koelega, A. C. S., Pols, H. A. P., Ridder, M. A. J., and Keizer-Schrama, S. M. P. F. (1995). Renal transplantation and osteoporosis. Arch. Dis. Child 72, 502–506. 26. Cohen, A., and Shane, E. (2003). Osteoporosis after solid organ and bone marrow transplantation. Osteoporos. Int. 14, 617–630. 27. Cruz, D. N., Wysolmerski, J. J., Brickel, H. M., Gundberg, C. G., Simpson, C. A., Mitnick, M. A., Kliger, A. S., Lorber, M. I., Basadonna, G. P., Friedman, A. L., Insogna, K. L., and Bia, M. J. (2001). Parameters of high bone-turnover predict bone loss in renal transplant patients: a longitudinal study. Transplantation 72, 83–88. 28. Farmer, C. K. T., Hampson, G., Vaja, S., Abbs, I. C., Hilton, R. M., Koffman, G., Watkins, J., Sacks, S. H., and Fogelman, I. (2002). Late low dose steroid withdrawal in renal transplant recipients increases bone formation and bone mineral density without altering renal function: a randomized controlled trial. J. Bone Miner. Res. 17, S158. 29. Hurst, G., Alloway, R., Hathaway, D., Somerville, T., Hughes, T., and Gaber, A. (1998). Stabilization of bone mass after renal transplant with preemptive care. Transplant Proc. 30, 1327–1328. 30. Moreno, A., Torregrosa, J. V., Pons, F., Campistol, J. M., Martinez de Osaba, M. J., and Oppenheimer, F. (1999). Bone mineral density after renal transplantation: long-term follow-up. Transplant Proc. 31, 2322–2323. 31. Nowacka-Cieciura, E., Durlik, M., Cieciura, T., Lewandowska, D., Baczkowska, T., Kukula, K., Lao, M., Szmidt, J., and Rowinski, W. (2002). Steroid withdrawal after renal transplantation – risks and benefits. Transplant Proc. 34, 560–563. 32. Cueto-Manzano, A. M., Konel, S., Freemont, A. J., Adams, J. E., Mawer, B., Gokal, R., and Hutchison, A. J. (2000). Effect of 1,25dihydroxyvitamin D3 and calcium carbonate on bone loss associated with long-term renal transplantation. Am. J. Kid. Dis. 35, 227–236. 33. Grotz, W. H., Mundinger, A., Gugel, B., Exner, V., Kirste, G., and Schollmeyer, P. J. (1994). Bone fracture and osteodensitometry with dual energy x-ray absorptiometry in kidney transplant recipients. Transplantation 58, 912–915. 34. Nisbeth, U., Lindh, E., Ljunghall, S., Backman, U., and Fellstrom, B. (1994). Fracture frequency after kidney transplantation. Transplant Proc. 26, 1764. 35. Nisbeth, U., Lindh, E., Ljunghall, S., Backman, U., and Fellstrom, B. (1999). Increased fracture rate in diabetes mellitus and females after renal transplantation. Transplantation 67, 1218–1222. 36. Smets, Y. F., van der Pijl, J. W., de Fijter, J. W., Ringers, J., Lemkes, H. H., and Hamdy, N. A. (1998). Low bone mass and high incidence of fractures after successful simultaneous pancreas-kidney transplantation. Nephrol. Dial. Transplant. 13, 1250–1255. 37. Chiu, M. Y., Sprague, S. M., Bruce, D. S., Woodle, E. S., Thistlethwaite, J. R., Jr., and Josephson, M. A. (1998). Analysis of fracture prevalence in kidney-pancreas allograft recipients. J. Am. Soc. Nephrol. 9, 677–683. 38. Ramsey-Goldman, R., Dunn, J. E., Dunlop, D. D., Stuart, F. P., Abecassis, M. M., Kaufman, D. B., Langman, C. B., Salinger, M. H., and Sprague, S. M. (1999). Increased risk of fracture in patients receiving solid organ transplants. J. Bone Miner. Res. 14, 456–463.

Chapter 42 Transplantation Osteoporosis: Biochemical Correlates of Pathogenesis and Treatment 39. Ball, A. M., Gillen, D. L., Sherrard, D., Weiss, N. S., Emerson, S. S., Seliger, S. L., Kestenbaum, B. R., and Stehman-Breen, C. (2002). Risk of hip fracture among dialysis and renal transplant recipients. JAMA 288, 3014–3018. 40. Delmas, P. D., Wilson, D. M., Mann, K. G., and Riggs, B. L. (1983). Effect of renal function on plasma levels of bone Gla-protein. J. Clin. Endocrinol. Metab. 57, 1028–1030. 41. Briner, V. A., Thiel, G., Monier-Faugere, M.-C., Bognar, B., Landmann, J., Kamber, V., and Malluche, H. (1995). Prevention of cancellous of bone loss but persistence of renal bone disease despite normal 1,25 vitamin D levels two years after kidney transplantation. Transplantation 59, 1393–1400. 42. Boiskin, I., Epstein, S., Ismail, F., Thomas, S. B., and Raja, R. (1989). Serum osteocalcin and bone mineral metabolism following successful renal transplantation. Clin. Nephrol. 31, 316–322. 43. Pietschmann, P., Vychtil, A., Woloszczuk, W., and Kovaril, J. (1991). Bone metabolism in patients with functioning kidney grafts: increased serum levels of osteocalcin and parathyroid hormone despite normalization of kidney function. Nephron. 59, 533–536. 44. Withold, W., Degenhard, S., Castelli, D., Heins, M., and Grabensee, B. (1994). Monitoring of osteoblast activity with an immunoradiometric assay for determination of bone alkaline phosphatase mass concentration in patients receiving renal transplants. Clin. Chim. Acta. 225, 137–146. 45. Withold, W., Degenhard, S., Grabensee, B., and Reinauer, H. (1995). Comparison between serumlevels of bone alkaline phosphatase and the carboxy-terminal propeptide of type I procollagen as markers of bone formation in patients following renal transplantation. Clin. Chim. Acta. 239, 143–151. 46. Hamdy, N.A.T., Risteli, J., Risteli, L., Harris, S., Beneton, M. N., Brown, C. B., and Kanis, J. A. (1994). Serum type I procollagen peptide: a non-invasive index of bone formation in patients on haemodialysis? Nephrol. Dial. Transplant 9, 511–516. 47. Coen, G., Mazzafero, S., Ballanti, P., Bonucci, E., Bondatti, F., Manni, M., Pasquali, M., Perruzza, I., Sardella, D., and Spurio, A. (1992). Procollagen type I C-terminal extension peptide in predialysis chronic renal failure. Am. J. Nephrol. 12, 242–251. 48. Gonzalez, M. T., Bonnin, R., Cruzado, J. M., Garcia, R., Moreso, F., Fulladosa, X., Alsina, J. A., Navarro, M., and Grino, J. M. (1995). Course of three biochemical bone markers after kidney transplantation. Transplant Proc. 27, 2266–2271. 49. Lechleitner, P., Krimbacher, E., Genser, N., Fridrich, L., zur Nedden, D., Helweg, G., Koenig, P., and Joannidis, M. (1994). Bone mineral densitometry in dialyzed patients: quantitative computed tomography versus dual photon absorptiometry. Bone 15, 387–391. 50. Rambausek, M., Ritz, E., Pomer, S., Mohring, K., and Rohl, L. (1988). Alkaline phosphatase levels in renal transplant recipients receiving cyclosporine or azathioprine/steroids. Lancet 1, 247. 51. Aubia, J., Masramon, J., and Serrano, S. (1988). Bone histology in renal patients receiving cyclosporin. Lancet 1, 1048. 52. Wilmink, J. M., Bras, J., Surachno, S., Heyst, J. L. A. M., and Horst, J. M. V. (1989). Bone repair in cyclosporine treated renal transplant patients. Transplant Proc. 21, 1492–1494. 53. Dumoulin, G., Hory, B., Nguyen, N. U., Henriet, M.-T., Bresson, C., Regnard, J., and Saint-Hillier, Y. (1995). Lack of evidence that cyclosporine treatment impairs calcium-phosphorus homeostasis and bone remodeling in normocalcemic long-term renal transplant recipients. Transplantation 59, 1690–1694. 54. Amado, J. A., Riancho, J. A., Francisco, A. L. M. D., Cotorruelo, J. G., Feijanes, J., Arias, M., Napal, J., and Gonzalez-Macias, J. (1996). Hyperparathyroidism is responsible for the increased levels of osteocalcin in patients with normally functioning kidney grafts. Kidney Int. 50, 1726–1733. 55. Lee, A. H., Mull, R. L., Keenan, G. F., Callegari, P. E., Dalinka, M. K., Eisen, H. J., Manicini, D. M., DiSesa, V. J., and Attie, M. F. (1994).

56.

57.

58.

59.

60.

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63.

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65.

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67.

68.

69.

70.

71. 72.

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Osteoporosis and bone morbidity in cardiac transplant recipients. Am. J. Med. 96, 35–41. Muchmore, J. S., Cooper, D. K. C., Ye, Y., Schlegel, V. J., and Zudhi, N. (1991). Loss of vertebral bone density in heart transplant patients. Transplant Proc. 23, 1184–1185. Shane, E., Mancini, D., Aaronson, K., Silverberg, S. J., Seibel, M. J., Addesso, V., and McMahon, D. J. (1997). Bone mass, vitamin D deficiency and hyperparathyroidism in congestive heart failure. Am. J. Med. 103, 197–207. Shane, E., Addesso, V., Namerow, P., McMahon, D., Lo, S., Staron, R., Zucker, M., Pardi, S., Maybaum, S., and Mancini, D. (2004). Alendronate versus calcitriol for the prevention of bone loss after cardiac transplantation. New. Engl. J. Med. 350, 767–776. Rich, G. M., Mudge, G. H., Laffel, G. L., and LeBoff, M. S. (1992). Cyclosporine A and prednisone-associated osteoporosis in heart transplant recipients. J. Heart Lung Transplant 11, 950–958. Leidig-Bruckner, G., Hosch, S., Dodidou, P., Ritchel, D., Conradt, C., Klose, C., Otto, G., Lange, R., Theilmann, L., Zimmerman, R., Pritsch, M., and Zeigler, R. (2001). Frequency and predictors of osteoporotic fractures after cardiac or liver transplantation: a follow-up study. Lancet. 357, 342–347. Sambrook, P. N., Kelly, P. J., Keogh, A., Macdonald, P., Spratt, P., Freund, J., and Eisman, J. A. (1994). Bone loss after cardiac transplantation: a prospective study. J. Heart Lung Transplant 13, 116–121. Shane, E., Rivas, M., Staron, R. B., Silverberg, S. J., Seibel, M., Kuiper, J., Mancini, D., Addesso, V., Michler, R. E., and Factor-Litvak, P. (1996). Fracture after cardiac transplantation: A prospective longitudinal study. J. Clin. Endocrinol. Metab. 81, 1740–1746. Shane, E., Rivas, M., McMahon, D. J., Staron, R. B., Silverberg, S. J., Seibel, M. J., Mancini, D., Michler, R., Aaronson, K., Addesso, V., and Lo, S. H. (1997). Bone loss and turnover after cardiac transplantation. J. Clin. Endocrinol. Metab. 82, 1497–1506. Van Cleemput, J., Daenen, W., Nijs, J., Geusens, P., Dequeker, J., and Vanhaecke, J. (1995). Timing and quantification of bone loss in cardiac transplant recipients. Transpl. Int. 8, 196–200. Berguer, D. G., Krieg, M. A., Thiebaud, D., Burckhardt, P., Stumpe, F., Hurni, M., Sadeghi, H., Kappenberger, L., and Goy, J. J. (1994). Osteoporosis in heart transplant recipients: a longitudinal study. Transplant Proc. 26, 2649–2651. Henderson, N. K., Sambrook, P. N., Kelly, P. J, Macdonald, P., Keogh, A. M., Spratt, P., and Eisman, J. A. (1995). Bone mineral loss and recovery after cardiac transplantation [letter]. Lancet 346, 905. Sambrook, P. N., Kelly, P. J., Fontana, D., Nguyen, T., Keogh, A., Macdonald, D., Spratt, P., Freund, J., and Eisman, J. A. (1994). Mechanisms of rapid bone loss following cardiac transplantation. Osteoporosis. Int. 4, 273–276. Thiebaud, D., Krieg, M., Gillard-Berguer, D., Jaquet, A., Goy, J., and Burckhardt, P. (1996). Cyclosporine induces high turnover and may contribute to bone loss after heart transplantation. Eur. J. Clin. Invest. 26, 549–555. Prummel, M. F., Wiersinga, W. M., Lips, P., Sanders, G. T. P., and Sauerwein, H. P. (1991). The course of biochemical parameters of bone turnover during treatment with corticosteroids. J. Clin. Endocrinol. Metab. 72, 382–386. Lems, W. F., Gerrits, M. I., Jacobs, J. W. G., Vugt, R. M. V., Rijn, H. J. M. V., and Bijlsma, J. W. J. (1996). Changes in (markers of) bone metabolism during high dose corticosteroid pulse treatment in patients with rheumatoid arthritis. Ann. Rheum. Dis. 55, 288–293. Guo, C., Johnson, A., Locke, T., and Eastell, R. (1998). Mechanism of bone loss after cardiac transplantation. Bone 22, 267–271. Monegal, A., Navasa, M., Guanabens, N., Peris, P., Pons, F., Osaba, M. J. M. D., Rimola, A., Rodes, J., and Munoz-Gomez, J. (1997). Osteoporosis and bone mineral metabolism in cirrhotic patients referred for liver transplantation. Calcif. Tissue Int. 60, 148–154.

714 73. Ninkovic, M., Love, S. A., Tom, B., Alexander, G. J., and Compston, J. E. (2001). High prevalence of osteoporosis in patients with chronic liver disease prior to liver transplantation. Calcif. Tissue Int. 69, 321–326. 74. Hodgson, S. F., Dickson, E. R., Wahner, H. W., Johnson, K. A., Mann, K. G., and Riggs, B. L. (1980). Bone loss and reduced osteoblast function in primary biliary cirrhosis. Ann. Int. Med. 103, 855–860. 75. Guanabens, N., Pares, A., Alvarez, L., Osaba, M. J. M. D., Monegal, A., Peris, P., Ballesta, A. M., and Rodes, J. (1998). Collagen-related markers of bone turnover reflect the severity of liver fibrosis in patients with primary biliary cirrhosi. J. Bone Miner. Res. 13, 731–738. 76. Guanabens, N., Pares, A., Marinoso, L., Brancos, M. A., Piera, C., Serrano, S., Rivera, F., and Rodes, J. (1990). Factors influencing the development of metabolic bone disease in primary biliary cirrhosis. Am. J. Gastroenterol. 85, 1356–1362. 77. Hodgson, S. F., Dickson, E. R., Eastell, R., Eriksen, E. F., Bryant, S. C., and Riggs, B. L. (1993). Rates of cancellous bone remodeling and turnover in osteopenia associated with primary biliary cirrhosis. Bone 14, 819–827. 78. Vedi, J., Greer, S., Skingle, S., Garrahan, N., Ninkovic, M., Alexander, G., and Compston, J. (1999). Mechanism of bone loss after liver transplantation: A histomorphometric analysis. J. Bone Miner. Res. 14, 281–287. 79. Janes, C. H., Dickson, E. R., Okazaki, R., Bonde, S., McDonagh, A. F., and Riggs, B. L. (1995). Role of hyperbilirubinemia in the impairment of osteoblast proliferation associated with cholestatic jaundice. J. Clin. Invest. 95, 2581–2586. 80. Haagsma, E. B., Thijn, C. J. P., Post, J. G., Slooff, M. J. H., and Gisp, C. H. (1988). Bone disease after liver transplantation. J. Hepatol. 6, 94–100. 81. Arnold, J. C., Hauser, R., Ziegler, R., Kommerelli, B., Otto, G., Theilmann, L., and Wuster, C. (1992). Bone disease after liver transplantation. Transplant Proc. 24, 2709–2710. 82. McDonald, J. A., Dunstan, C. R., Dilworth, P., Sherbon, K., Sheil, A. G. R., Evans, R. A., and McCaughan, G. W. (1991). Bone loss after liver transplantation. Hepatology 14, 613–619. 83. Naiker, I. P., Govender, S., Naicker, S., Dawood, S., Haffejee, A. A., and Seedat, Y. K. (1993). Avascular necrosis of bone following renal transplantation. S. Afr. Med. J. 83, 646–649. 84. Eastell, R., Dickson, R. E., Hodgson, S. F., Wiesner, R. H., Porayko, M. K., Wahner, H. W., Cedel, S. L., Riggs, B. L., and Krom, R. A. F. (1991). Rates of vertebral bone loss before and after liver transplantation in women with primary biliary cirrhosis. Hepatology 14, 296–300. 85. Meys, E., Fontanges, E., Fourcade, N., Thomasson, A., Pouyet, M., and Delmas, P. (1994). Bone loss after orthotopic liver transplantation. Am. J. Med. 97, 445–450. 86. Navasa, M., Monegal, A., Guanabens, N., Peris, P., Rimola, A., MunozGomez, J., Visa, J., and Rodes, J. (1994). Bone fractures in liver transplant patients. Brit. J. Rheumatol. 33, 52–55. 87. Ninkovic, M., Love, S., Tom, B. D., Bearcroft, P. W., Alexander, G. J., and Compston, J. E. (2002). Lack of effect of intravenous pamidronate on fracture incidence and bone mineral density after orthotopic liver transplantation. J. Hepatol. 37, 93–100. 88. Compston, J., Greer, S., Skingle, S., Stirling, D., Price, C., Friend, P., and Alexander, G. (1996). Early increase in plasma parathyroid hormone level following liver transplantation. J. Hepatol. 25, 715–718. 89. Hawkins, F. G., Leon, M., Lopez, M. B., Valero, M. A., Larrodera, L., Garcia-Garcia, I., Loinaz, C., and Gonzales, E. M. (1994). Bone loss and turnover in patients with liver transplantation. HepatoGastroenterol. 41, 158–161. 90. Valero, M., Loinaz, C., Larrodera, L., Leon, M., Morena, E., and Hawkins, F. (1995). Calcitonin and bisphosphonate treatment in bone loss after liver transplantation. Calcif. Tissue Int. 57, 15–19. 91. Rabinowitz, M., Shapiro, J., Lian, J., Bloch, G. D., Merkel, I. S., and Thiel, D. H. V. (1992). Vitamin D and osteocalcin levels in liver

Carolina A. Moreira Kulak and Elizabeth Shane

92.

93.

94.

95.

96.

97.

98. 99.

100.

101.

102.

103.

104.

105.

106.

107.

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transplant recipients. Is osteocalcin a reliable marker of bone turnover in such cases? J. Hepatol. 16, 50–55. Watson, R. G., Coulton, L., Kanis, J. A., Lombard, M., Williams, R., Neuberger, J., and Elias, E. (1990). Circulating osteocalcin in primary biliary cirrhosis following liver transplantation and during treatment with ciclosporin. J. Hepatol. 11, 354–358. Abdelhadi, M., Eriksson, S. A., Ljusk Eriksson, S., Ericzon, B. G., and Nordenstrom, J. (1995). Bone mineral status in end-stage liver disease and the effect of liver transplantation. Scand. J. Gastroenterol. 30, 1210–1215. Aris, R., Neuringer, I., Weiner, M., Egan, T., and Ontjes, D. (1996). Severe osteoporosis before and after lung transplantation. Chest 109, 1176–1183. Shane, E., Silverberg, S. J., Donovan, D., Papadopoulos, A., Staron, R. B., Addesso, V., Jorgensen, B., McGregor, C., and Shulman, L. (1996). Osteoporosis in lung transplantation candidates with end stage pulmonary disease. Am. J. Med. 101, 262–269. Tschopp, O., Boehler, A., Speich, R., Weder, W., Seifert, B., Russi, E. W., and Schmid, C. (2002). Osteoporosis before lung transplantation: association with low body mass index, but not with underlying disease. Am. J. Transplant. 2, 167–172. Aris, R. M., Renner, J. B., Winders, A. D., Buell, H. E., Riggs, D. B., Lester, G. E., and Ontjes, D. A. (1998). Increased rate of fractures and severe kyphosis: sequelae of living into adulthood with cystic fibrosis. Ann. Intern. Med. 128, 186–193. Bachrach, L. K., Loutit, C. W., and Moss, R. B. (1994). Osteopenia in adults with cystic fibrosis [see comments]. Am. J. Med. 96, 27–34. Donovan, D. S., Jr., Papadopoulos, A., Staron, R. B., Addesso, V., Schulman, L., McGregor, C., Cosman, F., Lindsay, R. L., and Shane, E. (1998). Bone mass and vitamin D deficiency in adults with advanced cystic fibrosis lung disease. Am. J. Respir. Crit. Care Med. 157, 1892–1899. Grey, A. B., Ames, R. W., Matthews, R. D., and Reid, I. R. (1993). Bone mineral density and body composition in adult patients with cystic fibrosis [see comments]. Thorax 48, 589–593. Aringer, M., Kiener, H., Koeller, M., Artemiou, O., Zuckerman, A., Wieselthaler, G., Klepetko, W., Seidl, G., Kainberger, F., Barnecker, P., Smolen, J., and Pietschmann, P. (1998). High turnover bone disease following lung transplantation. Bone 23, 485–488. Ferrari, S. L., Nicod, L. P., Hamacher, J., Spiliopoulos, A., Slosman, D. O., Rochat, T., Bonjour, J.-P., and Rizzoli, R. (1996). Osteoporosis in patients undergoing lung transplantation. Eur. Respir. J. 9, 2378–2382. Shane, E., Papadopoulos, A., Staron, R. B., Addesso, V., Donovan, D., McGregor, C., and Schulman, L. L. (1999). Bone loss and fracture after lung transplantation. Transplantation 68, 220–227. Spira, A., Gutierrez, C., Chaparro, C., Hutcheon, M. A., and Chan, C. K. (2000). Osteoporosis and lung transplantation: a prospective study. Chest 117, 476–481. Trombetti, A., Gerbase, M. W., Spiliopoulos, A., Slosman, D. O., Nicod, L. P., and Rizzoli, R. (2000). Bone mineral density in lungtransplant recipients before and after graft: prevention of lumbar spine post-transplantation-accelerated bone loss by pamidronate. J. Heart Lung Transplant. 19, 736–743. Schulte, C., Beelen, D., Schaefer, U., and Mann, K. (2000). Bone loss in long-term survivors after transplantation of hematopoietic stem cells: a prospective study. Osteoporos. Int. 11, 344–353. Stern, J. M., Chesnut, C. H., 3rd, Bruemmer, B., Sullivan, K. M., Lenssen, P. S., Aker, S. N., and Sanders, J. (1996). Bone density loss during treatment of chronic GVHD. Bone Marrow Transplant 17, 395–400. Ebeling, P., Thomas, D., Erbas, B., Hopper, L., Szer, J., and Grigg, A. (1999). Mechanism of bone loss following allogeneic and autologous hematopoeitic stem cell transplantation. J. Bone Miner. Res. 14, 342–350.

Chapter 42 Transplantation Osteoporosis: Biochemical Correlates of Pathogenesis and Treatment 109. Kang, M. I., Lee, W. Y., Oh, K. W., Han, J. H., Song, K. H., Cha, B. Y., Lee, K. W., Son, H. Y., Kang, S. K., and Kim, C. C. (2000). The shortterm changes of bone mineral metabolism following bone marrow transplantation. Bone 26, 275–279. 110. Valimaki, M., Kinnunen, K., Volin, L., Tahtela, R., Loyttniemi, E., Laitinen, K., Makela, P., Keto, P., and Ruutu, T. (1999). A prospective study of bone loss and turnover after allogeneic bone marrow transplantation: effect of calcium supplementation with or without calcitonin. Bone Marrow Transplant. 23, 355–361. 111. Lee, W. Y., Cho, S. W., Oh, E. S., Oh, K. W., Lee, J. M., Yoon, K. H., Kang, M. I., Cha, B. Y., Lee, K. W., Son, H. Y., Kang, S. K., and Kim, C. C. (2002). The effect of bone marrow transplantation on the osteoblastic differentiation of human bone marrow stromal cells. J. Clin. Endocrinol. Metab. 87, 329–335. 112. Carlson, K., Simonsson, B., and Ljunghall, S. (1994). Acute effects of high dose chemotherapy followed by bone marrow transplantation on serum markers of bone metabolism. Calcif. Tissue Int. 55, 408–411. 113. Tauchmanova, L., Serio, B., Del Puente, A., Risitano, A. M., Esposito, A., De Rosa, G., Lombardi, G., Colao, A., Rotoli, B., and Selleri, C. (2002). Long-lasting bone damage detected by dual-energy x-ray absorptiometry, phalangeal osteosonogrammetry, and in vitro growth of marrow stromal cells after allogeneic stem cell transplantation. J. Clin. Endocrinol. Metab. 87, 5058–5065. 114. Lien, D., Jackson, K., Monkhouse, P., Halenar, J., Mullen, J., Winton, T., Modry, D., and Siminoski, K. (2001). Intervention prevents progression of osteoporosis in patients awaiting lung transplantation. J. Heart Lung Transplant. 20, 225. 115. Fan, S. L., and Cunningham, J. (2001). Bisphosphonates in renal osteodystrophy. Curr. Opin. Nephrol. Hypertens. 10, 581–588. 116. Braith, R. W., Mills, J., Welsch, M. A., Keller, J. W., and Pollack, M. L. (1996). Resistance exercise training restores bone mineral density in heart transplant recipients. J. Am. Coll. Cardiol. 28, 1471–1477. 117. Braith, R. W., Magyari, P. M., Fulton, M. N., Aranda, J., Walker, T., and Hill, J. A. (2003). Resistance exercise training and alendronate reverse glucocorticoid-induced osteoporosis in heart transplant recipients. J. Heart Lung Transplant. 22, 1082–1090. 118. Mitchell, M. J., Baz, M. A., Fulton, M. N, Lisor, C. F., and Braith, R. W. (2003). Resistance training prevents vertebral osteoporosis in lung transplant recipients. Transplantation 76, 557–562. 119. Valimaki, M. J., Kinnunen, K., Tahtela, R., Loyttyniemi, E., Laitinen, K., Makela, P., Keto, P., and Nieminen, M. (1999). A prospective study of bone loss and turnover after cardiac transplantation: effect of calcium supplementation with or without calcitonin. Osteoporos. Int. 10, 128–136. 120. Hay, J. E., Malinchoc, M., and Dickson, E. R. (2001). A controlled trial of calcitonin therapy for the prevention of post-liver transplantation atraumatic fractures in patients with primary biliary cirrhosis and primary sclerosing cholangitis. J. Hepatol. 34, 292–298. 121. Cremer, J., Struber, M., Wagenbreth, I., Nischelsky, J., Demertzis, S., Graeter, T., Abraham, C., and Haverich, A. (1999). Progression of steroid-associated osteoporosis after heart transplantation. Ann. Thorac. Surg. 67, 130–133. 122. Meys, E., Terreaux-Duvert, F., Beaume-Six, T., Dureau, G., and Meunier, P. J. (1993). Effects of calcium, calcidiol, and monofluorophosphate on lumbar bone mass and parathyroid function in patients after cardiac transplantation. Osteoporosis. Int. 3, 329–332. 123. Talalaj, M., Gradowska, L., Marcinowska-Suchowierska, E., Durlik, M., Gaciong, Z., and Lao, M. (1996). Efficiency of preventive treatment of glucocorticoid-induced osteoporosis with 25-hydroxyvitamin D3 and calcium in kidney transplant patients. Transplant. Proc. 28, 3485–3487. 124. Torres, J., Garcia, S., Gomez, A., Gonzalez, A., Barrios, Y., Concepcion, M. T., Hernandez, D., Garcia, J. J., Checa, M. D., Lorenzo, V., Salido, E. (2004). Treatment with intermittent calcitriol

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and calcium reduces bone loss after renal transplantation. Kidney Int. 65, 705–712. Sambrook, P., Henderson, N. K., Keogh, A., MacDonald, P., Glanville, A., Spratt, P., Bergin, P., Ebeling, P., and Eisman, J. (2000). Effect of calcitriol on bone loss after cardiac or lung transplantation. J. Bone Miner. Res. 15, 1818–1824. Fan, S., Almond, M. K., Ball, E., Evans, K., and Cunningham, J. (2000). Pamidronate therapy as prevention of bone loss following renal transplantation. Kidney Int. 57, 684–690. Fan, S. L., Kumar, S., and Cunningham, J. (2003). Intravenous pamidronate as treatment for osteoporosis after kidney transplantation: a prospective study. Kidney Int. 63, 2275–2279. Aris, R. M., Lester, G. E., Renner, J. B., Winders, A., Blackwood, A. D., Lark, R. K., and Ontjes, D. A. (2000). Efficacy of pamidronate for osteoporosis in cystic fibrosis patients following lung transplantation. Am. J. Respir. Crit. Care Med. 162, 941. Krieg, M., Seydoux, C., Sandini, L., Goy, J. J., Berguer, D. G., Thiebaud, D., and Bruckhardt, P. (2001). Intravenous pamidronate as a treatment for osteoporosis after heart transplantation: a prospective study. Osteoporos. Int. 12, 112–116. Shane, E., Rodino, M. A., McMahon, D. J., Addesso, V., Staron, R. B., Seibel, M. J., Mancini, D., Michler, R. E., and Lo, S. H. (1998). Prevention of bone loss after heart transplantation with antiresorptive therapy: a pilot study. J. Heart Lung Transplant. 17, 1089–1096. Reeves, H., Francis, R., Manas, D., Hudson, M., and Day, C. (1998). Intravenous bisphosphonate prevents symtomatic osteoporotic vertebral collapse in patients after liver tranplantation. Liver Transpl. Surg. 4, 404–409. Coco, M., Glicklich, D., Faugere, M. C., Burris, L., Bognar, I., Durkin, P., Tellis, V., Greenstein, S., Schechner, R., Figueroa, K., McDonough, P., Wang, G., Malluche, H. (2003). Prevention of bone loss in renal transplant recipients: A prospective randomized trial of intravenous pamidronate. J. Am. Coll. Nephrol. 14, 2669–2676. Kovac, D., Lindic, J., Kandus, A., and Bren, A. F. (2001). Prevention of bone loss in kidney graft recipients. Transplant. Proc. 33, 1144–1145. Schwarz, C., Mitterbauer, C., Heinze, G., Woloszczuk, W., Haas, M., and Oberbauer, R., (2004). Nonsustained effect of short-term bisphosphonate therapy on bone turnover three years after renal transplantation. Kidney Int. 65, 304–309. Hommann, M., Abendroth, K., Lehmann, G., Patzer, N., Kornberg, A., Voigt, R., Seifert, S., Hein, G., and Scheele, J. (2002). Effect of transplantation on bone: osteoporosis after liver and multivisceral transplantation. Transplant. Proc. 34, 2296–2298. Grotz, W., Nagel, C., Poeschel, D., Cybulla, M., Petersen, K. G., Uhl, M., Strey, C., Kirste, G., Olschewski, M., Reichelt, A., and Rump, L. C. (2001). Effect of ibandronate on bone loss and renal function after kidney transplantation. J. Am. Soc. Nephrol. 12, 1530–1537. Ugur, A., Guvener, N., Isiklar, I., Karakayali, H., and Erdal, R. (2000). Efficiency of preventive treatment for osteoporosis after renal transplantation. Transplant. Proc. 32, 556–557. Stempfle, H. U., Werner, C., Siebert, U., Assum, T., Wehr, U., Rambeck, W. A., Meiser, B., Theisen, K., and Gartner, R. (2002). The role of tacrolimus (FK506)-based immunosuppression on bone mineral density and bone turnover after cardiac transplantation: a prospective, longitudinal, randomized, double-blind trial with calcitriol. Transplantation 73, 547–552. Stempfle, H. U., Werner, C., Echtler, S., Wehr, U., Rambeck, W. A., Siebert, U., Uberfuhr, P., Angermann, C. E., Theisen, K., and Gartner, R. (1999). Prevention of osteoporosis after cardiac transplantation: a prospective, longitudinal, randomized, double-blind trial with calcitriol. Transplantation 68, 523–530. Dodidou, P., Bruckner, T., Hosch, S., Haass, M., Klar, E., Sauer, P., Ziegler, R., and Leidig-Bruckner, G. (2003). Better late than never?

716 Experience with intravenous pamidronate treatment in patient with low bone mass or fractures following cardiac or liver transplantation. Osteoporos. Int. 14, 82–88. 141. Idilman, R., de Maria, N., Uzunalimoglu, O., and van Thiel, D. H. (1997). Hepatic osteodystrophy: a review. Hepatogastroenterol 44, 574–581. 142. Grotz, W., Rump, A. L., Niessen, H., Schmidt-Gayt, A., Reichelt, G., Kirste, G., Olchewski, G., and Schollmeyer, P. (1998). Treatment of osteopenia and osteoporosis after kidney transplantation. Transplantation 66, 1004–1008. 143. Giannini, S., Dangel, A., Carraro, G., Nobile, M., Rigotti, P., Bonfante, L., Marchini, F., Zaninotto, M., Dalle Carbonare, L.,

Carolina A. Moreira Kulak and Elizabeth Shane Sartori, L., and Crepaldi, G. (2001). Alendronate prevents further bone loss in renal transplant recipients. J. Bone Miner. Res. 16, 2111–2117. 144. Jeffrey, J. R., Leslie, W. D., Karpinski, M. F., Nickerson, P. W., and Rush, D. N. (2003). Prevalence and treatment of decreased bone density in renal transplant recipients: a randomized prospective trial of calcitriol versus alendronate. Transplantation 76, 1498–1502. 145. Koc, M., Tuglular, S., Arikan, H., Ozener, C., and Akoglu, E. (2002). Alendronate increases bone mineral density in long-term renal transplant recipients. Transplant. Proc. 34, 2111–2113.

Chapter 43

Secondary Osteoporosis Jean E. Mulder, M.D. Carolina A. Moreira Kulak, M.D. Elizabeth Shane, M.D.

I. Introduction II. Hyperthyroidism and Osteoporosis III. Osteoporosis Secondary to Hypogonadism

Division of Endocrinology and Metabology of Hospital de Clinicas, Federal university of Parana (SEMPR), Curitiba-/Brazil Department of Medicine, College of Physicians and Surgeons, Columbia University, New York, New York 10032

IV. Anticonvulsant Drugs and Osteoporosis References

II. HYPERTHYROIDISM AND OSTEOPOROSIS

I. INTRODUCTION Osteoporosis is the most common metabolic bone disease worldwide. The term primary osteoporosis is generally used to describe the progressive bone loss which occurs with aging, in both men and women, but especially in postmenopausal women. Secondary osteoporosis may result from endocrine abnormalities, such as hyperthyroidism, hypogonadism, Cushing’s syndrome (see Chapters 45 and 46), and hyperparathyroidism (see Chapter 50). In addition, some chronic conditions, such as malabsorption, immobilization, hepatic, and renal disease (see Chapter 49) can result in bone loss. Finally many drugs also cause secondary osteoporosis (Table I). In this chapter, we will review the impact of some of these more commonly encountered disorders of the skeleton, with specific emphasis on bone mineral density and parameters of bone remodeling. Dynamics of Bone and Cartilage Metabolism

Department of Medicine, Brigham and Women’s Hospital, Harvard University, Boston, MA 02115

The relationship between hyperthyroidism and bone loss was described more than 100 years ago, when a postmortem study in a thyrotoxic patient revealed excessive bone destruction [1]. Subsequent histomorphometric studies of bone in hyperthyroidism support this early observation. Reductions in the resorption and formation phases of the bone remodeling cycle have been noted, resulting in an overall increase in the activation frequency of the remodeling cycle. Both osteoblast and osteoclast activities are enhanced. Specifically, an increased calcification rate, increased resorption surface, and a decreased mineralization lag time have been described in cancellous bone of hyperthyroid patients. In cortical bone, increases in active resorption and cortical porosity have also been reported [2, 3]. Typical of the adult skeleton subjected 717

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Secondary Osteoporosis

Hypogonadal states Turner’s syndrome Athletic amenorrhea Anorexia nervosa Hyperprolactinemia GnRH therapy Hypothalamic amenorrhea Thyroid disorders Other endocrine disorders Hyperparathyroidism Diabetes mellitus Hemochromatosis Acromegaly Medications Glucocorticoids Anticonvulsant drugs Cyclosporine Diuretics Anticoagulants Methotrexate Renal disorders Gastrointestinal disorders Primary biliary cirrhosis Inflammatory bowel disease Celiac disease and sprue Hepatic diseases Hematologic disorders Multiple myeloma Systemic mastocytosis Lymphoproliferative disorders Rheumatologic disorders Immobilization Genetic disorders Osteogenesis imperfecta Vitamin D-resistant osteomalacia Hypophosphatasia Marfan syndrome Ehlers–Danlos Glycogen storage diseases

Chapter 43

Chapter 43 Chapter 46

to subclinical hyperthyroidism. Osteoporosis is also multifactorial in etiology and is influenced by genetics, hormonal, nutritional, and lifestyle factors. Therefore, the impact of hyperthyroidism on bone loss and its long-term consequences are variable, depending on the nature and severity of the thyroid disease as well as on individual bone health and the presence of other risk factors for osteoporosis. The increased availability of bone densitometry and biochemical markers of bone remodeling has resulted in a greater understanding of thyroid-related bone loss. In this section, we will review the impact of hyperthyroidism on bone remodeling, with special emphasis on changes in bone mineral density (BMD) and parameters of bone remodeling.

Chapters 18, 41, 42 Chapter 43 Chapter 42

A. Hyperthyroidism (Endogenous) Chapter 45

Chapter 48

Chapter 51 Chapter 40 Chapters 28, 36 Chapters 49, 50 Chapters 18, 44

to events associated with high bone turnover, untreated hyperthyroidism results in a net negative bone balance and reduced bone mass [4]. The mechanism of the deleterious effect of excessive thyroid hormone on bone has not been fully clarified. Stimulation of bone-resorbing cytokines, such as interleukin-6 and interleukin 1B, by thyroid hormone is one proposed mechanism [5, 6]. Recent evidence also suggests that TSH may have direct effects on skeletal remodeling, mediated by the TSH receptor on osteoblast and osteoclast precursors [7]. Hyperthyroidism is multifactorial in etiology and may result from autoimmune disease (Graves’ disease), toxic nodular or multinodular goiter, thyroiditis, or excessive exogenous thyroid hormone intake. In addition, hyperthyroidism can vary in severity, from severe thyrotoxicosis

BMD is decreased in both premenopausal and postmenopausal women with thyrotoxicosis [8–12]. Values 10–30% lower than age-matched controls have been reported [8–12]. Normalization of thyroid function results in improvement in BMD. Lupoli reported a 26% increase in spinal bone density (dual energy X-ray absorptiometry; DXA) in premenopausal women treated with antithyroid drugs for 12 months and an 18.6% increase in postmenopausal patients treated with antithyroid drugs plus hormone replacement therapy (HRT) [11]. However, final BMD in these patients was still lower than BMD in the control group matched for age and menopausal status. Interestingly, both pre- and postmenopausal patients treated with alendronate, in addition to antithyroid drugs and HRT (postmenopausal women only), achieved greater increases in spinal BMD after 12 months. Final BMD in this subgroup was not significantly different from the control population. It is unclear whether BMD would have normalized in the patients treated solely with antithyroid drugs with longer follow-up. In this regard, a 5-year evaluation of lumbar spine BMD in a very small group of men and women with treated hyperthyroidism has demonstrated complete restoration of vertebral bone density after antithyroid therapy [13]. The single postmenopausal woman in the study experienced a 9.8% increase in lumbar spine BMD over 5 years. Most studies report an increase in BMD of 1–11% in women after resolution of hyperthyroidism [8–10, 12, 13]. Of note, intranasal calcitonin has not been shown to improve BMD beyond the improvement noted with correction of thyroid function [12]. Few studies have addressed the effects of thyrotoxicosis on bone mass in men [8, 14–16]. A single prospective study addressing the effect of Graves’ disease on radial BMD in men revealed a lower baseline BMD in hyperthyroid patients compared with age-matched controls [15].

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Treatment with either radioactive iodine or antithyroid drugs resulted in an initial decrease in BMD followed by a return to baseline after 2 years. Radial BMD (assessed by single photon absorptiometry; SPA) in men with Graves’ disease remained lower than controls at 2 years. However, in a 2-year prospective follow-up study of ten patients with Graves’ disease (seven males), mean LS BMD (DXA) increased significantly after treatment with antithyroid drugs [17]. The decrease in BMD associated with hyperthyroidism is secondary to an increase in bone remodeling with bone resorption exceeding bone formation. Well-documented increases in serum and urinary markers of bone turnover in thyrotoxic patients support this premise (Fig. 1). Urinary markers of bone resorption (hydroxyproline, pyridinium cross-links) are increased in both pre- and postmenopausal women with untreated Graves’ disease [18, 19] and in hyperthyroidism in general [20]. Pyridinium cross-links, whether measured by HPLC [19, 20] or by direct immunoassay [21], are elevated in nearly all thyrotoxic patients and correlate well with free thyroxine (T4) and free triiodothyronine (T3) levels [19, 21]. Urinary assays which measure type I collagen C-telopeptide also correlate well with free T3 and T4 levels [22]. Elevated serum markers of bone formation (total and bone-specific alkaline phosphatase, osteocalcin, or PICP, the carboxyterminal propeptide of type 1 procollagen) have also been reported in Graves’ disease [10, 16–18, 21, 23] and in patients with toxic nodular goiter and amiodarone-induced hyperthyroidism [21]. Osteocalcin [18, 21] and PICP [18] levels correlate well

Figure 1

with free T3 [21, 23], total T4 [23], and free T4 levels [18]. Osteocalcin levels have also been reported to correlate negatively with BMD Z scores at the spine, femoral neck, and trochanter [16]. However, Garnero et al. demonstrated three-fold higher elevations in bone resorption markers than in formation markers, supporting the hypothesis of an imbalance between resorption and formation, favoring bone loss [21]. Beta receptor antagonists, frequently used in the therapy of Graves’ disease, do not appear to alter the rise in pyridinium cross-links, and therefore probably do not inhibit thyroid hormone-mediated bone resorption [19]. Radioactive iodine or antithyroid drug therapy, on the other hand, result in a prompt reduction in pyridinium crosslinks, with normalization by approximately 10–12 weeks. In two studies, osteocalcin levels remained unchanged during this time interval [17, 19]. In contrast, Garrel et al. demonstrated normalization of osteocalcin levels by 16 weeks after treatment of hyperthyroidism in the 70% of patients (men and women) with elevated baseline levels [23]. In premenopausal women, euthyroidism after 6 months of methimazole therapy was associated with significant reductions in serum PICP and urinary hydroxyproline excretion. There was a non-significant decrease in osteocalcin and total alkaline phosphatase activity [18]. In contrast, Garnero et al. observed that osteocalcin, measured by ELISA, decreased significantly by 1 month after therapy with antithyroid drugs, radioactive iodine, or surgery in 11 of 27 patients evaluated [19]. In the same study, the pattern of change of bone-specific alkaline phosphatase

Serum osteocalcin (A), bone-specific alkaline phosphatase (B), and urinary pyridinoline crosslinks (C) in patients with endogenous hyperthyroidism. (N), age-matched normal subjects; (P), patient values. Dotted lines represent the normal ranges +/− 2 SD. Reproduced with permission from: Garnero et al. [21] (1994). The Endocrine Society. Markers of bone turnover in hyperthyroidism and the effects of treatment. JCEM 78: 955–959.

720 differed, increasing after 1 month of therapy and decreasing 4 months after therapy. The discrepancy between normalization of osteocalcin, bone-specific alkaline phosphatase, and PICP, all markers of bone formation, may be explained by differences in assay technique, osteoblast function, release from bone, and clearance of the markers [18, 21]. Given that bone-specific alkaline phosphatase values were elevated in <50% of hyperthyroid patients studied and that levels did not correlate with the degree of hyperthyroidism, it appears to be a less sensitive marker of hyperthyroidinduced bone turnover than osteocalcin or PICP [21]. In summary, the decrease in bone mineral density and the elevation of markers of bone remodeling in patients with overt hyperthyroidism support the histologic findings of increased cortical porosity and increased cancellous resorption surfaces and calcification rates, indicating accelerated bone turnover and net loss of bone. The increase in BMD and improvement of markers of bone remodeling with normalization of thyroid function suggest that the bone loss is, at least in part, reversible. However, complete recovery, especially in postmenopausal women, is not documented consistently. Furthermore, Franklyn [24] and others [25, 26] have demonstrated that a previous history of thyrotoxicosis in postmenopausal women conferred an increased risk of low BMD, even in patients who had been euthyroid for many years. In contrast, premenopausal women with a history of thyrotoxicosis did not have an increased risk of low BMD [24]. Thus, estrogen may protect premenopausal women from the deleterious effects of excess endogenous thyroxine production on BMD. Further prospective longitudinal studies are needed to determine if bone loss is completely reversible in all patients and whether intervention with antiresorptive drugs is of benefit, when recovery is not complete.

JEAN E. MULDER ET AL.

thyroid nodules and subclinical hyperthyroidism, Foldes et al. reported no significant difference in Z scores (DXA) at any site compared with the control group [29]. In contrast, 52 postmenopausal women with subclinical hyperthyroidism had significantly lower Z scores at both the femoral neck and radius, but not at the lumbar spine, compared with the postmenopausal control group (Fig. 2). Osteopenia was present, but osteoporosis was rarely detected. The cross-sectional study design prohibits the assessment of the impact of the duration of subclinical hyperthyroidism on bone density. In addition, there are often

B. Subclinical Hyperthyroidism (Endogenous) Patients with multinodular goiter with autonomy, solitary autonomous functioning nodules, or inadequately treated Graves’ disease often develop thyroid hormone abnormalities characterized by a suppressed TSH level with a normal T4 level, a pattern which has been designated subclinical hyperthyroidism. Endogenous subclinical hyperthyroidism has also been associated with a decrease in bone mineral density in pre- and postmenopausal women [27], although the data in premenopausal women are conflicting. Faber et al. [28] observed no change in bone mineral density of the distal forearm or lumbar spine over 2 years in premenopausal women with goiter and subclinical hyperthyroidism. Similarly, in a cross-sectional study of 39 premenopausal women with autonomously functioning

Figure 2

Z-scores for vertebral (A), femoral neck (B), and midshaft radial (C) BMD values in premenopausal (closed circles) and postmenopausal (open circles) women with nodular goiter and either normal thyroid function (left panel), endogenous subclinical hyperthyroidism (middle panel), or endogenous overt hyperthyroidism (right panel). The horizontal bars represent the mean values of the groups. Asterisks denote significant differences from a sex- and age-matched population. *P < 0.05, **P < 0.01, ***P < 0.001. Reproduced with permission from: Foldes et al. [29] (1993). Bone mineral density in patients with endogenous subclinical hyperthyroidism: is this thyroid status a risk factor for osteoporosis. Clin. Endocrinol. 39: 521–527.

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large individual variations in BMD values within patient subgroups. Therefore, it is difficult to conclude from the Foldes study [29] if postmenopausal women had lower bone density Z scores at the femoral neck and radius because they had a longer duration of subclinical hyperthyroidism, whether bone loss due to estrogen deficiency is accelerated by the presence of subclinical hyperthyroidism, or whether some women had suboptimal peak bone densities earlier in life, independent of thyroid dysfunction. Faber et al. [30] have reported an increase in femoral neck (FN) BMD (no change in LS BMD) in 16 postmenopausal women with nodular goiter and subclinical hyperthyroidism who received treatment with RAI. In contrast, patients who did not receive RAI (n = 12) had progressive decreases in both FN and spine BMD. These data suggest that subclinical hyperthyroidism in postmenopausal women can result in bone loss, which may be reversible with normalization of thyroid function. Studies of markers of bone remodeling in patients with endogenous subclinical hyperthyroidism are conflicting. Faber et al. demonstrated elevated osteocalcin levels in six of 44 patients with non-toxic goiter. Osteocalcin levels correlated negatively with TSH levels in both pre- and postmenopausal women [31]. However, Foldes et al. did not report a significant difference in osteocalcin levels in pre- or postmenopausal women with subclinical hyperthyroidism compared with an appropriately matched control population [29]. More recent studies have demonstrated increased urinary deoxypyridinoline (DPD) excretion in pre- and postmenopausal patients with subclinical hyperthyroidism compared with controls, indicating excessive bone resorption [32–34]. The discrepancies among these reports may be related to the duration of subclinical hyperthyroidism or to the impact of excessive thyroid hormone on bone in the presence of estrogen deficiency. It is also possible that other local growth factors, such as insulin-like growth factor (IGF-I), may be involved in mediating the effects of thyroid hormone on the skeleton. IGF-I has stimulatory effects on osteoblasts and is a known stimulator of bone formation. IGF-I levels have been measured in both pre- and postmenopausal women with hyperthyroidism [34]. Whereas premenopausal women with overt hyperthyroidism had elevated IGF-I levels, postmenopausal women with either subclinical or overt hyperthyroidism did not. This relative IGF-I deficiency may contribute to the reduced bone mass observed in postmenopausal women. Taken together, the above data suggest that endogenous subclinical hyperthyroidism can result in increased bone resorption and a decrease in bone mineral density in postmenopausal women. However, premenopausal women with endogenous subclinical hyperthyroidism appear to be at lower risk for

reduced bone mineral density, despite increased bone resorption.

C. Hyperthyroidism (Exogenous) Overt hyperthyroidism (elevated T4 and/or T3 with suppressed TSH) secondary to excessive levothyroxine (L-T4) therapy occurs infrequently, given the advent of ultrasensitive TSH assays and an increased awareness of the potential adverse effects of excess exogenous L-T4 on BMD and cardiac function. The data suggest that exogenous L-T4, in doses sufficient to cause hyperthyroidism, results in decreased BMD and an increased risk of fractures [35]. Histomorphometric analysis of bone in such patients has revealed increased cancellous and cortical bone turnover similar to that reported in endogenous hyperthyroidism [3, 35].

D. Subclinical Hyperthyroidism (Exogenous) Suppressive therapy with L-T4 is often used in the treatment of nodular thyroid disease and in thyroid cancer. Biochemical testing in these patients often reveals subclinical hyperthyroidism. The effects of excess exogenous thyroid hormone on bone mineral density are conflicting, with some studies demonstrating a decrease in bone density [25, 26, 36–42] and others suggesting no effect [43–48]. The relationship between L-T4 suppressive therapy and biochemical markers of bone remodeling is similarly controversial [39, 40, 43–46, 49–51]. These discrepancies are explained, in part, by the heterogeneity of the patient populations studied, the heterogeneity of the underlying thyroid disease, differences in methodology and site of bone density determination, and by the small sample size and cross-sectional design of many of the studies. There is also great variability in the TSH assays (second- or thirdgeneration assays), the degree of suppression of TSH levels, the duration and type of thyroid hormone suppressive therapy, and the frequency of past endogenous hyperthyroidism in the patients studied. Finally, other factors which impact on BMD are not controlled for in many studies, such as calcium and vitamin D intake, physical activity, sun exposure, and ethnicity and race. Recent cross-sectional studies have suggested that exogenous subclinical hyperthyroidism does not significantly alter BMD when patients are compared with controls matched for age, menopausal status, height, and weight. Giannini et al. [45] evaluated 25 women (12 premenopausal) who were treated with suppressive doses of L-T4 for thyroid cancer for a mean of 7.6 years. TSH levels were

722 below normal in all patients. There was no difference in lumbar spine Z scores (DXA) between the premenopausal patients and controls nor between the postmenopausal patients and controls [45]. Similarly, Franklyn et al. [44] reported no effect of L-T4 suppressive therapy on femoral or vertebral BMD in pre- or postmenopausal women treated for a mean of 7.9 years for thyroid cancer. In contrast, Kung et al. [40] evaluated 34 postmenopausal Chinese women treated with suppressive doses of L-T4 for a mean of 12.2 years, after thyroidectomy for thyroid cancer. Patients had lower BMD (DXA) at the lumbar spine, femoral neck, Ward’s triangle, and trochanter compared to the control population. In addition, two patients in the treated group had a history of fracture (femoral neck, wrist). The differences among these studies may be accounted for by the longer duration of L-T4 use or by ethnic differences. In support of this, Ross et al. have demonstrated that premenopausal women treated with suppressive doses of L-T4 for more than 10 years had the most significant reduction in cortical BMD [36], suggesting that duration of suppressive therapy is an important factor in predicting bone mass. However, Franklyn et al. did not find any correlation between duration of L-T4 suppressive therapy and BMD [44]. Finally, two meta-analyses evaluating the effect of exogenous subclinical hyperthyroidism on bone mass revealed no significant change in bone mineral density in premenopausal women with suppressed TSH levels. However, a small but significant decrease in BMD was noted in postmenopausal women [52, 53]. Cross-sectional studies evaluating biochemical markers of bone remodeling in exogenous subclinical hyperthyroidism have been similarly conflicting. Total alkaline phosphatase activity has been either elevated [40] or unchanged [43, 44] in postmenopausal women with suppressed TSH levels. Elevated levels of bone-specific alkaline phosphatase (BAP) have been reported in both pre- and postmenopausal women taking suppressive doses of L-T4 compared to controls matched for age and menopausal status [45]. Elevated osteocalcin levels have been reported in pre- and postmenopausal women with suppressed TSH levels by some [39, 40, 51], but not all investigators [43, 46, 49]. Elevated urinary hydroxyproline levels have been reported in both premenopausal [45] and postmenopausal women with suppressed TSH levels [40, 51]. More specific urinary markers of bone resorption, such as urinary DPD and pyridinoline (PYD) excretion, have been demonstrated to be elevated in postmenopausal women with subclinical hyperthyroidism, but not in premenopausal women with similarly suppressed TSH levels [46, 54]. Urinary N-telopeptide and serum CrossLaps have not been elevated in either pre- or postmenopausal women

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on suppressive LT4 therapy [50]. Many studies have reported no significant change in PTH, calcitriol, or calcitonin levels [40, 45, 46]. There are few longitudinal studies assessing the impact of suppressive L-T4 therapy on BMD and markers of bone remodeling [46, 55, 56]. In one very small study involving older premenopausal women, there was no change in radial BMD assessed by SPA, but mean spinal BMD, assessed by DPA, decreased over a 3-year interval compared to age-matched controls [55]. This decrease may have been related to subtle changes in estrogen levels during the perimenopausal period, although all women continued to have regular menses during the study. Guo et al. recently evaluated longitudinal changes in BMD and markers of bone turnover in a homogeneous group of postmenopausal women on suppressive (n = 41) or replacement (n = 23) doses of levothyroxine, compared with agematched healthy postmenopausal women (n = 36) [56]. There was no difference in mean BMD (DXA: lumbar spine, femoral neck, or total body) or markers of bone turnover in any of the groups at baseline or at the end of the 2-year observation period. However, 2 years after the dose of L-T4 was decreased in the subgroup of patients with primary hypothyroidism who were inappropriately taking suppressive doses (n = 18), there was a significant increase in BMD in the lumbar spine and femoral neck and a significant decrease in all three markers of bone turnover compared with the women who remained on suppressive therapy for thyroid cancer (n = 23) and the healthy control population (Fig. 3). Therefore, it appears that suppressive L-T4 therapy can increase bone turnover, but bone markers in this study remained within the normal range for age. Furthermore, bone loss was not accelerated over a 2-year observation period. However, a reduction in L-T4 dose resulted in a decrease in bone turnover and an increase in BMD [56]. Taken together, the above data suggest that suppressive doses of L-T4 can increase bone remodeling in a subset of pre- and postmenopausal women with exogenous subclinical hyperthyroidism. Thus, these patients are predisposed to an excessive increase in bone resorption relative to bone formation, and subsequently are at a higher risk of decreased BMD. The risk of decreased BMD appears to be greater in postmenopausal women. However, BMD in patients on suppressive L-T4 therapy is not only dependent on the degree of TSH suppression, but perhaps also on the duration of use, as well as on lifestyle, hormonal, nutritional, genetic, and other factors. Consequently, patients on suppressive L-T4 do not necessarily develop osteopenia or osteoporosis. Cross-sectional studies of BMD and subclinical hyperthyroidism often fail to account for individual variation

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be reduced. If this is not possible, antiresorptive therapy should be considered. One cross-sectional study revealed that postmenopausal women taking both high-dose thyroid hormone and estrogen had BMD levels similar to women taking only estrogen. In contrast, women taking high-dose thyroid hormone without estrogen had lower BMD values in the spine, hip, and forearm [41]. These data suggest that estrogen protects against thyroid hormone-related bone loss. Intravenous pamidronate [57], but not calcitonin [58], has been shown to improve bone mass in patients taking suppressive doses of L-T4. There are insufficient data to support the treatment of all postmenopausal women on suppressive doses of L-T4 with antiresorptive agents. Measuring biochemical markers of bone remodeling may serve to identify an “at-risk” population who may benefit from antiresorptive therapy, although this strategy has not been rigorously evaluated in patients with subclinical hyperthyroidism.

E. Conclusion

Figure 3 The rate of change of serum osteocalcin (A), bone-specific alkaline phosphatase (B), and urinary NTX/Cr (C) in three groups of postmenopausal women taking L-thyroxine (Group 1 – hypothyroidism with normal TSH levels, Group 2 – hypothyroidism with previously suppressed TSH levels. The dose of L-thyroxine was reduced after the baseline measurement, and TSH levels normalized 1–4 months later. Group 3 – thyroid cancer with persistently suppressed TSH levels), and a control group with normal TSH levels (Group 4). The middle horizontal lines represent the mean values. The upper and lower horizontal lines represent the 95% confidence interval. The decrease in serum osteocalcin and urinary NTX/Cr in group 2 was significantly different than the other groups (P < 0.05). Groups with the same letters were not different from each other. Reproduced with permission from: Guo et al. [56] (1997). Longitudinal changes of bone mineral density and bone turnover in postmenopausal women on thyroxine. Clin. Endocrinol. 46: 301–307.

in BMD as a result of these other factors. A definitive understanding of the relationship between suppressive-dose L-T4 and BMD requires longitudinal studies of greater duration. Given the potential risk of bone loss, patients expected to continue long-term L-T4 suppressive therapy, especially patients with other risk factors for osteoporosis, require a careful evaluation of BMD and markers of bone remodeling. Postmenopausal patients and those with elevated markers of bone remodeling require long-term monitoring of BMD. If BMD declines, the dose of L-T4 should

Women with thyrotoxicosis secondary to Graves’ disease or thyroid autonomy have increased bone remodeling and decreased bone density, which is at least partially reversible with correction of thyroid function. Simultaneous treatment with bisphosphonates resulted in a more rapid normalization of BMD in one study [11]. Further studies are required to evaluate whether bisphosphonates confer additional longterm benefit, beyond that achieved with normalization of thyroid function. Subclinical hyperthyroidism in postmenopausal women with toxic multinodular goiter or solitary autonomous nodules appears to be a risk factor for osteoporosis and should be evaluated and treated in the context of other osteoporosis risk factors. Premenopausal women with similar thyroid dysfunction are at lower risk, possibly because of a protective effect of estrogen. Subclinical hyperthyroidism secondary to L-T4 suppressive therapy is associated with an increase in markers of bone remodeling and a decrease in BMD in some women. Again, postmenopausal women appear to be at greater risk than premenopausal women (Table II). Longitudinal studies are required to determine if antiresorptive therapy diminishes this risk in postmenopausal women. In the interim, patients requiring thyroid hormone replacement therapy should be treated with the lowest dose of L-T4 to maintain a normal TSH level. Patients requiring suppressive doses of L-T4 require monitoring of BMD. The utility of biochemical markers of bone remodeling in identifying the subset of patients at greatest risk for L-T4-related bone loss requires further investigation.

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BMD Bone turnover

Menopausal status Pre Post ↓ ↓ Increased Increased

Hyperthyroidism and Osteoporosis

Subclinical hyperthyroidism (endogenous) Menopausal status Pre Post No change ↓ Variable Variable

III. OSTEOPOROSIS SECONDARY TO HYPOGONADISM The effects of gonadal steroids on bone and mineral metabolism are relatively well known. Gonadal hormones, particularly estradiol, play a crucial role in the achievement of peak bone mass [59–61]. In addition, they are of fundamental importance in maintaining bone mass [60]. In later life, bone mass is dependent on both peak bone density attained at an earlier age as well as on the rate of bone loss. Therefore, hypogonadism at any time in life may increase the risk of osteoporosis and fractures. The beneficial effects of gonadal hormones on bone at the cellular level are not completely understood. In adults, the major effect of estrogen on bone remodeling is to decrease resorption rather than to affect bone formation [62]. Estrogen receptors and post-receptor signaling in response to estrogen administration have been demonstrated in osteoblasts, osteoclasts, and bone marrow stromal cells in culture. Osteoblasts and stromal cells produce cytokines, such as interleukin-1, interleukin-6, tumor necrosis factor, and macrophage-colony stimulating factor, that regulate osteogenesis. Estrogen appears to modulate the production of osteoclastogenic cytokines and hence prevents bone resorption [63, 64]. Estrogen deficiency, on the other hand, results in increased production of bone-resorbing cytokines leading to an increase in osteoclast activity and consequently greater bone resorption. In this section, we will review the consequences of hypogonadism on bone mineral density and indices of bone remodeling.

A. Turner’s Syndrome Turner’s syndrome is characterized by total or partial X chromosome monosomy, associated with gonadal dysgenesis and diminished or absent ovarian estrogen production [65]. Skeletal abnormalities, including short stature, cubitus valgus, delayed skeletal maturation, and osteoporosis, are commonly described in patients with Turner’s syndrome [66–69]. BMD values from 19–27%

Hyperthyroidism (levothyroxine treatment)

Subclinical hyperthyroidism (levothyroxine treatment)

Menopausal status Pre Post ↓ ↓ Insufficient Insufficient data data

Menopausal status Pre Post No change ↓ or No change Variable Variable

below control values have been reported in patients with Turner’s syndrome [67, 68, 70]. However, earlier studies are limited by less-sophisticated bone densitometry technology, heterogeneity in the patient population studied, and variability in treatment regimens, with respect to the timing and dose of estrogen or growth hormone therapy and use of anabolic steroids, such as oxandrolone, all of which may significantly impact BMD. Finally, an appropriate control population is difficult to identify. Girls and women with Turner’s syndrome have short stature and weigh less than other girls. BMD is an areal rather than a volumetric measurement, and therefore BMD is often underestimated in smaller individuals. Thus, height- and weight-matched controls are preferable to age-matched controls when comparing BMD values in Turner’s patients to those in a normal control population. Although earlier studies failed to account for the small stature of Turner’s patients, more recent studies address this issue [71, 72]. For example, Ross et al. [71] evaluated BMD, assessed by SPA and DPA, in 78 prepubertal girls with Turner’s syndrome. Bone density of the wrist was decreased compared to a prepubertal control group matched for chronological age, but wrist BMD was similar to the control group after adjusting for height in a multiple regression analysis. BMD of the spine did not differ between girls with Turner’s syndrome and the age-matched prepubertal control population. There are few longitudinal studies evaluating BMD in girls with Turner’s syndrome [72–74]. Shaw et al. [73] measured lumbar spine BMD sequentially for 2 years in 18 girls with Turner’s syndrome. BMD values, assessed by DXA, were compared with two separate reference data populations, an age-matched control group, and a control group matched for body weight and pubertal status. At baseline, mean standard deviation scores (SDS) were lower in the girls with Turner’s syndrome compared with age-matched controls. In contrast, mean SDS scores in Turner’s girls were normal when compared with the reference population matched for weight and pubertal status. Subsequently, 14 girls had BMD evaluations 2.5 years later. Only 11% of the 11–13-year-old girls had normal

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increments in BMD for age, whereas 63% of the 5–11-year-old girls had normal age-related increases in BMD. One possible hypothesis to account for the observation that bone accretion slows as girls reach pubertal age is that the dose, type, or pattern of exogenous estrogen replacement therapy, traditionally initiated at age 12 years, may not mimic normal puberty sufficiently to allow girls with Turner’s syndrome to achieve optimal peak bone density. In this study, there was no correlation between growth hormone or estrogen dose and increments in BMD. However, all girls received similar doses of ethinyl estradiol. An estrogen regimen which more closely replicates normal physiology may enable girls with Turner’s syndrome to achieve appropriate age-related increases in BMD. In contrast to prepubertal girls, adults with Turner’s syndrome have consistently demonstrated low BMD compared both with age-matched controls [75, 76] and with controls matched for height [67]. Davies et al. [67] evaluated lumbar spine BMD in 40 women with Turner’s syndrome. Mean vertebral BMD, assessed by DXA, was more than two standard deviations below the age-matched control population. Furthermore, when vertebral BMD was controlled for height and weight, it remained similarly low compared to the normal population. There was no relationship between duration of estrogen use and vertebral BMD in women with Turner’s syndrome. Taken together, the above data suggest that prepubertal girls with Turner’s syndrome have similar lumbar spine and wrist BMD values compared with age-matched girls when BMD values are adjusted for height and weight. In contrast, women with Turner’s syndrome have lower BMD than age-matched controls, regardless of correction for height and weight. Osteopenia in adult women with Turner’s syndrome appears to be related primarily to estrogen deficiency during the critical period of bone acquisition in the teenage years [67, 76, 77]. The observation that Turner’s patients who experience spontaneous vaginal bleeding have higher BMD values than typical Turner’s patients who require estrogen treatment to induce puberty supports this premise [67, 78]. In addition, girls who begin estrogen therapy before 11 years of age have been shown to have better radial bone mineral content than girls started on therapy after 12 years of age [69]. In cross-sectional studies, adult women with Turner’s syndrome on estrogen replacement therapy invariably have higher BMD values than untreated patients [68, 75–77, 79]. The results of small longitudinal studies evaluating the effects of estrogen therapy on radial BMC or BMD [68, 69, 80], or vertebral BMD [81, 82], suggest that BMC or BMD may increase slightly [69, 81] or remain unchanged [68, 80] with estrogen therapy. BMD does not typically reach a normal range for age and height, even with estrogen replacement therapy [69, 80, 82].

725 Because BMD does not completely normalize with estrogen treatment, some authors have hypothesized that an intrinsic bone defect related to the chromosomal abnormality, rather than estrogen deficiency, may contribute to bone loss in patients with Turner’s syndrome [68, 69, 83]. In support of this hypothesis is the finding that women with Turner’s syndrome had lower cortical BMD (as measured by DXA at the third radial site) compared with age-matched women with karyotypically normal premature ovarian failure [83]. However, cancellous bone (as measured by DXA of ultradistal forearm or DPA of LS) has not been demonstrated to be significantly different between the two groups [76, 83], suggesting that the chromosomal abnormality per se does not result in an intrinsic structural defect in cancellous bone [76]. Bone histomorphometric studies support the hypothesis that increased bone resorption secondary to estrogen deficiency is the primary cause of osteopenia in Turner’s patients. In 1974, Brown et al. [84] described bone morphology in eight girls (9–19 years of age) with Turner’s syndrome. None were treated with estrogen, despite the fact that many were past the normal age of puberty. The predominant histomorphometric feature was an increase in the percentage of bone surface undergoing resorption, with evidence of normal bone formation. Reports of biochemical markers of bone remodeling in Turner’s syndrome are conflicting. Saggese et al. [85] reported normal total alkaline phosphatase levels in 14 untreated females with Turner’s syndrome, aged 4.2–21.0 years. However, elevated total [75, 84] and bonespecific alkaline phosphatase [76] have been reported in five of eight untreated adolescent girls [84] and in estrogen-treated adults with Turner’s syndrome [75, 76]. In contrast, osteocalcin levels have been elevated in untreated adults with Turner’s syndrome, but not in estrogen-treated women [75, 76, 86]. In addition, Saggese et al. [85] have reported that serum osteocalcin concentrations increase after 1,25(OH)2 D administration in untreated patients (4.2–21.0 years of age), demonstrating that osteoblasts respond normally to physiologic stimulus in patients with Turner’s syndrome. Markers of bone resorption in Turner’s syndrome vary with age and estrogen therapy. Urinary hydroxyproline levels have been normal in prepubertal girls with Turner’s syndrome compared with controls matched for chronologic age [87] or bone age [88]. However, urinary hydroxyproline levels were elevated in girls older than 14 years and women, who were not receiving estrogen therapy, compared with age-matched controls [76, 87]. Stepan et al. [76] also reported increased urinary hydroxyproline levels in estrogen-treated adults with Turner’s syndrome. However, the values were lower than in the untreated group.

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Interestingly, urinary pyridinium cross-links, a more specific marker of bone resorption, has been demonstrated to be mildly elevated in prepubertal girls (4.6–14.8 years) with Turner’s syndrome compared with controls matched for bone age, suggesting that bone resorption may be increased even in some young girls with Turner’s syndrome [88]. However, whether rising pyridinium cross-link values in the oldest girls, who were entering puberty and not receiving estrogen therapy, accounted for the mild elevation was not addressed in the study. Taken together, these data suggest that markers of bone remodeling in Turner’s patients increase around the time of puberty, supporting the histomorphometric finding of increased bone resorption surfaces. Markers remain elevated in patients who are not treated with estrogen. However, since estrogen therapy does not consistently normalize markers, estrogen replacement alone may not be sufficient to reduce remodeling rates, at least in currently prescribed doses. Alternatively, other mechanisms of bone loss may be involved. Abnormal growth hormone (GH) secretory dynamics may also contribute to low bone mass in patients with Turner’s syndrome [89–91]. Although abnormal patterns of GH secretion have been observed in this syndrome, most patients are not GH deficient as classically defined by provocative testing [89]. Serum levels of IGF-I, as well as IGF binding protein-3 (IGFBP-3), have been reported to be normal in this syndrome. However, in vitro studies have shown that despite normal IGF-I plasma levels, the autocrine/paracrine actions of this growth factor are reduced [90]. Many therapeutic trials of supraphysiologic doses of recombinant human growth hormone have shown an increase in height velocity in girls with Turner’s syndrome [92]. One longitudinal study revealed improvement in volumetric cortical and cancellous BMD (as measured by phalangeal radiographic absorptiometry) in pubertal-aged girls treated with a combination of GH and estrogen. Three years after discontinuation of GH, during which time the dose of estrogen was increased, BMD increased and was not significantly different compared with healthy controls [74]. The study was limited by lack of a GH control group, and therefore the beneficial effects of GH vs estrogen on BMD could not be distinguished. Further studies are required to elucidate the relationship between low bone mass, growth factors, and cytokines in Turner’s syndrome.

B. Athletes with Amenorrhea The relationship between exercise and BMD is complex and incompletely understood. In general, exercise is associated with increased BMD. In women,

however, the beneficial effect of physical activity on bone mineral density can be lost if the amount of exercise is so intensive that it is associated with menstrual disturbances. The prevalence of amenorrhea among runners ranges from 25–50%. In contrast, only 12% of swimmers and cyclists have amenorrhea [93, 94]. Young girls who begin intensive athletic training before menarche often demonstrate delayed puberty, which may also be associated with lower bone mass [95]. The pathogenesis of exercise-induced amenorrhea is unclear. Risk factors include a prior history of menstrual irregularity, weight loss, and a low caloric intake. The absence of regular menses in exercising women is related to a disruption of the pulsatile release of gonadotropinreleasing hormone from the hypothalamus, resulting in low levels of gonadotropins and secondary reduction of estrogen and progesterone secretion [96]. Women who have missed 50% of their expected menstrual periods by 20 years of age are likely to have reduced bone mineral density as compared to normal controls. The most severe reductions in bone mass have been documented in the lumbar spine [96–99]. However, BMD is also lower at the femoral neck, trochanter, Ward’s triangle, and tibia (Fig. 4) [100]. In contrast, bone mineral density of the radius is generally normal [96–99]. Therefore, as in postmenopausal women, cancellous bone appears to be more severely affected by exercise-induced hypoestrogenism than cortical bone [101]. The effect of exercise on the growing skeleton also depends on the type of physical activity, the region of the skeleton involved [102], body weight, and dietary calcium intake. For example, gymnasts have been noted to have higher bone mineral density at the femoral neck than runners, despite similar past and current menstrual cycle patterns [103]. Similarly, Slemenda and Johnston [104] demonstrated that ice skaters with oligo- and amenorrhea had only a minimal (2%) decrement in bone mineral density compared to controls. Such data suggest that the mechanical forces associated with impact loading and muscular contraction during gymnastics and skating training have powerful osteogenic effects, which may counteract the deleterious effects of estrogen deficiency on the skeleton. Therefore, the type of exercise is an important factor in predicting bone loss in this young population. The effect of physical activity on serum concentrations of calcinotropic hormones and biochemical markers of bone remodeling has not been extensively studied. In one study of amenorrheic dancers, serum osteocalcin levels were elevated and inversely related to age, Tanner stage, bone density, and weight [105]. Osteocalcin declined with return of menses but not with estrogen therapy, suggesting that either exogenous therapy was inadequate or that

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Figure 4 Bone mineral density of amenorrheic (gray bars) and eumenorrheic (black bars) athletes (* P < 0.01). Reproduced with permission from: Rencken et al. [100] (1996). Bone density in amenorrheic athletes. JAMA 276: 239.

factors other than estrogen deficiency are operative in the increased bone turnover in this group. However, in another study, athletes who did not use oral contraceptives had higher levels of the bone resorption marker, serum beta-crosslaps (CTx), than athletes using oral contraceptives, which suggests that oral contraceptives might be somewhat protective of skeletal health in female athletes [106]. Few other studies have evaluated markers of bone turnover in exercising women. Zanker et al. [107] evaluated markers of bone remodeling in 33 female distance runners (mean age 27.2 years) in relation to their menstrual status. Serum levels of osteocalcin and BAP and urinary DPD were surprisingly lower in the amenorrheic runners compared with the eumenorrheic runners, indicative of reduced bone turnover. Markers of bone formation correlated with estradiol concentrations and BMI in the amenorrheic runners. The authors postulated that a chronic energy deficit resulted in decreased BMI, and a subsequent disruption of normal bone remodeling, in the amenorrheic runners. Similar metabolic changes are described in some patients with chronic eating disorders (see below). In summary, exercise-induced amenorrhea is a risk factor for osteoporosis. Although estrogen deficiency is an important cause of the bone loss, there may be other mechanisms for bone loss in these women which require further study, including inadequate caloric intake to match energy expenditure and type of excercise.

C. Eating Disorders Anorexia nervosa (AN) and bulimia affect 5–10% of women. The onset may be during early or late adolescence

or in the third or fourth decades of life. These eating disorders are often chronic and resistant to treatment, resulting in significant morbidity and even mortality. The reported prevalence of osteoporosis or osteopenia at one or more skeletal sites in patients with AN is 38% and 92%, respectively [108]. Osteopenia has been reported in both cancellous [109–113] and cortical [110, 114, 115] bone. However, cancellous bone loss is more frequent and more severe. While the etiology of osteoporosis in AN remains unclear, several metabolic consequences of AN may adversely affect skeletal mass. These include estrogen deficiency, secondary hyperparathyroidism due to low dietary calcium or vitamin D deficiency, endogenous cortisol excess, reduced IGF-I levels and protein-energy malnutrition. The interrelationship between body composition and regular menses may be particularly important for skeletal health in patients with AN. In one study, cancellous bone was relatively preserved in women with AN and normal menstrual cycles, compared to women with AN and amenorrhea [116]. Although BMI was similarly low in both groups, eumenorrheic subjects had a higher percent body fat and higher leptin levels, highlighting the importance of fat mass in the preservation of normal menstrual function, and therefore cancellous BMD, in underweight women. Histomorphometric studies in patients with anorexia report conflicting results [117, 118]. Kaplan et al. [117] reported a markedly reduced rate of bone formation and a concomitant increase in parameters of bone resorption in one patient with anorexia. In general, the level of bone remodeling activity was reduced, suggesting inactive or low-turnover osteoporosis. In contrast, Joyce et al. [118] reported a high turnover pattern in two anorectic women and three patients with other types of eating disorders.

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Data on markers of bone remodeling in anorectic women are similarly conflicting. Osteocalcin, a specific marker of osteoblast function, has been noted to be very low in some studies of women with AN [119, 120] (Fig. 5). Furthermore, a significant correlation between serum osteocalcin and body mass index has been demonstrated, suggesting a link between nutritional status and decreased rates of bone formation [120]. In contrast, other reports have shown normal [112, 115] or elevated [121] levels of osteocalcin. With respect to markers of bone resorption, urinary hydroxyproline was normal in one study of anorectic patients [112]. However, measurements of more specific markers of bone resorption, such as deoxypyridinoline and N-telopeptide excretion, have revealed high-normal or increased levels [120, 122, 123], which were inversely correlated with bone mineral density (Fig. 5) [120]. Moreover, in the latter study [120], serum osteocalcin was very low, thus supporting the hypothesis that uncoupling of bone remodeling with decreased bone formation and increased resorption contributes to bone loss in some young women with anorexia. Other factors may also contribute to metabolic bone disease in patients with anorexia. Serum vitamin D concentrations are highly variable in patients with eating disorders. Serum levels can be affected by reduced dietary intake and diminished hepatic synthesis or binding capacity of vitamin D-binding protein in patients with estrogen deficiency. Total 25-OHD levels are normal in the majority of AN patients [112, 114, 117, 121]. Elevated serum and urinary cortisol levels are common in women with AN [111, 112, 124, 125]. In a group of 53 women with eating disorders, hypercortisolism was present in 55% of the anorectic patients but just 6% of the bulimic women [112]. Hypercortisolism results from both increased production and decreased clearance of cortisol, secondary to enhanced activity of the hypothalamicpituitary-adrenal axis. High cortisol levels may contribute

to the severe bone loss seen in patients with AN. In this regard, Biller et al. reported elevated urinary cortisol levels in 26 amenorrheic women with AN compared with the control group [111]. Moreover, urinary cortisol levels correlated inversely with vertebral bone density, supporting the hypothesis that endogenous hypercortisolism is important in the development of osteopenia in women with AN. In addition to hypercortisolism, elevated GH and suppressed IGF-I levels have been described in patients with AN [126]. It has been hypothesized that prolonged nutritional deprivation in AN induces down-regulation of the GH receptor or its postreceptor processing, resulting in reduced production of IGF-I and elevated GH levels. IGF-I is a nutritionally dependent bone factor, which stimulates collagen formation by osteoblasts. Therefore, deficiency of IGF-I may contribute to decreased bone formation and bone loss in women with AN. In this regard, Klibanski et al. have demonstrated that IGF-I levels were predictive of the change in vertebral bone mass in untreated patients with AN [113]. In addition, Grinspoon et al. evaluated the effects of short-term recombinant IGF-I (rhIGF-I) administration on bone turnover and BMD in women with AN [120, 127]. There was a 1.8% increase in LS BMD in subjects treated with IGF-I and oral contraceptives for 9 months, whereas there was a 1% decrease in LS BMD in the untreated control group [127]. Markers of bone remodeling were stimulated in a dose-dependent manner. The highest dose of rhIGF-I stimulated markers of both bone formation (osteocalcin, PICP) and resorption (DPD), whereas the lower dose stimulated only a single marker of bone formation (PICP) [120]. These data support the hypothesis that low bone mass in AN is related, at least in small part, to disturbances in the GH–IGF-I axis. Medical treatment with estrogen, including oral contraceptives, or IGF-I has not been shown to normalize BMD. Weight gain has a positive impact on bone mass [113]. One small study suggested a beneficial effect of Actonel

Figure 5 Comparison of baseline bone turnover in women with anorexia nervosa (n = 22) and normal controls; (A) osteocalcin (* P < 0.001); (B) deoxypyridinoline (* P < 0.001); (C) N-telopeptide (*P < 0.01). Reproduced with permission from: Grinspoon et al. [120] (1996). Effect of short-term recombinant human insulinlike growth factor-1 administration on bone turnover in osteopenic women with anorexia nervosa. JCEM 81: 3866.

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on LS BMD in ten women with AN [122]. However, bisphosphonates are not approved for the treatment of premenopausal women with low bone mass, unless low bone mass is associated with glucocorticoid use. Further studies are required in this patient population to determine safety and efficacy. In summary, bone loss associated with anorexia nervosa is multifactorial in etiology and may be related to estrogen deficiency, vitamin D deficiency, cortisol excess, reduced IGF-I levels, and malnutrition. Biochemical markers of bone resorption are usually elevated. Markers of bone formation may be elevated or depressed and may vary with the severity or duration of the disease. It is unclear whether markers of bone remodeling will be useful in identifying patients who are most at risk for osteoporosis and future fracture.

D. Hyperprolactinemia Hyperprolactinemia commonly occurs secondary to pituitary or hypothalamic diseases, secondary to certain medications, such as antipsychotic drugs, and in some physiological conditions, such as pregnancy and lactation. High levels of prolactin inhibit gonadotropin-releasing hormone (GnRH) and consequently cause hypogonadotropic hypogonadism. As many as 25–30% of premenopausal women with amenorrhea have hyperprolactinemia [128]. Decreased bone mass has been reported in patients with hyperprolactinemia [129–133]. Cross-sectional studies have demonstrated BMD values 4–23% lower at appendicular sites [133–135] and 8–23% lower at axial sites [134–136], compared with age-matched controls. The importance of hypogonadism in the pathogenesis of bone loss in these patients has been well documented [130, 134–137]. Cicarreli et al. demonstrated that there was a positive correlation between BMC and estrogen levels in a group of women with hyperprolactinemia [131]. Furthermore, Kayath et al. reported that men and women with a history of prolactinoma and hypogonadism for more than 10 years had lower BMD values than patients who were more recently diagnosed [130]. Similarly, Biller et al. reported lower BMD in hyperprolactinemic women with amenorrhea than in those with hyperprolactinemia and oligoamenorrhea or normal menses [133]. These findings suggest that the severity of bone loss in patients with hyperprolactinemia is related to the presence and duration of hypogonadism, emphasizing the significant role of estrogen deficiency in the pathogenesis of osteoporosis associated with this condition. Few studies have addressed the reversibility of osteopenia in hyperprolactinemic amenorrheic patients [133, 138].

729 In Biller’s study [133], vertebral BMD, assessed by computerized tomography, declined progressively in untreated hyperprolactinemic amenorrheic women followed for a mean of 1.7 years. In contrast, 56% of treated patients, who had resumption of menstrual function, had improvement in BMD. The authors concluded that menstrual function is the most important predictor of progressive bone loss in hyperprolactinemic women. However, 44% of women did not have an increase in BMD, despite restoration of their menstrual function, suggesting that other factors may play a role in the bone loss seen in these women. One such factor may be PTH-related peptide (PTHrP), a potent stimulator of bone resorption. A significant elevation in serum levels of PTHrP has been reported in patients with hyperprolactinemia [139, 140]. Kovacs and Chik evaluated 33 lactating women and 16 patients with prolactin-secreting pituitary adenomas [140]. Both groups had higher levels of PTHrP than sex- and age-matched controls. Repeat measurements in four patients after therapy with either bromocriptine, surgery, or octreotide revealed a decrease in PTHrP levels. Furthermore Stiegler et al. have reported lower vertebral BMD Z scores in hyperprolactinemic patients with high PTHrP levels compared with hyperprolactinemic patients with normal levels [141]. There was a significant negative correlation between PTHrP and BMD Z scores. In addition, patients with higher PTHrP levels had significantly higher mean serum calcium, total alkaline phosphatase, and osteocalcin levels and lower mean serum phosphorous levels. Urinary calcium and phosphorus excretion were also higher in the group with the high PTHrP levels. Therefore, PTHrP may play a role in bone loss associated with hyperprolactinemia. Few studies have evaluated biochemical markers of bone remodeling in patients with hyperprolactinemia. In Ciccarelli study’s, there was no difference in urinary hydroxyproline levels between the hyperprolactinemic women with or without amenorrhea [131]. Similarly, total alkaline phosphatase levels have been reported to be normal in several studies [130, 132, 133, 136]. However, measurements of more specific markers of bone remodeling, such as osteocalcin and N-telopeptide, have revealed important changes. For example, Sartorio et al. reported low osteocalcin levels in 29 hyperprolactinemic women with prolactinomas compared with age-matched controls [142]. After 12 months of therapy with dopamine agonists, osteocalcin levels significantly increased and prolactin levels decreased. Similarly, Di Somma et al. recently reported low osteocalcin levels in 20 men with pituitary adenoma, compared with controls matched for age and BMI. In contrast, urinary N-telopeptide levels were increased [132]. BMD of the lumbar spine correlated positively with serum

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osteocalcin and inversely with urinary N-telopeptide excretion. Furthermore, a significant inverse correlation was observed between serum prolactin and osteocalcin levels. After 18 months of treatment with dopamine agonists, osteocalcin levels significantly increased and N-telopeptide levels decreased [132]. Taken together, these findings suggest that an uncoupling of bone remodeling, with an increase in bone resorption and a decrease in bone formation, occurs in patients with hyperprolactinemia. Further studies are needed to confirm these findings.

E. Conclusion In conclusion, patients with hypogonadism of any cause are at risk for developing osteoporosis. The decrease in bone mass is primarily due to deficiency of gonadal steroids. However, disturbances in nutrition and other hormonal factors, such as IGF-I, PTHrP, and cortisol may also contribute. The utility of biochemical indices of bone remodeling in the evaluation and treatment of patients with osteoporosis secondary to hypogonadism remains unclear. Further evaluation within each patient subgroup, particularly longitudinal studies, are required to determine whether markers of bone turnover predict rates of bone loss and fracture in affected individuals.

IV. ANTICONVULSANT DRUGS AND OSTEOPOROSIS Metabolic bone disease may complicate the management of adults and children receiving several of the commonly used anticonvulsant drugs [143, 144]. Diphenylhydantoin, phenobarbital, carbamazepine, and sodium valproate have all been implicated. In the past, anticonvulsant-induced bone disease typically presented as overt osteomalacia or rickets, with fractures, proximal muscle weakness, frank hypocalcemia, and hypophosphatemia [145, 146]. In the more common modern presentation, patients commonly present with an asymptomatic decrease in bone mineral density [147–152]. Importantly, however, fracture rates are higher than expected in epilepsy patients [153]. Institutionalized patients with epilepsy have a 4.3-fold increased risk of all fractures and a 3.2-fold increase for fractures not related to seizures [154]. However, fracture risk is also increased in community-dwelling patients. A recent study found that that the risk of fracture in patients with epilepsy was double that in controls; only 34% of these fractures were related to seizures [155]. The fractures may be related to the seizures themselves or to the increased incidence of falls in patients with epilepsy. The Study of

Osteoporotic Fractures found that community-dwelling women over age 65 who used antiepileptic drugs were 75% more likely to fall than women who did not require such therapy [156]. However, the fractures may also be related to low bone mass associated with the use of various antiepileptic drugs. Antiepileptic drugs likely affect bone and mineral metabolism by multiple mechanisms [157]. These mechanism have been reviewed recently [157]. Traditionally, abnormalities in bone and mineral metabolism in patients with epilepsy have been ascribed to effects of certain antiepileptic drugs (AEDs) that induce the hepatic cytochrome P450 system and cause abnormalities in the vitamin D–parathyroid hormone endocrine system. Diphenylhydantoin, phenobarbital, and carbamazepine stimulate hepatic mixed function oxidase activity and therapy with these drugs is associated with increased degradation of vitamin D sterols [158–160]. In contrast, sodium valproate does not induce the cytochrome P450 system and therefore is not thought to affect vitamin D metabolism directly. However, this drug has been associated with low bone density and increased bone turnover in adults with epilepsy and may affect bone and mineral metabolism through other mechanisms [151]. The effects of anticonvulsant drugs on serum 25-hydroxyvitamin D concentrations are unclear. Hahn and others have shown that anticonvulsant drug administration is associated with increases in the disappearance of vitamin D and 25-hydroxyvitamin D from the circulation and the appearance of inactive polar metabolites of vitamin D in the bile and urine [158–161]. In older studies, serum levels of 25-OHD were lower in longstanding epilepsy [159, 160, 162–167]. A more recent Finnish study also found serum levels of 25-OHD to be lower in patients taking various combinations of diphenylhydantoin, phenobarbital, and carbamazepine [152]. In contrast, other studies found serum 25-OHD levels in patients on diphenylhydantoin to be no different from normal controls [148, 168–170]. These discrepancies may be related to geographic differences in the study populations, since the studies with low 25-hydroxyvitamin D levels were conducted in more northern latitudes where there is less sunlight exposure. Thus, inadequate vitamin D stores may develop when taking these drugs, particularly when patients are from northern climates or institutionalized, and access to ultraviolet light may be diminished. In addition to the actions of anticonvulsant drugs upon hepatic vitamin D metabolism, direct effects upon cellular metabolism have been described. Diphenylhydantoin, which interferes with cation transport in many tissues, directly inhibits intestinal calcium transport [171], and together with phenobarbital, has been shown to inhibit

Chapter 43 Secondary Osteoporosis

vitamin D-mediated calcium absorption [172]. Furthermore, Hahn and Halstead demonstrated that higher levels of vitamin D metabolites are required to maintain normal intestinal calcium absorption in patients taking anticonvulsant drugs [173]. In addition, both phenobarbital and diphenylhydantoin inhibit parathyroid hormone-mediated bone resorption in vitro which could cause resistance to parathyroid hormone and secondary increases in serum parathyroid hormone [174, 175]. Diphenylhydantoin, which appears to exert this effect by inhibition of hormonesensitive adenylate cyclase activity, is the more potent of the two drugs. Diphenylhydantoin also directly inhibits collagen synthesis and lysozomal enzyme release by bone explants [176]. Finally, diphenylhydantoin may affect estrogen metabolism. One case has been reported of a 17-year-old epileptic boy who had been treated with multiple anticonvulsants; the boy had very low bone mass and open epiphyses and a serum estrogen below 10 pg/ml. After 10 months of treatment with estrogen, he had a significant increase in bone density and bone age [177]. In contrast, another study reported elevated total and bioavailable serum estradiol concentrations in men with epilepsy receiving diphenylhydantoin compared to those receiving other antiepileptic drugs [178]. Two histomorphometric studies by Mosekilde and Melsen [179] and Weinstein et al. [169] have contributed in a major way to our current understanding of the skeletal effects of anticonvulsant drugs, establishing that these drugs are generally associated with increased bone turnover rather than osteomalacia. Mosekilde and Melsen evaluated a group of 20 adults all treated with diphenylhydantoin for at least 10 years, some of whom were also taking other anticonvulsant drugs [179]. Compared to a group of normal controls, these patients had significantly reduced serum and urinary calcium levels and elevated serum total alkaline phosphatase activity. Analysis of transiliac crest bone biopsies revealed normal cancellous bone volume, and increases in osteoid volume, active osteoid surface, mineralizing surface, and resorption surface. Although osteoid seams were slightly thicker than a control population, the mineral apposition rate was normal. Weinstein et al. [169] evaluated bone biopsies in 20 patients who had taken anticonvulsant drugs for 18 years. They observed decreased serum ionized calcium and increased parathyroid hormone levels and low cortical bone mineral density. Although cancellous bone volume was normal, cortical bone was thin and porous. Osteoid volume and surface were increased, osteoid width was normal, and the mineralization rate was normal or increased. Their findings were essentially in agreement with those of Mosekilde and Melsen [179], thus establishing anticonvulsant bone disease as a disorder of high remodeling rather than abnormal mineralization.

731 Low 25-hydroxyvitamin D levels could cause decreased intestinal calcium absorption and secondary hyperparathyroidism, as well as the increased bone remodeling observed by Weinstein [169] and Mosekilde [179]. However, routine serum measurements of indices of bone and mineral metabolism are frequently unremarkable in patients taking anticonvulsant drugs. Biochemical evaluation may reveal normal or minimally depressed serum calcium and phosphorus levels and serum total alkaline phosphatase activity may be normal or only slightly elevated. Serum 25-hydroxyvitamin D levels are frequently reduced in anticonvulsant-treated patients and there may be mild elevations of parathyroid hormone. Serum 1,25-dihydroxyvitamin D levels may be normal, high, or low. Markers of bone formation such as osteocalcin [152, 180, 181], serum total [180, 182], and bone-specific alkaline phosphatase and procollagen carboxyterminal peptide [181] have been reported to be elevated in children and adults taking anticonvulsant drugs. With respect to markers of resorption, Ohishi et al. [183] documented that urinary hydroxyproline, pyridinoline, and deoxypyridinoline excretion were increased in 15 premenopausal women taking anticonvulsant drugs compared with 211 healthy age- and sex-matched controls. Elevated urinary excretion of hydroxyproline [180, 182] and cross-linked carboxyterminal peptide of type I collagen have also been reported in patients on antiepileptic drugs. Elevated markers of bone turnover have also been reported in studies of adults [184] and children [185] on carabamazepine monotherapy. Thus, most available data suggest that biochemical markers of bone turnover are elevated in adults and children on anticonvulsant drugs, consistent with the histomorphometric studies that have established this disorder as a state of increased bone remodeling [169, 179]. In contrast, a recent study of community-dwelling premenopausal women with well-controlled epilepsy did not reveal elevated bone turnover markers [170]. Data on bone mineral density in children and adults taking anticonvulsant drugs are also conflicting. Adults receiving anticonvulsant drugs have been reported to have decreased bone density at multiple sites, including ribs, femoral neck, and spine [147, 148, 150–152]. A recent study of 44 epileptics, on medication for more than 5 years, found significantly decreased BMD (by DEXA) at the hip and lumbar spine [149]. Valimaki et al. measured spine and hip bone density in 38 men and women between the ages of 20 and 49 who were taking diphenylhydantoin, carbamazepine, or both. BMD was abnormal only in women and only at the hip [152]. Sheth and colleagues evaluated both axial and appendicular bone mass by DEXA in 26 children who had been taking sodium valproate or carbamezepine for more than 18 months [186]. While BMD

732 was normal in patients on carbamezepine, those receiving sodium valproate had reduced BMD at both the lumbar spine and radius. Higher rates of osteopenia and osteoporosis are often [147, 148, 150–152], but not always [170], found in adults receiving anticonvulsant drugs. In summary, bone turnover is frequently elevated in patients taking anticonvulsant drugs and bone mass is often, but not always, reduced. Risk factors for the development of anticonvulsant bone disease include high dose, multiple drug regimens, long duration of therapy, institutionalization, vitamin D deficiency either due to inadequate dietary intake or reduced sunlight exposure, physical inactivity, use of ketogenic diets or acetazolamide to induce chronic metabolic acidosis, and concomitant therapy with other drugs that induce hepatic enzymes (Table III). Anticonvulsant bone disease should be sought in individuals with a history of chronic long-term therapy and one or more of these risk factors. The routine biochemical evaluation may reveal hypocalcemia, hypophosphatemia, and increased alkaline phosphatase activity or may be normal. Urinary calcium excretion and serum 25-hydroxyvitamin D levels may be reduced and there may be mild elevations of serum parathyroid hormone. Both serum osteocalcin concentrations and urinary pyridinium crosslink excretion may be increased. However, any or all of these biochemical parameters may be normal and decreased bone mineral density may be the only manifestation. Alternatively, bone density may be normal in the face of clear-cut biochemical abnormalities. In difficult cases, bone biopsy may be helpful in establishing the diagnosis. Management of patients on chronic anticonvulsant drugs should include routine prophylaxis with supplementation with at least 400 IU and up to 2000 IU of vitamin D daily [187]. This is particularly important in the case of elderly or institutionalized patients. Although Barden et al. have demonstrated that 400 IU will prevent bone loss in institutionalized adults [188], Collins et al. have shown that much higher doses (400–4000 IU) may be necessary

Table III. Risk Factors for Bone Disease in Patients Receiving Anticonvulsant Drugs Long-term therapy High-dose therapy Multiple drug regimens Reduced physical activity Low vitamin D intake Chronic illness Elderly or institutionalized patients Limited exposure to sunlight Regimens that include drugs that induce hepatic cytochrome P450 enzymes

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to normalize serum 25-OHD levels in both institutionalized and non-institutionalized patients [189]. The common side effects of vitamin D therapy – hypercalciuria and hypercalcemia – are unlikely to occur at the relatively small doses (usually < 2000 IU) that have been shown to be effective. Treatment of established anticonvulsant bone disease with 2000–4000 IU of vitamin D daily has been demonstrated to result in improved calcium absorption, decreases in parathyroid hormone and urinary hydroxyproline excretion, and increases in bone mineral content [188]. A calium intake of 600–1000 mg/day should also be ensured [187]. All of these approaches should prove relatively cost-effective, particularly when contrasted with medical costs associated with fractures. Treatment of low bone density with bisphosphonates may be required, but only if serum 25-OHD and parathyroid hormone concentrations are normal, and there is no clinical or biochemical evidence of osteomalacia. Routine use of bisphosphonates in patients receiving long-term anticonvulsant drug therapy cannot be recommended at present [187].

References 1. Allain, T. J., and McGregor, A. M. (1993). Thyroid hormones and bone. J. Endocrinol. 139, 9–18. 2. Mosekilde, L., Flemming, M., Bagger, J. P., Myhre-Jensen, O., and Sorensen, N. S. (1976). Bone changes in hyperthyroidism: interrelationships between bone morphometry, thyroid function and calciumphosphorous metabolism. Acta. Endocrinologica. 85, 515–525. 3. Mosekilde, L., Eriksen, E. F., and Charles, P. (1990). Effects of thyroid hormones on bone and mineral metabolism. Endocrinol. Metab. Clin. N. Am. 19, 35–63. 4. Eriksen, E. F., Mosekilde, L., and Melsen, F. (1985). Trabecular bone remodeling and bone balance in hyperparathyroidism. Bone 6, 421–428. 5. Bisbocci, D., Gallo, V., Damiano, P., Sidoli, L., Cantoni, R., Aimo, G., Priolo, G., Pagni, R., and Chiandussi, L. (1996). Spontaneous release of interleukin 1b from human blood monocytes in thyrotoxic osteodystrophy. J. Endocrinol. Invest. 16, 511–515. 6. Lakatos, P., Foldes, J., Horvath, C., Kiss, L., Tatrai, A., Takacs, I., Tarjan, G., and Stern, P. H. (1997). J. Clin. Endocrinol. Metab. 82, 78–81. 7. Abe, E., Marians, R. C., Yu, W., Wu, X., Ando, T., Yanan, L., Iqbal, J., Eldeiry, L., Rajendren, G., Blair, H. C., Davies, T. F., and Zaidi, M. (2003). TSH is a negative regulator of skeletal remodeling. Cell. 115, 151–162. 8. Linde, J., and Friis, T. (1979). Osteoporosis in hyperthyroidism estimated by photon absorptiometry. Acta. Endocrinolgica. 91, 437–448. 9. Krolner, B., Jorgensen, J. V., and Nielsen, S. P. (1983). Spinal bone mineral content in myxoedema and thyrotoxicosis. Effects of thyroid hormone(s) and anti-thyroid treatment. Clin. Endocrinol. 18, 439–446. 10. Diamond, T., Vine, J., Smart, R., and Butler, P. (1994). Thyrotoxic bone disease in women: a potentially reversible disorder. Ann. Intern. Med. 120, 8–11. 11. Lupoli, G., Nuzzo, V., Di Carlo, C., Affinito, P., Vollery, M., Vitale, G., Cascone, E., Arlotta, F., and Nappi, C. (1996). Effects of alendronate on bone loss in pre- and postmenopausal hyperthyroid women treated with methimazole. Gynecol. Endocrinol. 10, 343–348.

Chapter 43 Secondary Osteoporosis 12. Jodar, E., Munoz-Torres, M., Escobar-Jimenez, F., Quesada, M., Luna, J. D., and Olea, N. (1997). Antiresorptive therapy in hyperthyroid patients: longitudinal changes in bone and mineral metabolism. J. Clin. Endocrinol. Metab. 82, 1989–1994. 13. Rosen, C. J., and Adler, R. A. (1992). Longitudinal changes in lumbar bone density among thyrotoxic patients after attainment of euthyroidism. J. Clin. Endocrinol. Metab. 75, 1531–1534. 14. Fraser, S. A., Anderson, J. B., Smith, D. A., and Wilson, G. M. (1971). Osteoporosis and fractures following thyrotoxicosis. Lancet 1, 981–983. 15. Toh, S. H., Claunch, B. C., and Brown, P. H. (1985). Effect of hyperthyroidism and its treatment on bone mineral content. Arch. Intern. Med. 145, 883–886. 16. Lee, M. S., Kim, S. Y., Lee, M. C., Cho, B. Y., Lee, H. K., Koh, C., and Min, H. K. (1990). Negative correlation between the change in bone mineral density and serum osteocalcin in patients with hyperthyroidism. J. Clin. Endocrinol. Metab. 70, 766–770. 17. Nagasaka, S., Sugimoto, H., Nakamura, T., Kusaka, I., Fujisawa, G., Sakuma, N., Tsuboi, Y., Fukuda, S., Honda, K., Okada, K., Ishikawa, S., and Saito, T. (1997). Antithyroid therapy improves bony manifestations and metabolic markers in patients with Graves’ thyrotoxicosis. Clin. Endocrinol. 47, 215–221. 18. Legovini, P., De Menis, E., Rin, G. D., Roiter, I., Breda, F., Artuso, V., Di Virgilio, R., and Conte, N. (1994). Increased serum levels of carboxyterminal propeptide of type 1 collagen (PICP) in hyperthyroidism. Horm. Metab. Res. 26, 334–337. 19. MacLeod, J. M., McHardy, K. C., Harvey, R. D., Duncan, A., Reid, I. W., Bewsher, P. D., and Robins, S. P. (1993). The early effects of radioiodine therapy for hyperthyroidism on biochemical indices of bone turnover. Clin. Endocrinol. 38, 49–53. 20. Ohishi, T., Kushida, K., Takahashi, M., Kawana, K., Yagi, K., Kawakami, K., Horiuchi, K., and Inoue, T. (1994). Urinary bone resorption markers in patients with metabolic bone disorders. Bone 15, 15–20. 21. Garnero, P., Vassy, V., Bertholin, A., Riou, J. P., and Delmas, P. D. (1994). Markers of bone turnover in hyperthyroidism and the effects of treatment. J. Clin. Endocrinol. Metab. 78, 955–959. 22. Garnero, P., Gineyts, E., Riou, J. P., and Delmas, P. D. (1994). Assessment of bone resorption with a new marker of collagen degradation in patients with metabolic bone disease. J. Clin. Endocrinol. Metab. 79, 780–785. 23. Garrel, D. R., Delmas, P. D., Malaval, L., and Tourniaire, J. (1986). Serum bone gla protein: a marker of bone turnover hyperthyroidism. J. Clin. Endocrinol. Metab. 62, 1052–1055. 24. Franklyn, J., Betteridge, J., Holder, R., Daykin, J., Lilley, J., and Sheppard, M. (1994). Bone mineral density in thyroxine treated femals with our without a previous history of thyrotoxicosis. Clin. Endocrinol. 41, 425–432. 25. Adlin, E. V., Maurer, A. H., Marks, A. D., and Channick, B. J. (1991). Bone mineral density in postmenopausal women treated with L-thyroxine. Am. J. Med. 90, 360–366. 26. Greenspan, S. L., Greenspan, F. S., Resnick, N. M., Block, J. E., Friedlander, A. L., and Genant, H. K. (1991). Skeletal integrity in premenopausal and postmenopausal women receiving long-term L-thyroxine therapy. Am. J. Med. 91, 5–14. 27. Mudde, A. H., Reijnders, F. J. S., and Kruseman, A. C. N. (1992). Peripheral bone density in women with untreated multinodular goitre. Clin. Endocrinol. 37, 35–39. 28. Faber, J., Overgaard, K., and Jarlow, A. (1992). Bone mineral content in premenopausal women with nontoxic goiter and reduced serum TSH. Proceedings from the American Thyroid Association’s 66th meeting, Rochester, MN. Abstract 1. 29. Foldes, J., Tarjan, G., Szathmari, M., Varga, F., Krasznai, I., and Horvath, C. S. (1993). Bone mineral density in patients with endogenous subclincal hyperthyroidism: Is this thyroid status a risk factor for osteoporosis? Clin. Endocrinol. 39, 521–527.

733 30. Faber, J., Jensen, W., Petersen, L., Nygaard, B., Hegedus, L., and Siersbaek-Nielsen, K. (1998). Normalization of serum thyrotrophin by means of radioiodine treatment in subclinical hyperthyroidism: effect on bone loss in postmenopausal women. Clin. Endocrinol. 48, 285–290. 31. Faber, J., Perrild, H., and Johansen, J. S. (1990). Bone gla protein and sex-hormone-binding globulin in nontoxic goiter: parameters for metabolic status at the tissue level. J. Clin. Endocrinol. Metab. 70, 49–55. 32. Kumeda, Y., Inaba, M., Tahara, H., Kurioka, Y., Ishikawa, T., Morii, H., and Nishizawa, Y. (2000). Persistent increase in bone turnover in Graves’ patients with subclinical hyperthyroidism. J. Clin. Endocrinol. Metab. 85, 4157–4161. 33. Kisakol, G., Kaya, A., Gonen, S., and Tunc, R. (2003). Bone and calcium metabolism in subclinical autoimmune hyperthyroidism and hypothyroidism. Endocrine J. 50, 657–661. 34. Foldes, J., Lakatos, P., Zsadanyi, J., and Horvath, C. (1997). Decreased serum IGF-I and dehydroepiandrosterone sulphate may be risk factors for the development of reduced bone mass in postmenopausal women with endogenous subclinical hyperthyroidism. Eur. J. Endocrinol. 136, 277–281. 35. Fallon, M. D., Perry, H. M., Bergfeld, M., Droke, D., Teitelbaum, S. L., and Avioli, L. V. (1983). Exogenous Hyperthyroidism with osteoporosis. Arch. Intern. Med. 143, 442–444. 36. Ross, D. S., Neer, R. M., Ridgway, E. C., and Daniels, G. H. (1987). Subclinical hyperthyroidism and reduced bone density as a possible result of prolonged suppression of the pituitary-thyroid axis with L-thyroxine. Am. J. Med. 82, 1167–1170. 37. Paul, T. L., Kerrigan, J., Kelly, A. M., Braverman, L. E., and Baran, D. T. (1988). Long-term L-thyroxine therapy is associated with decreased hip bone density in premenopausal women. JAMA 259, 3137–3141. 38. Staff, G. M., Harris, S., Sokoll, L. J., and Dawson-Hughes, B. (1990). Accelerated bone loss in hypothyroid patients overtreated with L-thyroxine. Annals. Int. Med. 113, 265–269. 39. Diamond, T., Nery, L., and Hales, I. (1990). A therapeutic dilemma: suppressive doses of thyroxine significantly reduce bone mineral measurements in both premenopausal and postmenopausal woman with thyroid carcinoma. J. Clin. Endocrinol. Metab. 72, 1184–1188. 40. Kung, A. W. C., Lorentz, T., and Tam, S. C. F. (1993). Thyroxine suppressive therapy decreases bone mineral density in post-menopausal women. Clin. Endocrinol. 39, 535–540. 41. Schneider, D. L., Barrett-Connor, E. L., and Morton, D. J. (1994). Thyroid hormone use and bone mineral density in elderly women. JAMA 271, 1245–1249. 42. Sijanovic, S., and Karner, I. (2001). Bone loss in premenopausal women on long-term suppressive therapy with thyroid hormone. Medscape Womens Health 6, 3–7. 43. Gam, A. N., Jensen, G. F., Hasselstrom, K., Olsen, M., and Nielsen, K. S. (1991). Effect of thyroxine therapy on bone metabolism in substituted hypothyroid patients with normal or suppressed levels of TSH. J. Endocrinol. Invest. 14, 451–455. 44. Franklyn, J. A., Betteridge, J., Daykin, J., Holder, R., Oates, G. D., Parle, J. V., Lilley, J., Heath, D. A., and Sheppard, M. C. (1992). Longterm thyroxine treatment and bone mineral density. Lancet 340, 9–13. 45. Giannini, S., Nobile, M., Sartori, L., Binotto, P., Ciuffreda, M., Gemo, G., Pelizzo, M. R., D’Angelo, A., and Crepaldi, G. (1994). Bone density and mineral metabolism in thyroidectomized patients treated with long-term L-thyroxine. Clin. Sci. 87, 593–597. 46. Garton, M., Reid, I., Loveridge, N., Robins, S., Murchison, L., Beckett, G., and Reid, D. (1994). Bone mineral density and metabolism in premenopausal women taking L-thyroxine replacement therapy. Clin. Endocrinol. 41, 747–755. 47. Bauer, D. C., Nevitt, M. C., Ettinger, B., and Stone, K. (1997). Low thyrotropin levels are not associated with bone loss in older women: a prospective study. J. Clin. Endocrinol. Metab. 82, 2931–2936. 48. Nuzzo, V., Lupoli, G., Esposito Del Puente, A., Rampone, E., Carpinelli, A., Del Puente, A. E., and Oriente, P. (1998). Bone mineral

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64. 65.

66.

density in premenopausal women receiving levothyroxine suppressive therapy. Gynecol. Endocrinol. 12, 333–337. Ross, D. S., Ardisson, L. J., Nussbaum, S. R., and Meskell, M. J. (1990). Serum osteocalcin in patients taking L-thyroxine who have subclinical hyperthyroidism. J. Clin. Endocrinol. Metab. 72, 507–509. Mikosch, P., Jauk, B., Gallowitsch, H. J., Pipam, W., Kresnik, E., and Lind, P. (2001). Suppressive levothyroxine therapy has no significant influence on bone degradation in women with thyroid carcinoma: a comparison with other disorders affecting bone metabolism. Thyroid 11, 257–263. Toivonen, J., Tahtela, R., Laitinen, K., Risteli, J., and Valimaki, M. J. (1998). Markers of bone turnover in patients with differentiated thyroid cancer with and following withdrawal of thyroxine suppressive therapy. Eur. J. Endocrinol. 138, 667–673. Faber, J., and Galloe, A. M. (1994). Changes in bone mass during prolonged subclinical hyperthyroidism due to L-thyroxine treatment: a meta-analysis. Eur. J. Endocrinol. 130, 350–356. Uzzan, B., Campos, J., Cucherat, M., Nony, P., Boissel, J. P., and Perret, G. Y. (1996). Effects of bone mass of long term treatment with thyroid hormones: a meta-analysis. J. Clin. Endocrinol. Metab. 81, 4278–4289. Harvey, R. D., McHardy, K. C., Reid, I. W., Paterson, F., Bewsher, P. D., Duncan, A., and Robins, S. P. (1991). Measurement of bone collagen degradation in hyperthyroidism and during thyroxine replacement therapy using pyridinium cross-links as specific urinary markers. J. Clin. Endocrinol. Metab. 72, 1189–1194. Pioli, G., Pedrazzoni, M., Palummeri, E., Sianesi, M., Del Frate, R., Vescovi, P. P., Preisco, M., Ulietti, V., Costi, D., and Passeri, M. (1992). Longitudinal study of bone loss after thyroidectomy and suppresseive thyroxine therapy in premenopausal women. Acta. Endocrinol. 126, 238–242. Guo, C. Y., Weetman, A. P., and Eastell, R. (1997). Longitudinal changes of bone mineral density and bone turnover in postmenopausal women on thyroxine. Clin. Endocrinol. 46, 301–307. Rosen, H. N., Moses, A. C., Garber, J., Ross, D. S., Lee, S. L., Ferguson, L., Chen, V., Lee, K., and Greenspan, S. L. (1998). Randomized trial of pamidronate in patients with thyroid cancer: bone density is not reduced by suppressive doses of thyroxine, but is increased by cyclic intravenous pamidronate. J. Clin. Endocrinol. Metab. 83, 2324–2330. Kung, A. W., and Yeung, S. S. (1996). Prevention of bone loss induced by thyroxine suppressive therapy in postmenopausal women: the effect of calcium and calcitonin. J. Clin. Endocrinol. Metab. 81, 1232–1236. Dhuper, S., Warren, M. P., Brooks-Gunn, J. and Fox, R. (1990). Effects of hormonal status on bone density in adolescent girls. J. Clin. Endocrinol. Metab. 71, 1083–1088. Holmes, S. J., and Shalet, S. M. (1996). Role of growth hormone and sex steroids in achieving and maintaining normal bone mass. Horm. Res. 45, 86–93. Burckardt, P., and Michel, C. (1989). The peak bone mass concept. Clin. Rheumatol. 8 (Suppl 2), 16–21. Turner, R. T., Riggs, B. L., and Spesberg, T. C. (1994). Skeletal effects of estrogen. Endocr. Rev. 15, 275–300. Ernst, M., Parkert, M. G., and Rodan, G. A. (1991). Functional estrogen receptors in osteoblastic cells demonstrated by transfection with a reporter gene containing an estrogen response element. Mol. Endocrinol. 5, 1597–1606. Horowitz, M. C. (1993). Cytokines and estrogen in bone: Anti-osteoporotic effects. Science 260, 626–627. Ford, C. E., Jones, K. W., Polani, P. E., de Almeida, J. C., and Briggs, J. H. (1959). A sex-chromosome anomaly in a case of gonadal dysgenesis (Turner’s Syndrome). Lancet 1, 711–713. Beal, R. K. (1973). Orthopedic aspects of the XO (Turner’s) syndrome. Clin. Orthopaed. Relat. Res. 97, 19–30.

JEAN E. MULDER ET AL. 67. Davies, M. C., Gulekli, B., and Jacobs, H. S. (1995). Osteoporosis in Turner’s syndrome and other forms of primary amenorrhoea. Clin. Endocrinol. 43, 741–746. 68. Shore, R. M., Chesney, R. W., Mazess, R. B., Rose, P. B., and Bargman. (1982). Skeletal dimenaralization in Turner’s syndrome. Calcif. Tissue Int. 34, 519–522. 69. Mora, S., Weber, G., Guarneri, M. P., Nizzoli, G., Pasolini, D., and Chiumello, G. (1992). Effect of estrogen replacement therapy on bone mineral content in girls with Turner’s syndrome. Obstet. Gynecol. 79, 747–751. 70. Smith, M. A., Wilson, J., and Price, W. H. (1982). Bone demineralisation in patients with Turner’s syndrome. J. Med. Genet. 19, 100–103. 71. Ross, J. L., Long, L. M., Feuillan, P., Cassorla, F., and Cutler, G. B. (1991). Normal bone density of the wrist and spine and increased wrist fractures in girls with Turner’s syndrome. J. Clin. Endocrinol. Metab. 73, 355–359. 72. Neely, E. K., Marcus, R., Rosenfeld, R. G., and Bachrach, L. K. (1993). Turner’s syndrome adolescents receiving growth hormone are not osteopenic. J. Clin. Endocrinol. Metab. 76, 861–866. 73. Shaw, N. J., Rehan, V. K., Husain, S., Marshall, T., and Smith, C. S. (1997). Bone longitudinal density in turner’s syndrome – a longitudinal study. Clin. Endocrinol. 47, 367–370. 74. Sass, T. C., De Muinck Keizer-Schrama, S. M., Stijnen, T., Asarfi, A., Van Leuwen, W. J., Van Teunenbroek, A., Van Rijn, R. R., and Drop, S. L. (2000). A longitudinal study on bone mineral density until adulthood in girls with Turner’s syndrome participating in a growth hormone injection frequency-response trial. Clin. Endocrinol. 52, 531–536. 75. Kurabayashi, T., Yasuda, M., Fujimaki, T., Yamamoto, Y., Oda, K., and Tanaka, K. (1993). Effect of hormone replacement therapy on spinal bone mineral density and T lymphocyte subsets in premature ovarian failure and Turner’s syndrome. Int. J. Gynecol. Obstet. 42, 25–31. 76. Stepan, J. J., Musilova, J., and Pacovsky, V. (1989). Bone demineralization, biochemical indices of bone remodeling, and estrogen replacement therapy in adults with Turner’s syndrome. J. Bone Min. Res. 4, 193–198. 77. Louis, O., Devroey, P., Kalender, W., and Osteaux, M. (1989). Bone loss in young hypoestrogenic women due to primary ovarian failure: spinal quantitative computed tomography. Fertil. Steril. 52, 227–231. 78. Carrascosa, A., Gussinye, M., Terradas, P., Yeste, D., Audi, L., and Vicens-Calvet, E. (2000). Spontaneous, but not induced, puberty permits adequate bone mass acquisition in adolescent Turner syndrome patients. J. Bone Miner. Res. 15, 2005–2010. 79. Hanton, L., Axelrod, L., Bakalov, V., and Bondy, C. A. (2003). The importance of estrogen replacement in young women with Turner’s syndrome. J. Womens Health 12, 971–977. 80. Emans, S. J., Grace, E., Hoffer, F. A., Gundberg, C., Ravnikar, V., and Woods, E. R. (1990). Estrogen deficiency in adolescents and young adults: Impact on bone mineral content and effects of estrogen replacement therapy. Obstet. Gynecol. 76, 585–592. 81. Gulekli, B., Davies, M. C., and Jacobs, H. S. (1994). Effect of treatment on established osteoporosis in young women with amenorrhoea. Clin. Endocrinol. 41, 275–281. 82. Suganuma, N., Furuhashi, M., Hirooka, T., Moriwaki, T., Hasegawa, Y., Mori, O., and Ogawa, M. (2003). Bone mineral density in adult patients with Turner’s syndrome: Analyses of the effectiveness of GH and ovarian steroid hormone replacement therapies. Endocrine J. 50, 263–269. 83. Bakalov, V. K., Axelrod, L., Baron, J., Hanton, L., Nelson, L. M., Reynolds, J. C., Hill, S., Troendle, J., and Bondy, C. A. (2003). Selective reduction in cortical bone mineral density in Turner syndrome independent of ovarian hormone deficiency. J. Clin. Endocrinol. Metab. 88, 5717–5722. 84. Brown, D. M., Jowsey, J., Phil, D., and Bradford, D. S. (1974). Osteoporosis in ovarian dysgenesis. J. Pediatr. 84, 816–820.

735

Chapter 43 Secondary Osteoporosis 85. Saggese, G., Federico, G., Bertelloni, S., and Baroncelli, G. I. (1992). Mineral metabolism in Turner’s syndrome: evidence for impaired renal vitamin D metabolism and normal osteoblast function. J. Clin. Endocrinol. Metab. 75, 998–1001. 86. Zseli, J., Bosze, P., Lakatos, P., Vargha, P., Tarjan, G., Kollin, E., Horvath, C., Laszlo, J., and Hollo, I. (1991). Serum bone gla protein in streak gonad sydrome. Calcif. Tissue Int. 48, 387–391. 87. Jakubowski, L. (1981). Urinary total hydroxiproline excretion in patients with Turner’s syndrome and Klinefelter’s syndrome. Horm. Metab. Res. 13, 399–403. 88. Rauch, F., Seibel, M., Woitge, H., Kruse, K., and Schonau, E. (1995). Increased urinary excretion of collagen crosslinks in girls with Ullrich-Turner syndrome. Acta. Paediatr. 84, 66–69. 89. Holl, R. W., Kunze, D., Blum, W. F., Benz, R., Etzrodt, H., and Heinze. (1993). The somatotropin- somatomedin axis in adult patients with Turner syndrome: Measurements of stimulated GH, GH-BP, IGF-I, IGF-II and IGFBP-3 in 25 patients. Horm. Res. 39, 30–53. 90. Barreca, A., Larizza, D., Damonte, G., Arvigo, M., Ponzani, P., Cesarone, A., Curto, F., Severi, F., Giordano, G., and Minuto, F. (1997). Insulin-like growth factors (IGF-I and IGF-II) and IGF-binding protein-3 production by fibroblasts of patients with Turner’s syndrome in culture. J. Clin. Endocrinol. Metab. 82, 1041–1046. 91. Saggese, G., Federico, G., and Cinquanta, L. (1995). Plasma growth hormone-binding protein activity, insulin like growth factor I, and its binding protein levels in patients with Turner’s syndrome: effect of short- and long-term recombinant growth hormone administration. Pediatr. Res. 37, 106–111. 92. Rosenfeld, R. G. (1992). Growth hormone therapy in Turner’s syndrome: an update on final heights. Acta. Paediat. Scand. Suppl 383, 3–6. 93. Shangold, M. M., and Levine, H. S. (1979). The effect of marathon training upon menstrual function. Am. J. Obstet. Gynecol. 54, 47. 94. Sanborne, C. F., Martin, B. J., and Wagner, W. W. (1982). Is athletes amenorrhea specific to runners? Am. J. Obstet. Gynecol. 143, 882–886. 95. Warren, M. P. (1980). The effect of exercise on pubertal progression and reproductive functions in girls. J. Clin. Endocrinol. Metab. 51, 1150–1157. 96. Fisher, E. C., Nelson, M. E., Fronteira, W. R., Turksoy, R. N., and Evans, W. J. (1986). Bone mineral content and levels of gonadotropins and estroges in amenorrheic running women. J. Clin. Endocrinol. Metab. 62, 1232–1236. 97. Drikwater, B. L., Nilson, K., Chesnut, C. H., Bremner, W. J., Shainholtz, S., and Southworth, M. B. (1984). Bone mineral contend of amenorrheic and eumenorrheic athletes. N. Engl. J. Med. 311, 277–281. 98. Myerson, M., Gutin, B., Warren, M. P., Wang, J., Lichtman, S., and Pierson, R. N. (1992). Total body bone density in amenorrheic runners. Obstet. Gynecol. 79, 973–978. 99. Marcus, R., Cann, C., Madvig, P., Minkoff, J., Goddard, M., Bayer, M., Martin, M., Gaudiani, L., Haskell, W., and Genant, H. (1985). Menstrual function and bone mass in elite women distance runners. Ann. Inter. Med. 102, 158–163. 100. Rencken, M. L., Chesnut, C. H., and Drinkwater, B. L. (1996). Bone density at multiple skeletal sites in amenorrheic athletes. JAMA 276, 238–240. 101. Seeman, E., Wahner, H. W., Offord, K. P., Johnson, W. J., and Riggs, B. L. (1982). The differential effects of endocrine dysfunction on the axial and appendicular skeleton. J. Clin. Invest. 69, 1302–1309. 102. Young, N., Formica, C., Szmukler, G., and Seeman, E. (1994). Bone density at weight-bearing and nonweight-bearing sites in ballet dancers: the effect of exercise, hypogonadism, and body weight. J. Clin. Endocrinol. Metab. 78, 449–454. 103. Robinson, T. L., Snow-Harter, C., Taaffe, D. R., Gillis, D., Shaw, J., and Marcus, R. (1995). Gymnasts exhibit higher bone mass than runners

104.

105.

106.

107.

108.

109. 110.

111.

112. 113.

114.

115.

116.

117.

118.

119.

120.

121.

despite similar prevalence of amenorrhea and oligomenorrhea. J. Bone Min. Res. 10, 26–35. Slemenda, C. W., and Johnston, C. C. (1993). High intensity activities in young women: Site specific bone mass effects among female figure skaters. Bone Miner. Res. 20, 125–132. Takacs, C., Solidum, A. A., Zimmerli, E. J., and Warren, M. P. (1997). Osteocalcin in young women with secondary amenorrhea. pp.131 1997 Society for Gynecologic Investigation, Annual Meeting, San Diego, CA. Herrmann, M., and Herrmann, W. (2004). The assessment of bone metabolism in female elite endurance athletes by biochemical bone markers. Clin. Chem. Lab. Med. 42, 1384–1389. Zanker, C. L., and Swaine, I. L. (1998). Relation between bone turnover, estradiol, and energy balance in women distance runners. Br. J. Sports. Med. 32, 167–171. Grinspoon, S., Thomas, E., Pitts, S., Gross, E., Mickley, D., Miller, K., Herzog, D., and Klibanski, A. (2000). Prevalence and predictive factors for regional osteopenia in women with anorexia nervosa. Ann. Int. Med. 133, 790–794. Andersen, A. E., Woodward, P. J., and LaFrance, N. (1995). Bone mineral density of eating disorder subgroups. Int. J. Eating Disord. 18, 335–342. Davies, K. M., Pearson, P. H., Huseman, C. A., Greger, N. G., Kimmel, D. K., and Reccker, R. R. (1990). Reduced bone mineral in patients with eating disorders. Bone 11, 143–147. Biller, B. M. K., Saxe, V., Herzog, D. B., Rosenthal, D. I., Holzman, S., and Klibansk, A. (1989). Mechanisms of osteoporosis in adult and adolescent women with anorexia nervosa. J. Clin. Endocrinol. Metab. 68, 548–554. Carmichael, K. A., and Carmichael, D. H. (1995). Bone metabolism and osteopenia in eating disorders. Medicine 74, 254–267. Klibanski, A., Biller, B. M. K., Schoenfeld, D. A., Herzog, D. B., and Saxe, V. C. (1995). The effects of estrogen administration on trabecular bone loss in young women with anorexia nervosa. J. Clin. Endocrinol. Metab. 80, 898–904. Rigotti, N. A., Neer, R. M., Skates, S. J., Herzog, D. B., and Nussbaum, S. R. (1991). The clinical course of osteoporosis in anorexia nervosa. A longitudinal study of cortical bone mass. JAMA 265, 1133–1138. Maugars, Y. M., Berthelot, J. M. M., Forestier, R., Mammar, N., Ladande, S., Venisse, J. L., and Prost, A. M. (1996). Follow up of bone mineral density in 27 cases of anorexia nervosa. Euro. J. Endocrin. 135, 591–597. Miller, K. K., Grinspoon, S., Gleysteen, S., Grieco, K. A., Ciampa, J., Breu, J., Herzog, D. B., and Klibanski, A. (2004). Preservation of neuroendocrine control of reproductive function despite severe undernutrition. J. Clin. Endocrinol. Metab. 89, 4434–4438. Kaplan, F. S., Pertschuck, M., Fallon, M., and Haddad, J. (1986). Osteoporosis and hip fracture in a young woman with anorexia nervosa. Clin. Orthop. 212, 250–254. Joyce, J. M., Warren, D. L., Humphries, L. L., Smith, A. J., and Coon, J. S. (1990). Osteoporosis in women with eating disorders: comparison of physical parameters, exercise, and menstrual status with SPA and DPA evaluation. J. Nucl. Med. 31, 325–331. Fonseca, V. A., D’ Souza, V., Houlder, S., Thomas, M., Wakeling, A., and Dandona, P. (1988). Vitamin D deficiency and low osteocalcin concentrations in anorexia nervosa. J. Clin. Pathol. 41, 195–197. Grinspoon, S., Baum, H., Lee, K., Anderson, E., Herzog, D., and Klibanski, A. (1996). Effects of short-term recombinant human insulin-like growth factor I administration on bone turnover in osteopenic women with anorexia nervosa. J. Clin. Endocrinol. Metab. 81, 3864–3870. Maugars, Y., and Prost, A. (1989). Osteoporose trabeculaire compliquant une anorexie mentale meconnue. Rev. Rhum. Mal. Osteoartic. 56, 159–161.

736 122. Miller, K. K., Grieco, K. K., Mulder, J. E., Grinspoon, S., Mickley, S., Yehezkel, R., Herzog, D. B., and Klibanski, A. (2004). Effects of Risedronate on bone density in anorexia nervosa. J. Clin. Endocrinol. Metab. 89, 3903–3906. 123. Zipfel, S., Seibel, M. J., Lowe, B., Beumont, P. J., Kasperk, C., and Herzog, W. (2001). Osteoporosis in eating disorders: A follow-up study of patients with anorexia and bulimia nervosa. J. Clin. Endocrinol. Metab. 86, 5227–5233. 124. Boyar, R. M., Hellman, L. D., Roffwarg, H., Kartz, J., Zumoff, B., O’Connor, J., Bradlow, H. L., and Fukushima, D. K. (1977). Corisol secretion and metabolism in anorexia nervosa. N. Engl. J. Med. 296, 190–193. 125. Kontula, K., Mustajoki, P., and Pelkonen, R. (1984). Development of cushing’s disease in a patient with anorexia nervosa. J. Endocrinol. Invest. 7, 35–40. 126. Counts, D. R., Gwirtsman, H., Carlsson, L. M., Lesem, M., and Cutler, G. B. (1992). The effects of anorexia nervosa and refeeding on growth hormone-binding protein, the insulin-like growth factors (IGFs), and the IGF-binding proteins. J. Clin. Endocrinol. Metab. 75, 762–767. 127. Grinspoon, S., Thomas, L., Miller, K. K., Herzog, D. B., and Klibaski, A. (2002). Effects of recombinant human IGF-I and oral contraceptive administration on bone density in anorexia nervosa. J. Clin. Endocrinol. Metab. 87, 2883–2891. 128. Petersson, F., Fries, H., and Nillius, S. J. (1973). Epidemiology of secondary amenorrhea. Incidence and prevalence rates. Am. J. Obstet. Gynecol. 117, 80–86. 129. Klibanski, A., and Greenspan, S. L. (1986). Increase in bone mass after treatment of hyperprolactinemic amenorrhea. N. Engl. J. Med. 315, 542–546. 130. Kayath, M. J., Lengyel, A. M. J., and Vieira, J. G. H. (1993). Prevalence and magnitude of osteopenia in patients with prolactinoma. Brazilian J. Med. Biol. Res. 26, 933–941. 131. Ciccarelli, E., Savino, L., Carlevatto, V., Bertagna, A., Isaia, G. C., and Camanni, F. (1988). Vertebral bone density in non amenorrhoeic hyperprolactinaemic women. Clin. Endocrinol. 28, 1–6. 132. Di Somma, C., Colao, A., Di Sarno, A., Klain, M., Landi, M. L., Facciollo, G., Pivonello, R., Panza, N., Salvatore, M., and Lombardi, G. (1998). Bone marker and bone density responses to dopamine agonist therapy in hyperprolactinemic males. J. Clin. Endocrinol. Metab. 83, 807–813. 133. Biller, B. M. K., Baum, H. B. A., Rosenthal, D. I., Saxe, V. C., Charpie, P. M., and Klibanski, A. (1992). Progressive trabecular osteopenia in women with hyperprolactinemic amenorrhea. J. Clin. Endocrinol. Metab. 75, 692–697. 134. Schlechte, J., El-Khoury, G., Kathol, M., and Walkner, L. (1987). Forearm and vertebral bone mineral in treated and untreated hyperprolactinemia amenorrhea. J. Clin. Endocrinol. Metab. 64, 1021–1026. 135. Klibanski, A., Biller, B. M. K., Rosenthal, D. I., Schoenfeld, D. A., and Saxe, V. (1988). Effects of prolactin and estrogen deficiency in amenorrheic bone loss. J. Clin. Endocrinol. Metab. 67, 124–130. 136. Koppelman, M. C. S., Kurtz, D. W., Morrish, K. A., Bou, E., Susser, J. K., Shapiro, J. R., and Loriax, D. L. (1994). Vertebral body bone mineral content in hyperprolactinemic women. J. Clin. Endocrinol. Metab. 59, 1050–1053. 137. Greenspan, S. L., Oppenheim, D. S., and Klibanski, A. (1989). Importance of gonadal steroids to bone mass in men with hyperprolactinemic hypogonadism. Ann. Intern. Med. 110, 526–531. 138. Schlechte, J., Walkner, L., and Kathol, M. (1992). A longitudinal analysis of premenopausal bone loss in healthy women and women with hyperprolactinemia. J. Clin. Endocrinol. Metab. 75, 698–703. 139. Suva, L. J., Winslow, G. A., Wettenhall, R. E. H., Hammonds, R. G., Mosely, J. M., Diefenbach-Jagger, H., Rodda, C. P., Kemp, B. E., Rodriguez, H., and Chen, E. Y. (1987). A parathyroid hormone-related

JEAN E. MULDER ET AL.

140.

141.

142.

143. 144. 145.

146. 147.

148.

149.

150.

151.

152.

153. 154.

155.

156.

157. 158.

159.

protein implicated in malignant hypercalcemia: cloning and expression. Science 237, 893–896. Kovacs, C. S., and Chik, C. L. (1995). Hyperprolactinemia caused by lactation and pituitary adenomas is associated with altered serum calcium, phosphate, parathyroid hormone (PTH), and PTH- related peptide levels. J. Clin. Endocrinol. Metab. 80, 3036–3042. Stiegler, C., Leb, G., Kleinert, R., Warnkross, H., RamschakSchwarzer, S., Lipp, R., Clarici, G., Krejs, G. J., and Dobnig, H. (1995). Plasma levels of parathyroid hormone-related peptide are elevated in hyperprolactinemia and correlated to bone density status. J. Bone Min. Res. 10, 751–759. Sartorio, A., Conti, A., Ambrosi, B., Muratori, M., and Faglia, G. (1990). Osteocalcin levels in patients with microprolactinoma before and during medical treatment. J. Endocrinol. Invest. 13, 419–422. Pack, A. M., and Morrell, M. J. (2004). Epilepsy and bone health in adults. Epilepsy Behav. 5, S24–29. Sheth, R. D. (2004). Bone health in pediatric epilepsy. Epilepsy Behav. 5, S30–35. Dent, C., Richens, A., Rowe, D. F. J., and Stamp, T. (1970). Osteomalacia with long-term anticonvulsant therapy in epilepsy. B. M. J. 4, 69–72. Richens, A., and Rowe, D. F. J. (1970). Disturbance of calcium metabolism by antoconvulsant drugs. B. M. J. 4, 73–76. Andress, D. L., Ozuna, J., Tirschwell, D., et al. (2002). Antiepileptic drug-induced bone loss in young male patients who have seizures. Arch. Neurol. 59, 781–786. Farhat, G., Yamout, B., Mikati, M. A., Demirjian, S., Sawaya, R., and El Hajj Fuliehan, G. (2002). Effect of antiepileptic drugs on bone density in ambulatory patients. Neurology 58, 1348–1353. Kubota, F., Kifune, A., Shibata, N., Akata, T., Takeuchi, K., Takahashi, S., Ohsawa, M., and Takama, F. (1999). Bone mineral density of epileptic patients on long-term antiepileptic drug therapy: a quantitative digital radiography study. Epilepsy Res. 33, 93–97. Pack, A. M., Olarte, M., Morrell, M. J., Flaster, E., Resor, S. R., and Shane, E. (2003). Bone mineral density in an outpatient population receiving enzyme-inducing antiepileptic drugs. Epilepsy Behav. 4, 169–174. Sato, Y., Kondo, I., Ishida, S., et al. (2001). Decreased bone mass and increased bone turnover with valproate therapy in adults with epilepsy. Neurology 57, 445–449. Valimaki, M. J., Tiihonen, M., Laitinen, K., Tahtela, R., Karkkainen, M., Lamberg-Allardt, C., Makela, P., and Tunninen, R. (1994). Bone mineral density measured by dual-energy X-ray absorptiometry and novel markers of bone formation and resorption in patients on antiepileptic drugs. J. Bone Miner. Res. 9, 631–637. Mattson, R. H., and Gidal, B. E. (2004). Fractures, epilepsy, and anticonvulsant drugs. Epilepsy Behav. 5, S36–40. Desai, K. B., Ribbans, W. J., and Taylor, G. J. (1996). Incidence of five common fracture types in an institutional epileptic population. Injury 27, 97–100. Vestergaard, P., Tigaran, S., Rejnmark, L., Tigaran, C., Dam, M., and Mosekilde, L. (1999). Fracture risk is increased in epilepsy. Acta. Neurol. Scand. 99, 269–275. Ensrud, K. E., Blackwell, T. L., Mangione, C. M., et al. (2002). Central nervous system-actvei medications and risk for falls in older women. J. Am. Geriatr. Soc. 50, 1629–1637. Fitzpatrick, L. A. (2004). Pathophysiology of bone loss in patients receiving anticonvulsant therapy. Epilepsy Behav. 5, S3–S15. Matheson, R. T., Herbst, J. J., Jubiz, W., Freston, J. W., and Tolman, K. G. (1976). Absorption and biotransformation of cholecalciferol in drug-induced osteomalacia. J. Clin. Pharmacol. 16, 426–432. Hahn, T. J., Hendin, B. A., Scharp, C. R., and Haddad, J. G. (1972). Effect of chronic anticonvulsant therapy on serum 25-hydroxycholecalciferol levels in adults. N. Engl. J. Med. 287, 900–904.

Chapter 43 Secondary Osteoporosis 160. Hahn, T. J., Birge, S. J., Scharp, C. R., and Avioli L. V. (1972). Phenobarbital induced alterations in vitamin D metabolism. J. Clin. Invest. 51, 741–748. 161. Hahn, T. J., Hendin, B. A., Scharp, C. R., Boisseau, V. C., and Haddad, J. J. G. (1975). Serum 25-hydroxycalciferol levels and bone mass in children on chronic anticonvulsant therapy. N. Engl. J. Med. 292, 550–554. 162. Stamp, T. C. B., Round, J. M., Rowe, D. J. F., and Haddad, J. J. G. (1972). Plasma levels and therapeutic effect of 25-hydroxycholecalciferol in epileptic patients taking anticonvulsant drugs. B. M. J. 4, 9–12. 163. Gough, H., Goggin, T., Bissessar, A., Baker, M., Crowlay, M., and Callaghan, N. (1986). A comparative study of the relative influence of different anticonvulsant drugs, UV exposure and diet on vitamin D and calcium metabolism in out-patients with epilepsy. Q. J. Med. 230, 569–577. 164. Hoikka, V., Savolainen, K., Alhave, E. M., Sivenius, J., Karjalainen, P., and Repo, A. (1981). Osteomalacia in institutionalized epileptic patinets on long-term anticonvulsant therapy. Acta. Neurol. Scand. 64, 122–131. 165. Okesina, A. B., Donaldson, D., and Lascelles, P. T. (1991). Isoenzymes of alkaline phosphatase in epileptic patients receiving carbamazepine monotherapy. J. Clin. Pathol. 44, 480–482. 166. Davie, M. W., Emberson, C. E., Lawson, D. E., Roberts, G. E., Barnes, J. L., Barnes, N. D., and Heeleg, A. F. (1983). Low plasma 25-hydroxyvitamin D and serum calcium levels in institutionalized epileptic subjects: assocaited risk factors, consequences and response to treatment with vitamin, D. Q. J. Med. 205, 79–91. 167. Tjellesen, L., and Christiansen, C. (1982). Serum vitamin D metabolites in epileptic patients treated with 2 different anyconvulsants. Acta. Neurol. Scand. 66, 335–341. 168. Wark, J. D., Larkins, R. G., Perry-Keene, D., Peter, C. T., Ross, D. L., and Sloman, J. G. (1979). Chronic diphenylhydantoin therapy does not reduce plasma 25-hydroxy-vitamin, D. Clin. Endocrinol. 11, 267–274. 169. Weinstein, R. S., Bryce, G. F., Sappington, L. J., King, D. W., and Gallagher, B. B. (1984). Decreased serum ionized calcium and normal vitamin D metabolite levels with anticonvulsant drug treatment. J. Clin. Endocrinol. Metab. 58, 1003–1009. 170. Pack, A. M., Morrell, M. J., Marcus, R., Holloway, L., Flaster, E., Done, E., Randall, A., Seale, C., and Shane, E. (2005). Bone metabolism and turnover in women with epilepsy on antiepileptic drug monotherapy. Ann. Neurol. 57, 252–257. 171. Koch, H. C., Kraft, D., and Herrath, D. V. (1972). Influence of diphenylhydantoin and phenobarbital on intestinal calcium transport in the rat. Epilepsia 13, 829–841. 172. Harrison, H. C., and Harrison, H. E. (1976). Inhibition of vitamin Dstimulated active transport of calcium of rat intestine by diphenylhydantoin phenobarbital treatment. Proc. Soc. Exp. Biol. Med. 153, 220–224. 173. Hahn, T. J., and Halstead, L. R. (1979). Anticonvulsant drug-induced osteomalacia: Alterations in mineral metabolism and response to vitamin D3 administration. Calcif. Tissue Int. 27, 13–18. 174. Hahn, T. J., Scharp, C. R., Richardson, C. A., Halstead, L. R., Kahn, A. J., and Teitelbaum, S. L. (1978). Interaction of diphenylhydantoin (phenytoin) and phenobarbital with hormonal mediation of fetal rat bone resorption in vitro. J. Clin. Invest. 62, 406–414.

737 175. Jenkins, M. V., Harris, M., and Willis, M. R. (1974). The effect of phenytoin on parathyroid extract and 25-hydroxycholecalciferolinducedbone resorption: Adenosine 3’, 5’-cyclic monophosphate production. Calcif. Tissue Res. 16, 163–167. 176. Dietrich, J. W., and Duffield, R. (1980). Effects of diphenylhydantoin on synthesis of collagen and noncollagen protein in tissue culture. Endocrinology. 106, 606–610. 177. Yamano, Y., Sakane, S., Takamatsu, J., and Ohsawa, N. (1999). Estrogen supplementation for bone dematuration in young epileptic man treated with anticonvulsant therapy; a case report. Endocr. J. 46, 301–307. 178. Heroz, A. G., Levesque, L. A., Drislane, F. W., Ronthal, M., and Schomer, D. L. (1991). Phenytoin-induced elevaion of serum estradiol and reproductive dysfunction in men with epilepsy. Epilepsia 32, 550–553. 179. Mosekilde, L., and Melsen, F. (1980). Dynamic differences in trabecular bone remodeling between patients after jejuno-ileal bypass for obesity and epileptic patients receiving anticonvulsant therapy. Metab. Bone Dis. Relat. Res. 2, 77–82. 180. Takeshita, N., Seino, Y., Ishida, H., Tanaka, H., Tsutsumi, C., Ogata, K., Kiyohara, K., Kato, H., Nozawa, M., Akiyama, Y., Hara, K., and Imura, H. (1989). Increased circulating levels of a-carboxyglutamic acid-containing protein and decreased bone mass in children on anticonvulsant therapy. Cacif. Tissue Int. 44, 80–85. 181. Lau, K. H., Nakade, O., Barr, B., Taylor, A. K., Houchin, K., and Baylink, D. J. (1995). Phenytoin increases markers of osteogenesis for the human species in vitro and in vivo. J. Clin. Endocrinol. Metab. 80, 2347–2353. 182. Tjellesen, L., Hummer, L., Christiansen, C., and Rodbro, P. (1986). Different metabolism of vitamin D2/D3 in epileptic patients treated with phenobarbitone/phenytoin. Bone 7, 337–342. 183. Ohishi, T., Kushida, K., Takahashi, M., Kawana, K., Yagi, K., Kwakami, K., Horiuchi, K., and Inoue, T. (1994). Urinary bone resorption markers in patients with metabolic bone disorders. Bone 15, 15–20. 184. Verrotti, A., Greco, R., Morgese, G., and Chiarelli, F. (2000). Increased bone turnover in epileptic patients treated with carbamazepine. Ann. Neurol., 47, 385–388. 185. Verrotti, A., Greco, R., Latini, G., Morgese, G., and Chiarelli, F. (2002). Increased bone turnover in prepubertal, pubertal and postpubertal patients receiving carbamazepine. Epilepsia 43, 1488–1492. 186. Sheth, R. D., Wesolowski, C. A., Jacob, J. C., Penney, S., Hobbs, G. R., Riggs, J. E., and Bodensteiner, J. B. (1995). Effect of carbamazepine and valproate on bone mineral density [see comments]. J. Pediatr. 127, 256–262. 187. Drezner, M. K. (2004). Treatment of anticonvulsant drug-induced bone disease. Epilepsy Behav. 5, S41–S47. 188. Barden, H. S., Mazess, R. B., Rose, P. G., and McAweeney, W. (1980). Bone mineral status measured by direct photon absorptiometry in institutionalized adults receiving long-term anticonvulsant therapy and multivitamin supplementation. Calcif. Tissue Int. 31, 117–121. 189. Collins, N., Maher, J., Cole, M., Baker, M., and Callaghan, N. (1991). A prospective study to evaluate the dose of vitamin D required to correct low 25-hydroxyvitamin D levels, calcium and alkaline phosphatase in patients at risk of developing antiepileptic drug-induced osteomalacia. Quart. J. Med. 78, 113–122.

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Chapter 44

Osteomalacia and Rickets Marc K. Drezner, M.D.

I. II. III. IV.

Professor of Medicine, University of Wisconsin, Madison, Wisconsin, USA

V. Phosphopenic Rickets and Osteomalacia VI. Normal Mineral Rickets and Osteomalacia References

Definition Etiology Incidence and Epidemiology Calciopenic Rickets and Osteomalacia

I. DEFINITION

The calciopenic forms of the disease embrace disorders that affect the availability of calcium and/or vitamin D or its active metabolite. In general, these forms of the disease are marked by a low or marginally normal serum calcium level [3, 4], (secondary) hyperparathyroidism, and consequent increased bone turnover. While many of these calciopenic diseases are acquired or secondary to renal or gastrointestinal dysfunction, a subgroup of the disorders are due to genetic abnormalities of vitamin D metabolism [5, 6]. The phosphopenic disorders usually develop from an abnormality of transepithelial phosphate transport in the nephron, resulting in renal phosphate wasting. Often the underlying abnormality is due to a genetic defect, which directly or indirectly influences renal phosphate handling. Generally, patients with these disorders maintain a normal serum calcium concentration, whereas hypophosphatemia is characteristic. Unlike the calciopenic disorders, parathyroid hormone levels and bone turnover are often normal in phosphopenic forms of the disease. However, in Dent’s disease, a subtype of the phosphopenic diseases, which results from loss of function mutations of the CLCN5 gene, hypercalciuria is common and in most patients results in osteomalacia with increased bone turnover [7]. In those disorders marked by normal mineral availability, the abnormal calcification of cartilage and bone most

Rickets and osteomalacia are diseases characterized by defective bone and cartilage mineralization in children and bone mineralization in adults. The abnormal calcification of cartilage occurs at epiphyseal growth plates and contributes to an associated delayed maturation of the cartilage cellular sequence and disorganization of cell arrangement [1]. The resultant profusion of disorganized, non-mineralized, degenerating cartilage leads to widening of the epiphyseal plates with flaring or cupping, irregularity of the epiphyseal–metaphyseal junctions, attendant skeletal abnormalities and possibly retardation of growth. Abnormal calcification of bone is limited to the newly formed organic matrix deposited at the bone–osteoid interfaces of remodeling tissue. This defect results in an increase in the boneforming surface covered by incompletely mineralized osteoid, an enhanced osteoid volume and thickness, a decrease in the mineralizing surface (the percentage of bone surface undergoing calcification), and a likely heightened susceptibility to fractures and/or bone deformities [2]. The various rachitic and osteomalacic disorders recognized to date include more than 30 subtypes, which may be divided into disease forms characterized by calciopenia, phosphopenia, or normal mineral availability (Table I). Dynamics of Bone and Cartilage Metabolism

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The Rickets and Osteomalacia Syndromes

Calciopenic disorders

Phosphopenic disorders

Nutritional Calcium deficiency Vitamin D deficiency Vitamin D malabsorption Gastrointestinal disorders Partial/total gastrectomy Small bowel disease Intestinal bypass Pancreatic insufficiency Hepatobiliary disease Biliary atresia Biliary obstruction Biliary fistula Cirrhosis Loss of vitamin D metabolites Nephrotic syndrome Peritoneal dialysis Abnormal vitamin D metabolism Impaired hepatic 25-hydroxylation Liver disease Anticonvulsant therapy Impaired renal 1α-hydroxylation Vitamin D-dependent rickets, type 1 Chronic renal failure Hypoparathyroidism Pseudohypoparathyroidism Vitamin D resistance Vitamin D-dependent rickets, type 2 Hormone-binding negative Defective hormone-binding capacity Abnormal hormone-binding affinity Deficient nuclear localization Decreased hormone-receptor affinity

Nutritional Low phosphate intake Ingestion of phosphate-binding agents Impaired renal tubular phosphate transport Hereditary X-linked hypophosphatemia Hereditary hypophosphatemic rickets/hypercalciuria X-linked recessive hypophosphatemia (Dent’s disease) Autosomal dominant hypophosphatemia Fibrous dysplasia of bone Acquired Tumor-induced osteomalacia Mesenchymal, epidermal, endodermal tumors Fibrous dysplasia of bone Neurofibromatosis Linear nevus sebaceous syndrome Light chain nephropathy Sporadic hypophosphatemic osteomalacia General renal tubular disorders Fanconi’s syndrome, type 1 Hereditary Familial idiopathic Cystinosis Galactosemia Glycogen storage disease Wilson’s disease Lowe’s syndrome Acquired Renal transplantation Multiple myeloma Intoxication Cadmium Lead Fanconi’s syndrome, type 2 Normal mineral availability Primary mineralization defect Hereditary Hypophosphatasia Perinatal Infantile Childhood Adult-onset Pseudohypophosphatasia Acquired Drug-induced Fluoride Aluminum Bisphosphonates Abnormal matrix synthesis Fibrogenesis imperfecta ossium Axial osteomalacia Metabolic acidosis

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often results from the presence of a circulating inhibitor of bone mineralization (often a drug) or a genetic defect causing abnormal bone collagen or matrix. In general, these disease states are characterized by normal or low bone turnover and a variable predisposition to fracture. Although the phenotypic expression of the defective bone and cartilage mineralization is similar in this wide variety of disease forms, the associated genetic, hormonal, and biochemical abnormalities, as well as the resultant bone turnover, differ according to the pathogenetic defect. Therefore, identification of rickets and/or osteomalacia demands systematic analysis to determine cause and thereby appropriate therapeutic approaches for the disorder. Until recently, complete characterization of bone turnover in a rachitic or osteomalacic disorder and monitoring of therapeutic response and patient compliance required histomorphometric analysis of bone biopsies. However, the recent development of specific and sensitive biochemical markers, reflecting the overall rate of bone formation and bone resorption, provide relatively non-invasive measurements of potential value in assessment of the rachitic/osteomalacic diseases. Reports on the use of these markers in such disorders have increased substantially in recent years, albeit the data are still incomplete. Nevertheless, expected changes in markers of bone resorption and formation in these variable diseases is predictable for the most part, since changes in the biochemical markers reflect alterations in skeletal metabolism, which may vary with the underlying cause. Thus, coupling existent data regarding changes in bone markers in various rachitic/osteomalacic disorders with predicted alterations in the various disease states, paying particular attention to the complex mineralization process influencing cartilage and bone, should provide new insights into the pathogenesis and diagnosis of the calciopenic, phosphopenic and normal mineral variants of rickets/ osteomalacia.

II. ETIOLOGY Mineralization of cartilage and bone is a complex process in which the calcium–phosphorus inorganic mineral phase is deposited in an organic matrix in a highly ordered fashion. Such mineralization depends primarily upon the availability of sufficient calcium and phosphorus at mineralization sites. Other requisite conditions necessary for normal calcification include: (1) adequate metabolic and transport function of chondrocytes [8] and osteoblasts to regulate the concentration of calcium, phosphorus, and other ions at the mineralization sites; (2) maintenance of an optimal pH (approximately 7.6) for deposition of calcium–phosphorus complexes; (3) a low concentration of calcification

inhibitors (e.g. pyrophosphates, proteoglycans) in the circulation or bone matrix; and (4) the presence of collagen with unique type, number, and distribution of cross-links, remarkable patterns of hydroxylation and glycosylation, and abundant phosphate content, which collectively permit and facilitate deposition of mineral at gaps, “hole zones,” between the distal ends of collagen molecules.

A. Mineralization Bone mineral consists of Ca2+, PO43−, OH−, and CO32− ions, arranged in space in accordance with the crystal lattice of hydroxyapatite [9]. The mineralizing potential of extracellular fluid depends on the free ionic activity or effective concentration of Ca2+, HPO42− and H+ ions. For both calcium and phosphorus, ionic concentrations differ from the total concentrations because of protein binding and ion complexing, both of which are affected by pH. At sites of mineralization, the sucessive addition of Ca2+ and HPO42− ions present in ECF and simultaneous removal of protons generates a series of compounds beginning with secondary calcium phosphate or brushite (CaHPO4.2H2O), the first solid phase formed, and ending with hydroxyapatite [Ca10(PO4)6(OH)2] [10]. Mechanisms to accomplish initial mineral deposition include concentration gradients between mineralizing and non-mineralizing sites maintained by cells and by the ion-binding and releasing properties of macromolecules synthesized by cells, sequestration and subsequent release of calcium by mitochondria and heterogeneous nucleation by outside agents or substances [8, 10]. In contrast, mechanisms to retard the growth of hydroxyapatite crystals include the precise spatial relationships between mineral and matrix, the presence at critical locations of calcium chelators and inhibitors of mineralization such as pyrophosphate and albumin and the cellular and biochemical characteristics of the quiescent bone surface, the site of reversible mineral exchange [8]. 1. Cartilage

Within this general framework, mineral is deposited in growth plate cartilage in the form of spherical clusters of randomly oriented crystals of varying size [11]. The clusters, called calcospherulites, are spatially associated with matrix vesicles, which are small membrane-bound particles that are derived by an unknown mechanism from chondrocytes. The vesicles serve as nucleation sites and are distributed with highest density in the resting and hypertrophic zones and lowest density in the proliferative and calcifying zones of cartilage. The mineral is deposited in discrete patches, which aggregate to form a single longitudinal

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septum, 50 µm in width, extending across the growth plate [12]. Deposition of mineral is delayed approximately 24 hours after synthesis and secretion of matrix, likely reflecting changes in gene expression in chondrocytes [13]. Accumulation of uncalcified tissue in rachitic disorders is due entirely to a profound delay in this mineralization process, accompanied by structural disorganization of the metaphysic, secondary to mechanical effects and a markedly altered pattern of vascular invasion. Inadequate mineralization is confined to the maturation zone of the cartilage, whereas the resting and proliferative zones of the epiphyses exhibit normal histologic features. In the maturation zone, the height of the cell columns is increased and the cells are closely packed and irregularly aligned. Moreover, calcification in the interstitial regions of this hypertrophic zone is defective. These changes result in increased thickness of the epiphyseal plate, accompanied by an increase in transverse diameter, which may extend beyond the ends of the bone, causing characteristic cupping or flaring. The cause of this defective mineralization varies depending upon the subtype of rickets in question. However, in general, defective mineralization is secondary to decreased mineral availability or a combination of mineral and/or vitamin Ddependent or genetically impaired chondrocyte function. 2. Bone

In contrast, the mineralization of bone is marked by a paucity of matrix vesicles. Indeed, in lamellar bone, which is invariably formed in apposition to an existing surface, the matrix is compact in texture and the collagen fibrils are long and highly ordered. Mineral crystals are aligned with their long axis parallel to the collagen fibrils and are initially deposited within the hole zones by heterogeneous nucleation, but longer and wider crystals are subsequently formed on and between the fibrils [9, 12]. The deposition of mineral is, however, continuous and sharply demarcated, but delayed for approximately 2 weeks after matrix deposition. The delay likely reflects necessary maturation of the matrix, which includes completion of collagen fibril crosslinking and the development of precise orientation, conformation, and aggregation of a variety of non-collagenous proteins and proteoglycans [8, 14]. Abnormal mineralization results in accumulation of excess osteoid, a sine qua non for the diagnosis of osteomalacia in most instances. The accumulation of unmineralized osteoid in bone is generally due to a prolongation of the mineralization lag time and is manifest by an increase in the forming surface covered by incompletely mineralized osteoid and an increase in osteoid volume and thickness [15]. The mineralization process is likely influenced by the availability of mineral and vitamin D-dependent and/or genetic influences on osteoblast function. In this scenario, the osteoblast

probably functions by a direct effect on matrix maturation and by its effects on mineral transport. In any case, the evident difference in the mineralization processes in cartilage and bone accounts for the differences under some circumstances in severity and response to treatment of rickets and osteomalacia.

III. INCIDENCE AND EPIDEMIOLOGY Although vitamin D deficiency was a common form of calciopenic rickets/osteomalacia early in the 20th century, the therapeutic use of vitamin D and supplementation of foods with vitamin D has curtailed occurrence of this disease form. However, deficiency of vitamin D does persist in Asian populations, as well as in a growing number of elderly patients, vegans, children on macrobiotic diets and black children in the Western world [16–19], and vitamin D deficiency secondary to malabsorptive disorders continues to occur with relative frequency. Moreover, limited exposure to sunlight, dedicated use of sunscreen, increased frequency of exclusively breast-feeding infants and residence at northern latitudes have collectively re-established vitamin D deficiency as a significant cause of calciopenic rickets/ osteomalica throughout the world [19, 20]. In any case, over the past 20–30 years, additional subtypes of rickets/osteomalacia have been identified, which are in general characterized by a failure to respond to physiologic doses of vitamin D. These are related for the most part to calciopenic disorders due to primary vitamin D abnormalities (defective metabolism or postreceptor resistance) or deficient calcium supply (especially in many African countries) and phosphopenic diseases due to genetic abnormalities or acquired defects in phosphate transport (oncogenic osteomalacia). In addition, primary mineralization defects also occur, albeit infrequently, whether genetic (as in hypophosphatasia) or acquired (excess fluoride, aluminum, or bisphosphonate).

A. Clinical Findings The clinical manifestations of rickets are primarily related to skeletal pain and deformity, bone fractures, slipped epiphyses, and abnormalities of growth. In addition hypocalcemia, when present, may be sufficiently severe to produce tetany, laryngeal spasm, or seizures [21, 22]. In infants and young children, features include listlessness, irritability, profound hypotonia, and proximal muscle weakness. By age 6 months, frontal bossing with flattening at the back is evident. Later, a lateral collapse of both chest walls (Harrison’s sulcus) and rachitic rosary may

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appear. If untreated, progressive bony deformities result in bowing and fractures in the long bones. In addition, dental eruption may be delayed and, in those forms of the disease with hypocalcemia or hereditary hypophosphatemia, enamel defects and inadequate dentin calcification occur, respectively. Clinical signs of osteomalacia in adults are vague. Often the skeletal abnormalities are overlooked and features of an underlying disorder, such as malabsorption, may dominate. Nevertheless, diffuse skeletal pain and generalized muscular weakness are generally present [20]. Pain may be localized about the hips and result in an antalgic gait, and muscle weakness is primarily proximal and often associated with wasting and hypotonia. The myopathy is present in virtually all forms of osteomalacia, X-linked hypophosphatemia excepted [23]. In adults with a history of rickets, evidence of inactive rachitic abnormalities is often apparent. The radiographic and metabolic abnormalities characteristic of rickets/osteomalacia vary according to the presence of calciopenic (high turnover) or phosphopenic (normal to low turnover) disease, as well as disorders secondary to primary mineralization defects. Markers of bone turnover may be the discriminating factors that permit categorization and guide investigation of the disease. Moreover, serial evaluation of such markers may permit a simple means to judge efficacy of therapy. Common radiographic abnormalities in rickets are evident at the growth plate, which is increased in thickness, cupped, and displays an irregular, hazy appearance at the diaphyseal line secondary to uneven invasion of the recently calcified cartilage by adjacent bone tissue. In osteomalacia, there is usually a mild decrease in bone density associated with a coarsening of trabeculae and a blurring of their margins. Looser’s zones (or pseudofractures), ribbonlike zones of rarefaction-oriented perpendicular to the bone surface, are a more specific radiographic abnormality in osteomalacia.

IV. CALCIOPENIC RICKETS AND OSTEOMALACIA Currently recognized forms of calciopenic rickets/ osteomalacia include a group of disorders in which the availability of calcium or vitamin D and/or its active metabolite is compromised (Table I). These disorders are due to a nutritional deficiency, gastrointestinal malabsorption, abnormal vitamin D metabolism, or resistance to the effects of calcitriol. In general, the biochemical hallmarks of the disease (Table II) are a low or marginally normal serum calcium level, a decreased serum phosphorus

concentration, and (secondary) hyperparathyroidism. Commensurately, the urinary calcium is decreased due to diminished gastrointestinal absorption of calcium and the urinary phosphorus increased due to the influence of an increased circulating level of parathyroid hormone on renal function. In addition, serum 25-hydroxyvitamin D levels are decreased in states of vitamin D deficiency, albeit the range of values at which disease manifests varies with the assay employed and the severity of the disorder. Nevertheless, the 25-hydroxyvitamin D level is usually reduced to less than 10 ng/ml. In contrast, in these disorders the serum 1,25(OH)2D concentration is not always overtly decreased, and, in fact, may be elevated, secondary to the prevailing hyperparathyroidism and the stimulatory effects of parathyroid hormone on calcitriol production. Alternatively, a defect in vitamin D metabolism (impaired 1α-hydroxylation) results in an isolated deficiency of 1,25(OH)2D, whereas end-organ resistance to this active vitamin D metabolite increases the circulating level of calcitriol. Generally, alkaline phosphatase is elevated in all forms of calciopenic rickets/osteomalacia, although affected adults may manifest normal circulating levels. Unique radiographic and bone histomorphometric abnormalities of the calciopenic diseases reflect not only the presence of impaired mineralization but that of increased parathyroid hormone. In this regard, X-rays often display not only decreased bone density but subperiosteal resorption and bone cysts as evidence of longstanding secondary hyperparathyroidism. Concurrently, bone biopsies reveal the expected mineralization disorder, marked by increased osteoid and decreased mineralized surface, as well as the unambiguous signs of increased bone turnover, an increased number of osteoclasts, and osteoblasts along the bone surface (Table III).

A. Bone Markers Commensurate with the increased bone turnover due to secondary hyperparathyroidism in these calciopenic diseases, markers of bone resorption should be elevated (Table IV). However, only a few reports documenting such abnormalities are available. Nevertheless, in concert with the appearance of radiological rickets in the calciopenic disorders, documented elevations of biochemical markers have been reported. In this regard, many studies have documented that the commonly used and well-established measure of bone turnover, urinary hydroxyproline, is elevated in a wide variety of calciopenic rachitic/osteomalacic disorders [24]. However, as a consequence of the varied tissue origin and metabolism of hydroxyproline, this marker is poorly correlated with bone resorption assessed by bone

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Ν

⇑ ⇑ ⇑

⇑ ⇑ ⇑

PTH

_

Ν, ⇑

⇑ ⇑

_

_

Ν, ⇑

_

_

_

_

_

_

_

FGF-23

Metabolic Abnormalities in the Rachitic and Osteomalacic Disorders

Ν

Ν, ⇑7 Ν, ⇓

⇓, Ν ⇑

⇓, Ν ⇑ ⇑ ⇓, Ν ⇓, Ν

⇑

⇓ ⇓ ⇓

⇓ ⇓ ⇓

Calcium

Ν

Ν Ν

⇑4 ⇑4

⇑4 ⇑4 ⇑4 ⇑4 ⇑4

⇓

⇑, Ν ⇑, Ν ⇑, Ν

⇑, Ν ⇑, Ν ⇑, Ν

Phosphorus

Urinary

Ν

Ν Ν

⇑, Ν ⇑

Ν, ⇑ Ν Ν Ν Ν

Ν

⇑, Ν ⇑, Ν ⇑, Ν

⇑, Ν ⇑, Ν ⇑, Ν

Amino acids

3Although

2The

the serum calcium concentration in affected patients is generally low, in subjects with late presentation of disease the serum calcium level is normal. serum 1,25(OH)2D levels are low relative to the presence of hypophosphatemia, which should increase the circulating concentration. the serum PTH is generally normal in untreated patients, often those managed with phosphorus supplementation, with or without calcitriol, will develop an elevation of PTH. 4The 24 hour urinary excretion of phosphorus is increased relative to the serum phosphorus concentration and the renal TmP/GFR is decreased. 5Increased serum calcium concentrations occur frequently in patients with perinatal and infantile hypophosphatasia, apparently due to dyssynergy between gastrointestinal calcium absorption and defective skeletal growth and mineralization. 6Approximately 50% of patients are frankly hyperphosphatemic secondary to an increased TmP/GFR. 7Patients with hypercalcemia often exhibit hypercalciuria.

1Although

Calciopenic disorders Nutritional Calcium deficiency Vitamin D deficiency Vitamin D malabsorption Abnormal vitamin D metabolism Impaired 25-hydroxylation Impaired 1-hydroxylation Vitamin D-dependent rickets type II Phosphopenic disorders Nutritional Phosphate deficiency Hereditrary X-linked hypophosphatemia Hypophosphatemic rickets/hypercalciuria X-linked recessive hypophosphatemia Autosomal dominant hypophosphatemia Fibrous dysplasia of bone Acquired Tumor-induced osteomalacia Renal tubular disorders Normal mineral availability Primary mineralization defect Hypophosphatasia Drug-induced Abnormal matrix synthesis Fibrogenesis imperfecta ossium

Calcium

Table II.

744 MARC K. DREZNER

⇑ ⇑ ⇑ N, ⇑ N, ⇑ N, ⇑ N, ⇑ N, ⇑ N, ⇑ N, ⇑ N, ⇑

⇑ ⇑ ⇑ N, ⇑ N, ⇑ N, ⇑ N, ⇑ N, ⇑ N, ⇑ N, ⇑ N, ⇑

N N,⇓,⇑7 N

N1 N1 N1

⇓, N N,⇑2 N N N N N N,⇑2

N N N

N

N N,⇓,⇑7

⇑ ⇑ ⇑

⇑ ⇑ ⇑

PINP

N1 N1 N1

PICP

N

N N, ⇓8

N,⇓6 N N

N,⇑4 N,⇑5

N, ⇓ N N N, ⇓ N, ⇓

N, ⇓

⇑

⇑ ⇑

⇑ ⇑ ⇑

TRAP

N,⇑4 N,⇑5

N, ⇓ N3 N3 N, ⇓ N, ⇓

N, ⇓

⇑

⇑ ⇑

⇑ ⇑ ⇑

ICTP

Serum

N

N N

N,⇑4 N,⇑5

N, ⇓ N N N, ⇓ N, ⇓

N, ⇓

⇑

⇑ ⇑

⇑ ⇑ ⇑

OHP

N

N,⇓6 N,⇓8

N,⇑4 N,⇑5

N, ⇓ N3 N3 N, ⇓ N, ⇓

N, ⇓

⇑

⇑ ⇑

⇑ ⇑ ⇑

PYD

Urinary

Bone resorption markers

N

N,⇓6 N,⇓8

N,⇑4 N,⇑5

N, ⇓ N3 N3 N, ⇓ N, ⇓

N, ⇓

⇑

⇑ ⇑

⇑ ⇑ ⇑

DPD

OC, osteocalcin; PICP, C-terminal procollagen I extension peptides; PINP, N-terminal procollagen I extension peptides; ICTP, carboxyterminal telopeptide of type I collagen; TRAP, tartrate resistant acid phosphatase; OHP, hydroxyproline; PYD, pyridinoline crosslinks; DPD, deoxypyridinoline crosslinks. 1Osteocalcin measurements are generally normal to low in untreated patients; however, in early stages of the disease elevation of serum 1,25(OH) D levels, commensurate with increased parathyroid hormone, is 2 associated with above normal concentrations; with treatment of the disease the OC level will also increase to above normal levels. 2Although the serum osteocalcin is generally normal, in a select few an increased concentration has been observed; whether this elevation corresponds to the infrequent hyperparathyroidism encountered in subjects with this disease remains unknown. 3The possibility that a sensitive marker of resorption will be decreased is less in these disorders, since the serum 1,25(OH) D concentration is elevated which may increase osteoclast number/activity to a limited 2 degree. 4An increased level of a resorption marker may occur in the limited subset of patients with elevated urinary cyclic AMP. 5The elevated serum parathyroid hormone concentration that occurs in some affected subjects may increase the resorption markers. 6In the subset of patients with hypercalcemia and suppressed parathyroid hormone secretion bone resorption markers may be decreased. 7Collagen production and the markers reflecting this process may be quite variable in drug-induced disease, with a decrease realized secondary to aluminum and an increase due to fluoride. 8Sensitive markers of bone resorption may be decreased in drug-induced disease secondary to agents such as bisphosphonates.

Calciopenic disorders Nutritional Calcium deficiency Vitamin D deficiency Vitamin D malabsorption Abnormal vitamin D metabolism Impaired 25-hydroxylation Impaired 1-hydroxylation Vitamin D resistance Vitamin D-dependent rickets, type 2 Phosphopenic disorders Nutritional Phosphate deficiency Hereditrary X-linked hypophosphatemia Hypophosphatemic rickets/hypercalciuria X-linked recessive hypophosphatemia (Dent’s disease) Autosomal dominant hypophosphatemia Fibrous dysplasia of bone Acquired Tumor-induced osteomalacia Renal tubular disorders Normal mineral availability Primary mineralization defect Hypophosphatasia Drug-induced Abnormal matrix synthesis Fibrogenesis imperfecta ossium

OC

Serum

Bone formation markers

Table III. Expected/Documented Levels of Bone Turnover Markers in the Rachitic and Osteomalacic Disorders

Chapter 44 Osteomalacia and Rickets

745

746 Table IV.

MARC K. DREZNER Expected Alterations in Bone Density and Histomorphometry in the Rachitic and Osteomalacic Diseases Bone histomorphometry

Calciopenic disorders Nutritional Calcium deficiency Vitamin D deficiency Vitamin D malabsorption Abnormal vitamin D metabolism Impaired 25-hydroxylation Impaired 1-hydroxylation Vitamin D resistance Vitamin D-dependent rickets, type II Phosphopenic disorders Nutritional Phosphate deficiency Hereditrary X-linked hypophosphatemia Hypophosphatemic rickets/hypercalciuria X-linked recessive hypophosphatemia Autosomal dominant hypophosphatemia Fibrous dysplasia of bone Acquired Tumor-induced osteomalacia Renal tubular disorders Normal mineral availability Primary mineralization defect Hypophosphatasia Drug-induced Abnormal matrix synthesis Fibrogenesis imperfecta ossium

Bone density

OS/BS

Md.S/BS

Ob.Pm/B.Pm

Oc.Pm/B.Pm

⇓ ⇓ ⇓

⇑ ⇑ ⇑

⇓ ⇓ ⇓

⇑ ⇑ ⇑

⇑ ⇑ ⇑

⇓ ⇓

⇑ ⇑

⇓ ⇓

⇑ ⇑

⇑ ⇑

⇓

⇑

⇓

⇑

⇑

⇓

⇑

⇓

N

N, ⇑1

⇓, ⇑2 ⇓ ⇓ ⇓1 ⇓

⇑ ⇑ ⇑ ⇑ ⇑

⇓ ⇓ ⇓ ⇓ ⇓

⇓, N N ⇓, N N ⇓, N

N, ⇑3 N, ⇑1 N, ⇑ N N

⇓ ⇓

⇑ ⇑

⇓ ⇓

⇓, N ⇓, N

N, ⇑4 ⇑, N

⇓ ⇓

⇑ ⇑

⇓ ⇓

N N

N N

⇓

⇑

⇓

N

N

OS/BS, osteoid covered bone surface; Md.S/BS, mineralizing bone surface; Ob.Pm/B.Pm, osteoblast covered bone perimeter; Oc.Pm/B.Pm, osteoclast covered bone perimeter. 1The increased serum 1,25(OH) D levels in this syndrome may contribute to a modest increase in osteoclast activity. 2 2Although the bone density in untreated patients is characteristically decreased, long term disease often results in increased density in the lumbar spine. 3A minimal increase in the number of osteoclasts along the bone margin is often observed but may represent the diminished mineralized trabecular bone surface. 4In the subgroup of patients with elevated urinary cyclic AMP the circulating factor may increase osteoclast activity.

histomorphometry or calcium kinetics [25]. More recently, quantitation of the collagen pyridinium cross-links, pyridinoline (hyroxylysylpyridinoline) and deoxypyridinoline (lysylpyridinoline), has provided relatively specific markers of bone resorption. Indeed, these measurements correlate exceptionally well with rates of bone resorption and far better than hydroxyproline [26, 27]. Not surprisingly, therefore, measurement of this variable in patients with vitamin D deficiency osteomalacia has been invariably elevated [28]. In addition, several studies describe an increase in serum concentrations of pyridinoline crosslinked carboxyterminal telopeptide of type 1 collagen (ICTP/S-CTX) in children with rickets [29–31] and in centenarians with low vitamin D status and osteomalacia [32]. Although urinary hydroxylysine and plasma tartrateresistant acid phosphatase are additional sensitive markers of bone resorption and osteoclast activity, respectively, their value in assessment of the calciopenic rachitic/ osteomalacic diseases has not been evaluated.

Since bone resorption and formation are generally tightly coupled, both in health and in disease states, the high turnover calciopenic disorders should manifest elevated levels of formation markers. However, reports of these measurements have demonstrated significant variability and most notably disparate ability to document a high turnover state. In this regard, assay of the N- and C-terminal procollagen I extension peptides (PINP, PICP), as well as the N-terminal pro-peptide of type III collagen (PIIINP), have revealed significant elevations in patients with calciopenic disease [30, 31, 33]. Since the release of PICP and PINP into the circulation correlates with the incorporation of collagen into the bone matrix, the elevated levels likely reflect augmented deposition of poorly mineralized osteoid at the increased number of bone-resorptive sites created by secondary hyperparathyroidism. In contrast, measurements of osteocalcin in the calciopenic disorders, albeit variable, have largely been decreased to normal [34, 35]. Since osteocalcin synthesis marks the matrix

Chapter 44 Osteomalacia and Rickets

maturation and subsequent mineralization [36, 37], this finding implies a loss of, or an altered metabolic coupling between, aspects of matrix production and mineralization in affected patients. Hence, as expected, elevated PINP and PICP marks the increased bone turnover/matrix production, while the normal or decreased osteocalcin may reflect a disturbed osteoblastic/matrix maturation and the consequent characteristic bone mineralization defect. A subset of patients with osteomalacia due to vitamin D deficiency have elevated circulating levels of osteocalcin [38]. The occurrence of this increase correlates with an elevation of the serum 1,25(OH)2D concentration, a finding in early stages of vitamin D deficiency, when increased circulating levels of parathyroid hormone enhance production of this active vitamin D metabolite. With progression of the disease and progressive decline of serum 25(OH)D levels, the serum osteocalcin returns to normal, again reflecting poor mineralization of the collagen. However, with treatment of vitamin D deficiency osteomalacia of all types there is a rapid increase in serum osteocalcin, signaling the resolution of a reversible delay in osteoblast maturation [35]. In any case, assay of bone markers may play an important role in the diagnosis and treatment of the calciopenic rachitic/osteomalacic disorders. In many cases establishing a diagnosis of vitamin D deficiency osteomalacia presents formidable problems. Often the 25-hydroxyvitamin D level is not overtly decreased but remains in the low-normal range. Similarly, evidence for secondary hyperparathyroidism is marginal. Documentation of increased bone turnover, therefore, may facilitate diagnosis and certainly place the disease amongst the calciopenic disorders. Moreover, successful treatment of these diseases may be easily tracked, not only by following various biochemistries, such as serum calcium, phosphorus, 25-hydroxyvitamin D, and PTH, but by serial determination of bone markers to assure that therapeutic intervention has normalized bone turnover, albeit resolution of these abnormalities occurs more slowly than the calciopenia and secondary hyperparathyroidism. In addition, with normal mineralization, the measurement of serum osteocalcin will most commonly exhibit transient overt elevation [35].

V. PHOSPHOPENIC RICKETS AND OSTEOMALACIA Rickets and osteomalacia occur in association with a variety of disorders in which phosphate depletion predominates (Table I). Most typically, these diseases have in common abnormal proximal renal tubular function, which results in an increased renal clearance of inorganic phosphorus and consequent hypophosphatemia. However, the

747 biochemical abnormalities characteristic of these disorders are quite variable (Table II). Typically these disorders are marked by a normal serum calcium concentration, a decreased serum phosporus level secondary to renal phosphate wasting and normal parathyroid function. In addition, in most forms of the disease, the urinary calcium is decreased or low-normal and the gastrointestinal absorption of calcium and phosphorus mildly compromised. Affected children often manifest substantial elevations of alkaline phosphatase activity as well, while this variable is often, but not invariably, normal in affected adults. In all forms of the disease the serum 25-hydroxyvitamin D concentration remains normal, heralding the absence of vitamin D deficiency. However, secondary to an aberration of vitamin D metabolism, the serum calcitriol level is often overtly decreased or inappropriately normal in the presence of hypophosphatemia. In contrast, rachitic/osteomalacic disease due to nutritional phosphate deficiency, X-linked recessive hypophosphatemia, or hereditary hypophosphatemic rickets with hypercalciuria occurs despite an elevated serum calcitriol level. The recent molecular definition of three rare inherited hypophosphatemic rachitic/osteomalacic disorders has led to the identification of three key regulators affecting phosphate homeostasis, PHEX, FGF-23, and GALNT3. Lossof-function mutations in PHEX are the cause of X-linked hypophosphatemia in man and the murine homolog of the disease, the hyp-mouse [39–43]; gain-of-function mutations in FGF-23 are the cause of autosomal dominant hypophosphatemic rickets [44–46], while loss-of-function mutations in this gene are the cause of a familial form of tumoral calcinosis [47]; and loss-of-function mutations in GALNT3, which codes an enzyme involved in O-glycosylation, are the cause of a second form of tumoral calcinosis and the hyperostosis-hyperphosphatemia syndrome. Furthermore, FGF-23 is vastly overexpressed in tumors causing oncogenic osteomalacia [48–52]. As a result of these studies and others it has been established that FGF-23, both in vivo and in vitro, inhibits renal phosphate reabsorption and thereby provokes phosphate wasting and hypophosphatemia. In addition, to some extent this factor directly inhibits bone mineralization. Several other phosphaturic factors, overexpressed in tumors causing oncogenic osteomalacia, have been identified and include soluble frizzle-related protein 4 (sFRP-4) and matrix extracellular phosphoglycoprotein (MEPE) [45, 46]. Like FGF-23, recombinant sFRP-4 and MEPE both inhibit phosphate transport in vitro and are phosphaturic in vivo with potencies similar to those of FGF-23 and PTH [53, 54]. Interestingly, a C-terminal fragment of MEPE, comprising the ASARM motif that is also found in several closely related proteins, inhibits mineralization in vivo, and this fragment may be similar

748 to a small oncogenic osteomalacia tumor-derived protein that inhibits phosphate uptake in OK cells [54, 55]. Furthermore, MEPE forms, through the ASARM motif, a non-proteolytic protein complex with PHEX [56]. If this interaction between MEPE and PHEX/Phex is lost in osteoblasts of patients with X-linked hypophosphatemia, due to the PHEX mutation, impaired bone mineralization would result. In any case, consistent with a role in the bone mineralization process, MEPE gene expression is markedly increased during osteoblast-mediated matrix mineralization and elevated MEPE mRNA levels are observed in bone and osteoblasts derived from hyp-mice [57, 58]. Despite the potential role that MEPE and sFRP-4 may play in phosphate homeostasis, unlike FGF-23, the circulating concentrations of these proteins are not elevated in oncogenic osteomalacia [53, 59]. More recently, the identification of the role that these phosphatonins play in well known genetic diseases of phosphate homeostasis has led to discovery of new hypophosphatemic rachitic/osteomalacic disease forms. In this regard, Whyte et al. (personal communication) have described a patient with hypophosphatemic rickets and craniosynostosis, apparently caused by activating mutations in FGF receptor 1, raising the possibility that FGF-23 effects its actions at the kidney through this receptor. Similarly, Riminucci et al. [60] have linked the hypophosphatemic osteomalacia observed in a subgroup of patients with fibrous dysplasia to increased bone production and serum levels of FGF-23. Since secondary hyperparathyroidism is not a part of the phosphopenic rachitic/osteomalacic disorders, the radiographic abnormalities and bone histomorphometry of these diseases are distinctly different from those of the calciopenic diseases. In this regard, whereas aberrant mineralization of cartilage in X-linked hypophosphatemia is marked by widening of the growth plate and diminished calficification of the metaphysis, the metaphyseal margin is more distinct than in vitamin D deficiency rickets and there is less expansion in the width of the metaphysis. Indeed, in some cases rachitic changes are not radiographically apparent [61]. In concert, bone X-rays generally display decreased bone density but there is no subperiosteal resorption or bone cysts. In addition, patients with phosphopenic disorders of long duration often display Looser’s zones in sites different to those affected in calciopenic diseases. Most commonly such defects are present in the outer cortex of the bowed femur but they may occur along the medial cortex of the shaft. However, Looser’s zones in the ribs and pelvis are rare. While these pseudofractures may heal with appropriate treatment, as they do in vitamin D deficiency, often long-term defects persist despite therapy, due most likely to fibrous tissue deposition and no subsequent mineralization. In any case, although untreated patients with phosphopenic disease consistently demonstrate these

MARC K. DREZNER

radiographic abnormalities, which are distinctly different from those encountered in calciopenic rickets/osteomalacia, long-term treatment with large doses of phosphate, alone or in combination with vitamin D or calcitriol, may induce secondary hyperparathyroidism. Under these circumstances, the X-ray picture of incompletely healed rickets/ osteomalacia may be confused by subperiosteal erosions in the hand [62]. Genetic phosphopenic disorders such as X-linked hypophosphatemia also manifest a variety of unique radiographic findings. Although there is defective mineralization of osteoid in this disease, the long-term accumulation of unmineralized tissue on bone surfaces (and the absence of secondary hyperparathyroidism) results in increased bone mass and commonly a paradoxically increased bone density with a coarse and prominent trabecular pattern [63]. This appearance is not related to treatment and is most evident in the axial skeleton. Additionally, in this disease, the bones display developmental abnormalities such as decreased length and widening of the shafts in long bones. The histologic appearance of trabecular bone in children and adults with most phosphopenic diseases (Table III) reflects the presence of a low turnover osteomalacia [64–68]. In this regard, patients with X-linked hypophosphatemia have bone biopsies that display a marked reduction in the osteoblastic calcification rate, which results in prolongation of the mineralization lag time and increased duration of the formation period. In addition, a severe reduction of the extent of osteoblast-covered osteoid and of the tetracycline double-labeled areas shows that the number of active bone-forming centers is decreased, indicating that the birth rate of new bone metabolic units is profoundly depressed in the trabecular bone envelope. In contrast, the osteoclastic surface and the osteoclast number are normal or slightly increased, but this likely reflects a prolongation of the resorption period rather than normal or increased osteoclastic activity. Indeed, the normal to increased calcified bone volume present in affected patients indicates that the observed low bone formation rate is associated with an even more reduced rate of bone resorption. Hence, with a normal to increased extent of osteoclastic surface, the duration of the resorption phase must be as prolonged as the formation phase in order to maintain reasonably normal bone volume. A similar histomorphometric appearance characterizes the bone biopsies in a majority of patients with oncogenic osteomalacia [69, 70] and autosomal dominant hypophosphatemic rickets [71], consistent with the bone histology evident in murine models of the hypophosphatemic diseases, in which transgenic over-expression of FGF-23 is manifest [72, 73]. However, a few patients with this syndrome have tumors that secrete a non-parathyroid hormone factor(s), which activates adenylate cyclase, increases urinary cyclic AMP, and increases

Chapter 44 Osteomalacia and Rickets

bone turnover with changes similar to those encountered due to secondary hyperparathyroidism [74–76]. Similarly, patients with acquired forms of the phosphopenic rickets/ osteomalacia, such as those with Fanconi’s syndrome, and those with Dent’s disease, may exhibit increased bone resorption on bone biopsy, possibly secondary to moderate renal failure which may be part of their primary disease [77].

A. Bone Markers Available data regarding bone markers in the phosphopenic disorders remain remarkably sparse. However, for the majority of these diseases the expected levels of the various markers seem reasonably predictable (Table IV). In this regard, the normal or decreased bone turnover reflected in both roentgenograms of affected patients and histomorphometric analysis of bone biopsies dictates that serum and urine markers of bone resorption should be normal or mildly decreased. Measurement of urinary hydroxyproline excretion in children and adults with X-linked hypophosphatemia and oncogenic osteomalacia confirms this expectation (unpublished observations). However, as related above, sensitivity of this variable as a marker of bone resorption is limited. Similar assessment of urinary pyridinoline and deoxypyridinoline cross-links likewise reveals normal values (unpublished observations). However, assay of serum ICTP and TRAP is not available. In a small subset of patients with phosphopenic disease evidence of increased bone turnover is expected. In this regard, Shane et al. [74] have reported frankly elevated pyridinium cross-link excretion in a patient with oncogenic osteomalacia secondary to a non-parathyroid hormone factor which increases urinary cyclic AMP excretion. Similar data are expected, but not available, in three other patients with this disease and coincident elevation of urinary cyclic AMP [75, 76], one with coexistent hyperparathyroidism [78]. As noted above, the presence of secondary hyperparathyroidism in patients with renal failure and various forms of Fanconi’s syndrome or Dent’s disease would likewise predispose to evidence of increased bone resorption and consequently elevated urinary levels of markers, such as hydroxyproline and pyridinoline cross-links. Somewhat surprisingly, a similar increment in indices of bone resorption is not expected in patients with hereditary hypophosphatemic rickets with hypercalciuria. In these patients with significantly elevated serum 1,25(OH)2D levels, increased osteoclast activity secondary to the effects of the circulating calcitriol seems reasonable. However, extensive histomorphometric analysis of bone biopsies in affected patients clearly documents normal osteoclast activity, possibly due to the counter-balancing effects of suppressed parathyroid function [79].

749 In any case, documentation of normal bone turnover in the majority of patients with phosphopenic rickets/ osteomalacia provides a valuable diagnostic tool in establishing the presence of such a disorder. In this regard, hypophosphatemia is a common abnormality in patients with calciopenic and phosphopenic disease, while a calcium abnormality is often marginal in vitamin D-deficient disorders. Hence, the ability to detect normal bone resorption may support the diagnostic impression of a phosphopenic disease. Alternatively, the rare variant of increased bone turnover in a phosphopenic disorder may direct study of patients and lead to diagnosis of oncogenic osteomalacia and/or lead to the discovery of paraneoplastic factors with the potential to increase bone turnover and impact renal function through parathyroid hormone receptors. The expected levels of those bone markers reflecting bone formation in the phosphopenic diseases is decidedly less certain (Table IV). While the rate of bone formation seems undeniably decreased upon histomorphometric exam of bone biopsies of affected patients, several factors preclude the certainty that appropriate markers will mirror this status. In this regard, the presence of decreased bone turnover should result in decreased/normal serum levels of PICP and PINP, markers of collagen deposition. However, the potential that insufficient mineralization of bone matrix may increase the turnover of collagen [50] raises the possibility that these markers may be elevated in the phosphopenic disorders. In this regard, Ros et al. [80] reported increased PINP levels in a majority of patients with phosphopenic disease. In addition, Saggese et al. [81] reported an approximately five-fold elevation in PICP in patients with active rachitic/osteomalacic disease despite treatment with calcitriol and phosphorus. While this may have reflected a relatively enhanced bone formation secondary to the combination therapy in use, this seems unlikely since poor mineralization of bone matrix persisted. In any case, further data are clearly necessary to substantiate that PINP and PICP are elevated. In contrast, the limited data available regarding another measure of collagen turnover and hence potentially bone formation, serum PIIINP levels, indicate a significant reduction of this marker in patients with X-linked hypophosphatemia [79]. This aminoterminal propeptide of type II procollagen, however, is released by soft connective tissue and may reflect information about somatic growth, most particularly growth rate [81]. Thus, the depressed level observed in youths with X-linked hypophosphatemia may not represent a decreased bone formation rate but rather mark the poor growth of the affected patients. Limited data are available regarding serum osteocalcin levels in untreated patients with phosphopenic disease. However, in a limited number of children with X-linked hypophosphatemia, Whyte (personal communication) has

750

MARC K. DREZNER

found normal and Ros et al. [80] decreased serum calcitonin, consistent with the poor mineralization and the normal to decreased bone formation rate associated with this disease. Regulation of osteocalcin in X-linked hypophosphatemia, however, is a matter of debate. In hyp-mice, the murine homolog of the human disorder, serum osteocalcin levels are invariably elevated and are inappropriately decreased in response to 1,25(OH)2D administration [82]. While these findings suggest that osteoblastic activity in this mutation is abnormal, the absence of a similar marker of osteoblast dysfunction in affected humans remains poorly understood. The effects of treatment on expression of these bone markers in patients with phosphopenic disease is unknown. Traditional therapy includes use of oral phosphate supplements and calcitriol. A common complication is secondary hyperparathyroidism, likely due to phosphate mediated transient reductions in the serum calcium concentration. The occurrence of this abnormality, however, depends upon whether the calcitriol administered is sufficient to suppress parathyroid function and/or maintain the serum calcium levels. In any case, the hyperparathyroidism, when present, is generally expressed as a biochemical phenomenon manifest by an elevated concentration of parathyroid hormone without hypercalcemia. Moreover, bone biopsies from such affected patients do not exhibit overt evidence of increased bone turnover. Thus, it is difficult to predict if therapy will reveal evidence of increased bone resorption. Nevertheless, future studies to determine this possibility are extremely important as they will provide important information regarding the potential negative impact of the traditional therapeutic intervention. Similar confusion surrounds the effects of therapy on bone formation markers. In fact, the increased circulating level of 1,25(OH)2D, which occurs upon treatment, adds further doubt to the potential alterations in these markers, since osteocalcin may increase in response to such a stimulus. In any case, identification of the panorama of changes that do occur in bone markers upon successful therapy of phosphopenic rickets/osteomalacia is evidently of considerable importance, as it will provide a relatively noninvasive means to judge therapeutic efficacy in diseases generally remarkably refractory to treatment.

VI. NORMAL MINERAL RICKETS AND OSTEOMALACIA Rickets and osteomalacia occur infrequently in the absence of a reduced serum calcium and/or phosphorus concentration. Typically abnormal mineralization in these disorders is due to an intrinsic bone defect (in which

apparently abnormal matrix is produced) or to a circulating inhibitor of mineralization. Matrix abnormalities result from production of disordered collagen or other proteins in the matrix or expression of aberrant enzyme activity essential for normal mineralization. Fibrogenesis imperfecta ossium and axial osteomalacia are examples of diseases marked by abnormal collagen production (Table I). The former is a rare, sporadically occurring disorder characterized by the gradual onset of intractable skeletal pain in middle-aged men and women. Pathologic fractures are a prominent clinical feature, and patients typically become bedridden. Although the serum calcium and phosphorus are normal, alkaline phosphatase is invariably elevated. The bones have a dense, amorphous, mottled appearance radiologically and a disorganized arrangement of collagen with decreased birefringence histologically. Biopsy sections reveal no evidence of increased bone resorption. Axial osteomalacia generally affects only middle-aged men and the majority of patients present with vague dull chronic axial discomfort that usually affects the cervical region most severely. Abnormal radiographic findings are limited to the pelvis and spine, where the coarsened trabecular pattern is characteristic of osteomalacia. Although the alkaline phosphatase may be increased, histopathologic studies reveal a normal lamellar pattern of collagen and no increased resorption. However, marked thickening of the cortices and increased trabecular bone volume is evident [83] despite the osteoblasts appearing flat and inactive, suggesting that an osteoblastic defect and perhaps attendant abnormal matrix inhibit normal mineralization. Hypophosphatasia is an inheritable disorder in which a deficiency of the tissue-non-specific (liver/bone/kidney) isoenzyme of alkaline phosphatase results in the mineralization defect characteristic of rickets and osteomalacia. The severity of clinical expression is remarkably variable and spans intrauterine death from profound skeletal hypomineralization at one extreme to lifelong absence of symptoms at the other. As a consequence, six clinical disease types are distinguished (Table I). The age at which skeletal disease is initially noted delineates, in large part, the perinatal (lethal), infantile, childhood, and adult variants of the disorder. However, affected children and adults may manifest only the unique dental abnormalities of the syndrome and accordingly are classified as having odontohypophosphatasia. Finally, patients with the rare variant, pseudohypophosphatasia, have the clinical/radiologic/ biochemical features of the classic disease without a decrease in the circulating levels of alkaline phosphatase. These individuals have defects in cellular localization and substrate specificity of the enzyme. Affected infants exhibit hypercalcemia, hypercalciuria, and often hyperphosphatemia, as well as enlarged sutures of the skull,

751

Chapter 44 Osteomalacia and Rickets

craniostenosis, delayed dentition, enlarged epiphyses, and prominent costochondral junctions. Genu valgum or varum may develop subsequently. In older children disease may be limited to rickets. Surprisingly, the disorder in adults is mild despite the presence of osteopenia. Nevertheless, 50% of patients have a history of early exfoliation of deciduous teeth and/or rickets. The physiological basis for the bone disease likely relates to the role of alkaline phosphatase in cleaving pyrophosphate, an inhibitor of bone mineralization. Failure to hydrolyze this physiologic substrate results in inorganic pyrophosphate elevated to levels sufficiently high to inhibit the mineralization process. The consequence of this pathophysiological process is a block of the vectorial spread of mineral from initial nuclei within matrix vesicles outwards into the matrix of growth cartilage and bone. However, bone histomorphometry reveals no evidence of abnormal resorption or formation. Among the circulating inhibitors of bone mineralization responsible for rickets/osteomalacia are bisphosphonates, fluoride, and aluminum. The bone diseases resultant from exposure to these agents are highly variable, reflecting decreased resorption and formation secondary to aluminum and increased formation due to fluoride. While the serum calcium and phosphorus are routinely normal, the alkaline phosphatase is invariably elevated.

A. Bone Markers Since the majority of rachitic/osteomalacic disorders that occur without a reduced serum calcium or phosphorus concentration are marked by normal rates of bone turnover, investigators anticipate that markers of bone resorption and formation will fluctuate in the normal range. Thus, such tests can serve as supportive in efforts to ascertain the cause of a bone mineralization disorder. However, limited data are available to support such anticipation. Indeed, in select forms of normal mineral rickets/osteomalacia abnormalities occur which may variably influence bone marker levels. For example, the hypercalcemia often seen in children with hypophosphatasia may suppress parathyroid hormone secretion and decrease bone resorption, thereby depressing levels of urinary pyridinoline crosslinks among other tests. However, Girschick et al. [84] recently reported that hydroxyproline and free deoxypyridinoline excretion were normal in six children with childhood hypophosphatasia. Similarly, aluminum sequestration in parathyroid glands may limit parathyroid hormone secretion and effects on osteoblasts reduce collagen production. The net effect is likely the low turnover osteomalacia characteristically associated with aluminum exposure. Under such conditions profound decreases in markers of resorption and formation

are anticipated, provided that the often associated renal failure does not influence serum levels of these measures. Nevertheless, increased urine hydroxyproline excretion has been reported in a single patient with aluminum-induced osteomalacia [85]. In contrast to the aforementioned, osteomalacia secondary to fluoride ingestion is marked by increased osteoblast activity. Indeed, in an animal model of this disease bone resorption and formation were increased by 30% and 40%, respectively [86]. Thus, the possibility of increased levels of PINP and PICP are likely, correlating with enhanced incorporation of collagen into the bone matrix. However, the attendant defective mineralization may result in depressed serum osteocalcin levels, similar to that seen in calciopenic disorders in particular. The current availability of measurement for the bone markers should provide valuable tools to better understand the heterogeneous group of disorders that comprise the normal mineral rachitic and osteomalacic disorders.

References 1. Park, E. A. (1939). Observations on the pathology of rickets with particular reference to the changes at the cartilage-shaft junctions of the growing bones. Bull. NY Acad. Med. 15, 495–543. 2. Baylink, D., Stauffer, M., Wergedal, J., and Rich, C. (1970). Formation, mineralization and resorption of bone in vitamin D-deficient rats. J. Clin. Invest. 49, 1122–1134. 3. Stanbury, S. W., and Mawer, E. B. (1990). Metabolic disturbances in acquired osteomalacia. In: “The Metabolic and Molecular Basis of Acquired Disease” (R. D. Cohen, B. Lewis, K. G. M. M. Alberti, and A. M. Denman, eds.), pp. 1717–1782. Bailliere Tindall, London, UK. 4. Rao, D. S., Villanueva, A., Mathews, M., Bumo, B., Frame, B., Kleerekoper, M., and Parfitt, A. M. (1983). Histologic evolution of vitamin D depletion in patients with intestinal malabsorption or dietary deficiency. In: “Clinical Disorders of Bone and Mineral Metabolism” (B. Frame, and J. T. Potts, eds.), pp. 224–226. Excerpta Medica, Amsterdam. 5. Fraser, D., Kooh, S. W., Kind, H. O., Holick, M. F., Tanaka, Y., and DeLuca, H. F. (1973). Pathogenesis of hereditary vitamin D dependent rickets: an inborn error of vitamin D metabolism involving defective conversion of 25-hydroxyvitamin D to 1,25-dihydroxyvitamin D. N. Engl. J. Med. 289, 817–822. 6. Marx, S. J., Liberman, U. A., Eil, C., Gamblin, G. T., DeGrange, D. A., and Balsan, S. (1984). Hereditary resistance to 1,25-dihydroxyvitamin D. Recent Prog. Horm. Res. 40, 589–620. 7. Silva, I., Cebotaru, V., Wang, H., Wang, X., Wang, S. S., Guo, G., Devuyst, O., Thakker, R. V., Guggino, W. B., and Guggino, S. E. (2003). The ClC-5-knockout mouse model of Dent’s disease has renal hypercalciuria and increased bone turnover. J. Bone Min. Res. 18, 615–623. 8. Robey, P. G., and Boskey, A. L. (1996). The biochemistry of bone. In: “Osteoporosis” (R. Marcus, D. Feldman, and J. Kelsey, eds.), pp. 95–183. Academic Press, San Diego, CA. 9. Glimcher, M. J. (1992). The nature of the mineral component of bone and the mechanism of calcification. In: “Disorders of Bone and Mineral Metabolism” (F. L. Coe, and M. J. Favus, eds.), pp. 265–286. Raven, New York.

752 10. Neuman, W. F. (1980). Bone material and calcification mechanisms. In: “Fundamental and Clinical Bone Physiology” (M. R. Urist, ed.), pp. 83–107. Lippincott, Philadelphia, PA. 11. Christoffersen, J., and Landis, W. J. (1991). A contribution with review to the description of mineralization of bone and other calcified tissues in vivo. Anat. Rec. 230, 435–450. 12. Thorngren, K. G., Hansson, L. I., Menander-Sellman, K., and Stenstrom, A. (1973). Effect of hypophysectomy on longitudinal bone growth in the rat. Calcif. Tissue Res. 11, 281–300. 13. Caplan, A. I., and Boyan, B. D. (1994). Endochondral bone formation: The lineage cascade. In: “Bone: A Treatise” (B. K. Hall, ed.), Vol. 8: Mechanisms of Bone Development and Growth, pp. 1–46. CRC Press, Boca Raton, FL. 14. Boskey, A. L. (1990). Mineral-matrix interactions in bone and cartilage. Clin. Orthop. Related Res. 281, 244–274. 15. Parfitt, A. M. (1997). Vitamin D and the Pathogenesis of Rickets and Osteomalacia. In: “Vitamin D” (D. Feldman, F. H. Glorieux, and J. W. Pike, eds.), pp. 645–662. Academic Press, San Diego, CA. 16. Rudolf, M., Arulanantham, K., and Greenstein, R. M. (1980). Unsuspected nutritional rickets. Pediatrics 66, 72–76. 17. Edidin, D. V., Levitsky, L. L., Schey, W., Dumbuvic, N., and Campos, A. (1980). Resurgence of nutritional rickets associated with breastfeeding and special dietary practices. Pediatrics 65, 232–235. 18. Bachrach, S., Fisher, J., and Parks, J. S. (1979). An outbreak of vitamin D deficiency rickets in a susceptible population. Pediatrics 64, 871–877. 19. Hanley, D. A., and Davison, K. S. (2005). Vitamin D insufficiency in North America. J. Nutr. 135, 332–337. 20. Henderson, A. (2005). Vitamin D and the breastfed Infant. J. Obstet. Gynecol. Neonatal. Nurs. 34, 367–372. 21. Bonnici, F. (1978). Functional hypoparathyroidism in infantile hypocalcaemic stage I vitamin D deficiency rickets. S. Afr. Med. J. 54, 611–312. 22. Buchanan, N., Pettifor, J. M., Cane, R. D., and Bill, P. L. A. (1978). Infantile apnoea due to profound hypocalcaemia associate with vitamin D deficiency. S. Afr. Med. J. 54, 766–767. 23. Schott, G. D., and Wills, M. R. (1976). Muscle weakness in osteomalacia. Lancet 1, 626–629. 24. Kruse, K. (1995). Pathophysiology of calcium metabolism in children with vitamin D-deficiency rickets. J. Pediatr. 126, 736–741. 25. Delmas, P. D. (1990). Biochemical markers of bone turnover for the clinical assessment of metabolic disease. Endocrinol. Metab. Clin. North Am. 19, 1–18. 26. Eastell, R., Hampton, L., Colwell, A., Green, J. R., Assiri, A. M. A., Hesp, R., Russell, R. G. G., and Reeve, J. (1990). Urinary collagen crosslinks are highly correlated with radioisotopic measurement of bone resorption. In: “Osteoporosis 1990” (C. Christiansen, and K. Overquaard, eds.), Vol. 2, pp. 469–470. Osteopress, Aalborg, Denmark. 27. Delmas, P. D., Schenmer, A., Gineyts, E., Riis, B., and Christiansen, C. (1991). Urinary excretion of pyridinoline crosslinks correlate with bone turnover measured on iliac crest biopsy in patients with vertebral osteoporosis. J. Bone Min. Res. 6, 639–644. 28. Robins, S. P., Black, D., Paterson, C. R., Reid, D. M., Duncan, A., and Seibel, M. J. (1991). Evaluation of urinary hydroxypyridinium crosslink measurements as resorption markers in metabolic bone disease. Eur. J. Clin. Invest. 21, 310–315. 29. Scariano, J. K., Walter, E. A., Glew, R. H., Hollis, B. W., Henry, A., Ocheke, I., and Isichei, C. O. (1995). Serum levels of the pyridinoline crosslinked carboxyterminal telopeptide of type I collagen (ICTP) and osteocalcin in rachitic children in Nigeria. Clin. Biochem. 28, 541–545. 30. Sharp, C. A., Oginni, L. M., Worsfold, M., Oyelami, O. A., Risteli, L., Risteli, J., and M. W. J. D. (1997). Elevated collagen turnover in Nigerian children with calcium-deficiency rickets. Calcif. Tissue Res. 61, 87–94. 31. Li, F., Iqbal, J., Wassif, W., Kaddam, I., and Moniz, C. (1994). Carboxyterminal propeptide of type I procollagen in osteomalacia. Calcif. Tissue Res. 55, 90–93.

MARC K. DREZNER 32. Passeri, G., Pini, G., Troiano, L., Vescovini, R., Sanson, P., Passeri, M., Guerresi, P., Delsignore, R., Pedrazzoni, M., and Franceschi, C. (2003). Low Vitamin D status, high bone turnover and bone fractures in centenarians. J. Clin. Endocrinol. Metab. 88, 5109–5115. 33. Saggese, G., Bertelloni, S., Baroncelli, G. I., and Di Nero, G. (1992). Serum levels of carboxyterminal propeptide of type I procollagen in healthy children from 1st year of life to adulthood and in metabolic bone diseases. Eur. J. Pediatr. 151, 764–768. 34. Abitbol, V., Roux, C., Chaussade, S., Guillemant, S., Kolta, S., Dougados, M., Couturier, D., and Amor, B. (1995). Metabolic bone assessment in patients with inflammatory bowel disease. Gastroenterology 108, 417–422. 35. Daniels, E. D., Pettifor, J. M., and Moodley, G. P. (2000). Serum osteocalcin has limited usefulness as a diagnostic marker for rickets. Eur. J. Pediatr. 159, 730–733. 36. Price, P. A. (1983). The osteoblast. In: “Primer on the Metabolic Bone Diseases and Disorders of Mineral Metabolism” (M. J. Favus, ed.), pp. 15–21. Princeton University, Princeton, NJ. 37. Heinrichs, A. A. J., Bortell, R., Bourke, M., Lian, J. B., Stein, G. S., and Stein, J. L. (1995). Proximal promoter binding protein contributes to developmental tissue-restricted expression of the rat osteocalcin gene. J. Cell. Biochem. 57, 90–100. 38. Delmas, P. (1990). Biochemical markers of bone turnover for the clinical assessment of metabolic bone disease. In: “Metabolic Bone Disease, Part 2” (R. D. Tiegs, ed.), pp. 1–8. Saunders, Philadelphia, PA. 39. Holm, I. A., Huang, X., and Kunkel, L. M. (1997). Mutational analysis of the PEX gene in patients with X-linked hypophosphatemic rickets. Am. J. Hum. Genet. 60, 790–797. 40. Carpinelli, M. R., Wicks, I. P., Sims, N. A., O’Donnell, K., Hanzinikolas, K., Burt, R., Foote, S. J., Bahlo, M., Alexander, W. S., and Hilton, D. J. (2002). An ethyl-nitrosourea-induced point mutation in phex causes exon skipping, x-linked hypophosphatemia, and rickets. Am. J. Pathol. 161, 1925–1933. 41. Lorenz-Depiereux, B., Guido, V., Johnson, K., Zheng, Y., Gagnon, L., Bauschata, J., Davison, M. W., L., Donahue, L., Strom, T., and Eicher, E. M. (2004). Two new spontaneous intragenic deletions in the mouse Phex gene and a biological characterization of mice carrying these and two previously identified Phex mutations. Mamm. Genome. 42. The HYPConsortium. (1995). A gene (PEX) with homologies to endopeptidases is mutate in patients with X-linked hypophosphatemic rickets. Nat. Genet. 11, 130–136. 43. Strom, T. M., Francis, F., Lorenz, B., Boddrich, A., Econs, M. J., Lehrach, H., and Meitinger, T. (1997). Pex gene deletions in Gy and Hyp mice provide mouse models for X-linked hypophosphatemia. Hum. Mol. Genet. 6, 165–171. 44. (2000). Autosomal dominant hypophosphataemic rickets is associated with mutations in FGF23. The ADHR Consortium. Nat. Genet. 26, 345–348. 45. Shimada, T., Mizutani, S., Muto, T., Yoneya, T., Hino, R., Takeda, S., Takeuchi, Y., Fujita, T., Fukumoto, S., and Yamashita, T. (2001). Cloning and characterization of FGF23 as a causative factor of tumor-induced osteomalacia. Proc. Natl Acad. Sci. USA 98, 6500–6505. 46. Jan de Beur, S. M., Finnegan, R. B., Vassiliadis, J., Cook, B., Barberio, D., Estes, S., Manavalan, P., Petroziello, J., Madden, S., Cho, J. Y., Kumar, R., Levine, M. A., and Schiavi, S. C. (2002). Tumors associated with oncogenic osteomalacia express markers of bone and mineral metabolism. J. Bone Miner. Res. 17, 1102–1110. 47. Benet-Pages, A., Orlik, P., Strom, T., and Lorenz-Depiereux, B. (2004). An FGF23 missense mutation causes familial tumoral calcinosis with hyperphosphatemia. Hum. Mol. Genet. [Epub ahead of print]. 48. Jonsson, K. B., Zahradnik, R., Larsson, T., White, K. E., Hampson, G., Miyauchi, A., Econs, M., Lavigne, J., and Juppner, H. (2002). FGF-23 is a circulating factor that is elevated in oncogenic osteomalacia and X-linked hypophosphatemic rickets. N. Engl. J. Med. 348, 1656–1663.

Chapter 44 Osteomalacia and Rickets 49. Yamazaki, Y., Okazaki, R., Shibata, M., Hasegawa, Y., Satoh, K., Tajima, T., Takeuchi, Y., Fujita, T., Nakahara, K., Yamashita, T., and Fukumoto, S. (2002). Increased circulatory level of biologically active full-length FGF-23 in rickets/osteomalacia. J. Clin. Endocrinol. Metab. 87, 4957–4960. 50. Nelson, A. E., Bligh, R., Mirams, M., Gill, A., Au, A., Clarkson, A., Jueppner, J., Ruff, S., Stalley, P., Scolyer, R., Bobinson, B., Mason, R. S., and Bligh, P. (2003). FGF23: a new clinical marker for oncogenic osteomalacia. J. Clin. Endocrinol. Metab. 88, 4088–4094. 51. Ward, L., Rauch, F., White, K. E., Filler, G., Matzinger, M., Letts, M., Travers, R., Econs, M., and Glorieux, F. H. (2004). Resolution of severe, adolescent-onset hypophosphatemic rickets following resection of an FGF-23-producing tumour of the distal ulna. Bone 34, 905–911. 52. Mirams, M., Robinson, B. G., Mason, R., and Nelson, A. (2004). Bone as a source of FGF23: regulation by phosphate? Bone 35, 1192–1199. 53. Berndt, T., Craig, T. A., Bowe, A. E., Vassiliadis, J., Reczek, D., Finnegan, R., Jan de Beur, S. M., Schiavi, S. C., and Kumar, R. (2003). Secreted frizzled-related protein 4 is a potent tumor-derived phosphaturic agent. J. Clin. Invest. 112, 785–794. 54. Rowe, P. S. N., Kumagai, Y., Gutierrez, G., Garrett, I. R., Blacher, R., Rosen, D., Chen, D., Drezner, M. K., Quarles, L. D., and Mundy, G. R. (2003). MEPE regulates bone mineralization and phosphate transport: PHEX and the MEPE ASARM-peptide. Bone, In press. 55. Jonsson, K., Mannstadt, M., Miyauchi, A., Yang, I. M., Stein, G., Ljunggren, O., and Juppner, H. (2001). Extracts from tumors causing oncogenic osteomalacia inhibit phosphate uptake in opossum kidney cells. J. Endocrinol. 169, 613–620. 56. Rowe, P. S. N., Garrett, I. R., Schwarz, P. M., Carnes, D. L., Lafer, E. M., Mundy, G. R., and Guiterrez, G. E. (2005). Surface plasmon resonance (SPR) confirms that MEPE binds to PHEX via the MEPE-ASARM motif: a model for impaired mineralization in X-linked rickets (HYP). Bone 36, 33–46. 57. Argiro, L., Desbarats, M., Glorieux, F. H., and Ecarot, B. (2001). Mepe, the gene encoding a tumor-secreted protein in oncogenic hypophosphatemic osteomalacia, is expressed in bone. Genomics 74, 342–351. 58. Rowe, P. S., de Zoysa, P. A., Dong, R., Wang, H. R., White, K. E., Econs, M. J., and Oudet, C. L. (2000). MEPE, a new gene expressed in bone marrow and tumors causing osteomalacia. Genomics 67, 54–68. 59. Jain, A., Fedarko, N. S., Collins, M. T., Gelman, R., Ankrom, M., Tayback, M., and Fisher, L. (2004). Serum levels of matrix extracellular phosphoglycoprotein (MEPE) in normal humans correlate with serum phosphorus, parathyroid hormone and bone mineral density. J. Clin. Endocrinol. Metab. 89, 4158–4161. 60. Riminucci, M., Collins, M., Fedarko, N., Cherman, N., Corsi, A., White, K., Waguespack, S., Gupta, A., Hannon, T., Econs, M., Bianco, P., and Gehron Robey, P. (2003). FGF-23 in fibrous dysplasia of bone and its relationship to renal phosphate wasting. J. Clin. Invest. 112, 683–692. 61. Econs, M. J., Feussner, J. R., Samsa, G. P., Effman, E. L., Vogler, J. B., Martinez, S., Friedman, N. E., Quarles, L. D., and Drezner, M. K. (1991). X-linked hypophosphatemic rickets without “rickets”. Skeletal Radiol. 20, 109–114. 62. Steinbach, H. L., Kolb, F. O., and Crane, J. T. (1959). Unusual roentgen manifestations of osteomalacia. Am. J. Roentgenol. 82, 875–886. 63. Rivkees, S. A., el-Hajj-Fuleihan, G., Brown, E. M., and Crawford, J. D. (1992). Tertiary hyperparathyroidism during high phosphate therapy of familial hypophosphatemic rickets. J. Clin. Endocrinol. Metab. 76, 1514–1518. 64. Marie, P. J., and Glorieux, F. H. (1982). Bone histomorphometry in asymptomatic adults with hereditary hypophosphatemic vitamin Dresistant osteomalacia. Metab. Bone Dis. Relat. Res. 4, 249–253.

753 65. Lyles, K. W., Burkes, E. J. Jr., McNamara, C. R., Harrelson, J. M., Pickett, J. P., and Drezner, M. K. (1985). The concurrence of hypoparathyroidism provides new insights to the pathophysiology of X-linked hypophosphatemic rickets. J. Clin. Endocrinol. Metab. 60, 711–717. 66. Drezner, M. K., Lyles, K. W., and Harrelson, J. M. (1980). Vitamin D resistant osteomalacias: Evaluation of vitamin D metabolism and response to therapy. In: “Hormonal Control of Calcium Metabolism” (D.V. Choh, R. V. Talmage, and J. L. Matthews, eds.), pp. 243–251. Excerpta Medica, Amsterdam. 67. Lyles, K. W., Harrelson, J. M., and Drezner, M. K. (1982). The efficacy of vitamin D2 and oral phosphorus therapy in X-linked hypophosphatemic rickets and osteomalacia. J. Clin. Endocrinol. Metab. 54, 307–315. 68. Drezner, M. K., Lyles, K. W., Haussler, M. R., and Harrelson, J. M. (1980). Evaluation of a role for 1,25-dihydroxyvitamin D3 in the pathogenesis and treatment of X-linked hypophosphatemic rickets and osteomalacia. J. Clin. Invest. 66, 1020–1032. 69. Drezner, M. K., and Feinglos, M. N. (1977). Osteomalacia due to 1,25-dihydroxycholecalciferol deficiency: Association with a giant cell tumor of bone. J. Clin. Invest. 60, 1046–1053. 70. Lyles, K. W., Berry, W. R., Haussler, M., Harrelson, J. M., and Drezner, M. K. (1980). Hypophosphatemic osteomalacia: Association with prostatic carcinoma. Ann. Int. Med. 93, 275–278. 71. Econs, M. J., and McEnery, P. T. (1997). Autosomal dominant hypophosphatemic rickets/osteomalacia: clinical characterization of a novel renal phosphate-wasting disorder. J. Clin. Endocrinol. Metab. 82, 674–681. 72. Larsson, T. E. M., Marsell, R., Schipani, E., Ohlsson, C., Tenenhouse, H. S., Ljunggren, O., Jueppner, J., and Jonsson, K. B. (2003). Mice transgenic for fibroblast growth factor 23 exhibit growth retardation, osteomalacia and disturbed calcium/phosphate homeostasis. J. Bone Miner. Res. 18, S283 (SU431). 73. Shimada, T., Yoneya, T., Hino, R., Takeuchi, Y., Fukumoto, S., and Tamashita, T. (2001). Transgenic mice expressing fibroblast growth factor 23 demonstrate hypophosphatemia with low serum 1,25-dihydroxyvitamin D and rickets/osteomalacia. J. Bone Miner. Res. 16(Suppl 1), S151. 74. Shane, E., Parisien, M., Henderson, J. E., Dempster, D. W., Feldman, D., Hardy, M. A., Tohme, J. F., Karaplis, A. C., and Clemens, T. L. (1997). Tumor-induced osteomalacia: clinical and basis studies. J. Bone Miner. Res. 12, 1502–1511. 75. Miller, M. J., Marel, G., and Frame, B. (1982). Adult acquired vitamin D and PTH-resistant hypophosphatemic osteomalacia with multiple skeletal lesions. In: “Vitamin D-Chemical, Biochemical and Clinical Endocrinology of Calcium Metabolism” (A. W. Norman, K. Schaefer, D. V. Herrath, and H. G. Grigoleit, eds.), pp. 993–995. Walter deGruyter, New York. 76. Leicht, E., Biro, G., and Langer, H.-J. (1990). Tumor-induced osteomalacia: Pre- and post-operative biochemical findings. Horm. Metab. Res. 22, 640–643. 77. Clarke, B. L., Wynne, A. G., Wilson, D. M., and Fitzpatrick, L. A. (1995). Osteomalacia associated with adult Fanconi’s syndrome: clinical and diagnostic features. Clin. Endocrinol. 43, 479–490. 78. Reid, I. R., Teitelbaum, S. L., Dusso, A., and Whyte, M. P. (1987). Hypercalcemic hyperparathyroidism complicating oncogenic osteomalacia: Effect of successful tumor resection on mineral homeostasis. Am. J. Med. 83, 350–354. 79. Gazit, D., Tieder, M., Liberman, U. A., Passi-Even, L., and Bab, I. A. (1991). Osteomalacia in Hereditary Hypophosphatemic Rickets with Hypercalciuria: A correlative clinical-histomorphometric study. J. Clin. Endocrinol. Metab. 71, 229–235. 80. Ros, I., Alvarez, L., Guanabens, N., Peris, P., Monegal, A., Vazquez, I., Cerda, D., Ballesta, A. M., and Munoz-Gomez, J. (2005). Hypophosphatemic osteomalacia: a report of five cases and evaluation of bone markers. J. Bone Miner. Metab. 23, 266–269.

754 81. Saggese, G., Baroncelli, G. I., Bertelloni, S., and Perri, G. (1995). Long-term growth hormone treatment in children with renal hypophosphatemic rickets: Effects on growth, mineral metabolism and bone density. J. Pediatr. 127, 395–402. 82. Carpenter, T. O., and Gundberg, C. M. (1996). Osteocalcin abnormalities in Hyp mice reflect altered genetic expression and are not due to altered clearance, affinity for mineral, or ambient phosphorus levels. Endocrinology 137, 5213–5219. 83. Demiaux-Domenech, B., Bonjour, J. P., and Rizzoli, R. (1996). Axial osteomalacia: report of a new case with selective increase in axial bone mineral density. Bone 18, 633–637.

MARC K. DREZNER 84. Firschick, H. J., Schneider, P., Kruse, K., and Huppertz, H. I. (1999). Bone metabolism and bone mineral density in childhood hypophosphatasia. Bone 25, 361–367. 85. Kassem, M., Eriksen, E. F., Melsen, F., and Mosekilde, L. (1991). Antacid-induced osteomalacia: a case report with a histomorphometric analysis. J. Intern. Med. 229, 275–279. 86. Kragstrup, J., Richards, A., and Fejerskov, O. (1984). Experimental osteo-fluorosis in the domestic pig: a histomorphometric study of vertebral trabecular bone. J. Dent. Res. 63, 885–889.

Chapter 45

Assessment of Bone and Joint Diseases: Renal Osteodystrophy Esther A. González, Ziyad Al Aly and Kevin J. Martin

I. Introduction II. Biochemical Assessment of Renal Osteodystrophy III. Skeletal Imaging in Renal Osteodystrophy

IV. Summary References

I. INTRODUCTION

The spectrum of histologic abnormalities found in patients with chronic kidney disease is, therefore, broad and is generally classified according to the major abnormalities seen on bone histology [2, 3]. The prevalence of the various types of renal osteodystrophy found in patients with end-stage renal failure is shown in Table I; the data are divided according to the type of dialysis therapy being administered. Thus, hyperparathyroid bone disease is the most common abnormality in patients on hemodialysis whereas the adynamic bone lesion is the most common finding in patients on peritoneal dialysis. Some patients have mixed patterns of bone histology that are intermediate between these two extremes and some have only mild abnormalities [3]. Abnormalities in bone histology occur early in the course of renal insufficiency [4, 5]. The earliest findings are those of hyperparathyroid bone disease resulting from high levels of parathyroid hormone in serum, which arise as a consequence of phosphate retention and decreased calcitriol production by the diseased kidney [6]. As renal

The majority of patients with end-stage renal diseases have abnormal bone histology. These skeletal abnormalities, generically termed renal osteodystrophy, encompass the manifestations of hyperparathyroidism on bone (osteitis fibrosa), a disorder of low bone turnover (adynamic bone), abnormalities of bone mineralization (osteomalacia), and combinations of these abnormalities (mixed renal osteodystrophy), together with the accumulation of retained ions such as aluminum, iron, cadmium, oxalate, as well as the deposition of proteins such as β2 microglobulin [1]. It is also important to consider that the final manifestations in bone may be tempered by prior therapy for specific renal diseases, e.g. corticosteroids, inactivity as a consequence of chronic kidney disease, ovarian failure (either due to drug therapy or uremia) as well as advancing age. These confounding factors may complicate the final pattern of bone histology and, in addition, may influence the changes over time and the responses to therapeutic interventions. Dynamics of Bone and Cartilage Metabolism

Division of Nephrology, Saint Louis University School of Medicine, St. Louis, Missouri, USA

755

Copyright © 2006 by Academic Press. All rights of reproduction in any form reserved.

756 Table I.

Esther A. González ET AL. Prevalence of Renal Osteodystrophy in Patients with ESRD Hemodialysis

Mixed ROD Osteomalacia Mild/Normal

11 2 13

Peritoneal dialysis 4 6 21

The numbers shown represent the percent of patients with each type of renal osteodystrophy according to dialysis modality. There were 117 patients in the hemodialysis group and 142 in the peritoneal dialysis group. (Modified from Sherrard et al. with permission [3].)

disease progresses, other factors such as hypocalcemia, acidosis, intrinsic alterations of parathyroid function, and skeletal resistance to the actions of PTH all contribute to aggravate the hypersecretion of parathyroid hormone [7]. The finding of osteomalacia is a diminishing feature of skeletal disease in chronic kidney disease [8] and, when present, it is usually accompanied by the accumulation of aluminum at the mineralization front [9, 10]. The major sources of aluminum in this setting include the water used for hemodialysis and the administration of aluminumcontaining phosphate binders which may be used in the treatment of secondary hyperparathyroidism. With careful monitoring of the water and the use of alternative phosphate binders, aluminum-induced osteomalacia is becoming less of a problem [8]. The pathogenesis of adynamic bone disease, which is being increasingly recognized, remains unclear [11–13]. It may be the result of over-aggressive therapy of hyperparathyroidism in some cases, but it could also represent the accumulation of inhibitors of osteoblast function occurring as a consequence of renal failure or a deficiency of stimulators of osteoblast growth or differentiation. The symptoms and signs of renal osteodystrophy are generally vague and non-specific (Table II). Bone pain, often presenting as vague aches particularly in the feet and legs, is common. Muscular aches and muscle weakness are also very common complaints. Occasional spontaneous fractures can occur, particularly in the ribs, as a result of minimal trauma or coughing. Additional manifestations Table II.

Clinical Features of Renal Osteodystrophy

Bone pain Fractures Skeletal deformities Growth retardation Muscle weakness Extraskeletal calcifications Pruritis Periarthritis Spondyloarthropathy

include skeletal deformities, particularly in children who also demonstrate impaired growth, pruritis, joint pains, carpal tunnel syndrome and extraskeletal calcifications. Because of the non-specific nature of the symptoms, it becomes important to utilize biochemical and radiological diagnostic techniques to clarify or to confirm the precise diagnosis, and to serve as a guide for administration of specific therapy. The overall framework for the assessment of the factors involved in renal osteodystrophy is depicted in Figure 1. Thus, in the setting of renal failure, a series of events takes place, the most important being phosphorus retention and calcitriol deficiency, which lead to the development of hyperparathyroidism. The high levels of PTH serve to alter the cellular events involved in the process of skeletal remodeling with the resultant adverse consequences, which lead to osteitis fibrosa. Osteitis fibrosa is characterized by an increase in bone remodeling, with increased osteoblast and osteoclast activities, and excess fibrous tissue deposition in the peritrabecular areas of the bone marrow. Accordingly, assessment of parathyroid activity is of paramount importance in the evaluation and therapy of renal osteodystrophy. PTH also regulates the serum concentrations of calcium and phosphorus, the disturbances of which may also provide information regarding PTH activity (see also Chapter 16). Because PTH acts on osteoblast precursors as well as mature osteoblasts, it would appear useful to measure parameters of osteoblast activity to assess the cellular consequences of the prevailing parathyroid activity. Thus, osteoblast markers such as alkaline phosphatase (see Chapter 9) and osteocalcin (see Chapter 3) are useful in this regard. In addition, osteoblast secretory products such as IL-6 and IL-11 produced in response to PTH interact with the osteoclast pathway to produce an increase in osteoclast number and activity and an increased bone resorption (see Chapter 7). Similarly, osteoblastic cells may secrete factors that inhibit osteoclast development and activity such as osteoprotegerin (OPG). These intermediary substances may also be useful targets for measurement to gauge the cellular activity of the skeleton. Osteoclasts resorb bone and the products of resorption may be released into the circulation and provide an index of osteoclast activity. Likewise, other osteoclast markers such as tartrate-resistant acid phosphatase (TRAP; Chapter 10) may be released into the peripheral circulation where it can be measured. The histology may be modified by other systemic factors such as acidosis, which is common in advanced renal insufficiency and leads to mobilization of bone mineral as the hydrogen ion is buffered by bone carbonate. In addition, aluminum may accumulate at the mineralization front and impair mineralization. β2-microglobulin may also accumulate in bones and joint ligaments. Assessment of the validity of these diagnostic

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Figure 1

Diagrammatic representation of a framework for the assessment of the factors involved in renal osteodystrophy. See text for complete description.

aids requires relating the abnormalities to the “gold standard” for the diagnosis of renal osteodystrophy, which is the histologic examination of undecalcified sections of bone [14]. Bone biopsy is a minimally invasive technique that is not widely utilized in clinical practice; nonetheless, it is an important tool to demonstrate the validity of non-invasive diagnostic techniques and is essential in selected cases. Non-invasive radiographic techniques may also provide information with regard to the overall density and integrity of various parts of the skeleton.

II. BIOCHEMICAL ASSESSMENT OF RENAL OSTEODYSTROPHY A. Measurements of Calcium and Phosphorus in Serum Levels of serum calcium, particularly ionized calcium, are usually normal in mild to moderate renal insufficiency and tend to fall with advanced renal insufficiency; levels only decrease below the normal range in a minority of patients [15, 16]. The maintenance of normal values, during the course of renal insufficiency, is the result of compensatory secondary hyperparathyroidism. Decreases in total serum calcium are more common in patients with proteinuric renal diseases in whom hypoalbuminemia is present, but ionized calcium may be normal. The occurrence of frank hypocalcemia is even less common if calcium-containing antacids are used as phosphate binders; however, if hypocalcemia is present, it is a powerful stimulus for PTH secretion and should be corrected. Calciminetic agents have recently been introduced for the management of secondary

hyperparathyroidism and their use may be associated with hypocalcemia [17]. Hypercalcemia is uncommon in mild or moderate renal failure but may occur in advanced renal failure or end-stage renal disease. It may occur as a consequence of the ingestion of large amounts of calciumcontaining antacids (used as phosphate binders), severe (tertiary) hyperparathyroidism, states of abnormally low bone turnover such as adynamic bone, or overtreatment with vitamin D or vitamin D analogs [1, 18, 19]. The diagnosis of the precise cause of hypercalcemia is extremely important because the therapy required differs greatly depending on the cause. Occasionally, hypercalcemia may be due to the overproduction of calcitriol or coexistent granulomatous diseases such as sarcoidosis or tuberculosis [20, 21]. The levels of serum phosphorus are usually normal or slightly decreased in early to moderate renal failure [15, 16]. Hyperphosphatemia does not become evident until renal function has declined to less than 20% of normal. Again, the maintenance of normal serum phosphorus concentration is due to the development of secondary hyperparathyroidism [22]. Efforts are usually made to prevent the development of hyperphosphatemia during the course of renal insufficiency by dietary phosphorus restriction and the use of phosphate-binding antacids. Hyperphosphatemia may aggravate hyperparathyroidism by decreasing the levels of ionized calcium, decreasing the production of calcitriol, and by a direct effect on the parathyroid gland to increase PTH secretion and regulate the growth of parathyroid cells [23–26]. In addition to its contribution to hyperparathyroidism, hyperphosphatemia has also been associated with increased risk of mortality [27, 28]. Hyperphosphatemia is common when GFR falls to very low levels unless treatment for hyperphosphatemia is pursued aggressively.

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In some instances, increased resorption of phosphorus from bone as a consequence of severe hyperparathyroidism may contribute to the development of hyperphosphatemia. Apart from the frequent development of hypercalcemia (especially after the administration of low doses of calcitriol), which is suggestive of a low bone turnover syndrome, the levels of serum calcium do not have specific diagnostic value as to the precise nature of the underlying osteodystrophy. Likewise, measurements of serum phosphorus do not indicate specific abnormalities in bone histology. These measurements should be considered in conjunction with direct measurements of PTH. Figure 2

B. Measurement of PTH Since hyperparathyroidism is a major component of renal osteodystrophy, the accurate measurement of PTH in serum is important, not only as a diagnostic tool but also as a guide to therapy (see also Chapters 16 and 32). Difficulties with different assays for PTH directed towards different regions of the PTH molecule, complicated by the dependence of the kidney for removal of hormone fragments from the circulation [29], have largely been obviated by the advent of two-site assays for intact PTH by immunoradiometric (IRMA) or immunochemiluminescence (ICMA) techniques [30, 31]. The first generation of these two-site assays, referred to as ‘‘intact’’ PTH assays are now known to detect a large hormone fragment (PTH 7-84) in addition to the intact peptide (PTH 1-84) [32]. Refinement of the two-site assays has given rise to the second-generation PTH assays which detect PTH 1-84 exclusively [33]. Although the values obtained with the ‘‘intact’’ PTH assay correlate well with those obtained with the PTH 1-84 assays, the latter being 50–60% lower, a distinct advantage of the PTH 1-84 assays is that they may offer improved standardization between different laboratories [34]. The use of twosite PTH assays provides an accurate assessment of the current state of parathyroid activity and correlates well with the predominant type of osteodystrophy on bone histology, as illustrated in Figure 2. Thus, elevated PTH levels are the hallmark of osteitis fibrosa, whereas PTH values are much lower in the low bone turnover syndromes of adynamic bone or osteomalacia [35]. Hyperparathyroidism occurs early in the course of renal insufficiency, and PTH levels are often increased with histological evidence of hyperparathyroidism on bone. Precise guidelines for correlations of PTH levels with histological parameters of bone in early renal failure are relatively scarce, and most such correlations have been made in patients with end-stage renal disease using the firstgeneration two-site assays. The correlations of PTH values with bone histology in patients with end-stage renal disease

Diagnostic separation of the major types of renal bone disease by the measurement of intact PTH. Modified from Coburn and Salusky with permission [35].

also have shown that there is a resistance of the skeleton to PTH in renal failure, because supranormal levels appear to be required to maintain bone turnover in the normal range [36]. Thus, it has been found that ‘‘intact’’ PTH values of two to three times the upper limit of normal are necessary to achieve normal values for bone turnover. In view of these data, it is felt that values of ‘‘intact’’ PTH within 150 and 300 pg/ml (which correspond to 75 and 150 pg/mL in the PTH 1-84 assays) are generally associated with normal bone turnover in patients with end-stage renal disease. The implications of these findings are that these values for PTH should be the target values to be achieved with therapy. Thus, values below this range are often associated with abnormally low bone turnover, and conversely, levels above this range are generally associated with histologic evidence of hyperparathyroidism. The correlations, however, are far from perfect, and other diagnostic tools, as well as the use of serial measurements of PTH, may be helpful in improving the diagnostic utility. Since C-terminal PTH fragments such as PTH 7-84 may oppose the actions of PTH in bone [37, 38], it has been suggested that using the ratio between PTH 1-84 to PTH 7-84 may be of value in determining bone turnover in patients with chronic kidney disease; however, considerable controversy exists in this regard and the clinical utility of this ratio remains to be determined [39–41].

C. Measurement of Vitamin D Metabolites Abnormalities of vitamin D metabolism play a major role in the development of the secondary hyperparathyroidism characteristic of chronic kidney disease (as a result of impaired production of calcitriol from the diseased kidney as renal mass decreases [42]). In addition,

Chapter 45 Assessment of Bone and Joint Diseases: Renal Osteodystrophy

impaired glomerular filtration may compromise the delivery of substrate to the proximal tubular cell, thus contributing to decreased production of calcitriol [43]. Accordingly, levels of calcitriol in blood progressively decline as renal function declines [16]. Levels are maintained within the normal range in mild to moderate renal insufficiency, because high levels of PTH stimulate the production of calcitriol by the kidney. Thus, even normal levels of calcitriol could be considered inappropriately low for the prevailing levels of PTH. When GFR falls below 30% of normal, however, the reduction in renal mass limits the ability to produce sufficient amounts of calcitriol and, in spite of hyperparathyroidism, the levels of calcitriol in blood decline. Although the decrease in the levels of calcitriol is important as a pathogenetic factor for renal osteodystrophy, its measurement in chronic kidney disease offers little diagnostically unless extrarenal production of calcitriol is suspected (e.g. in sarcoidosis). Measurement of 25-hydroxyvitamin D provides the best index of vitamin D status and is useful if vitamin D deficiency is suspected (see also Chapter 33). Levels of this metabolite may decline in proteinuric renal diseases as a result of the loss of vitamin D binding protein in the urine. However, recent studies suggest that the presence of renal disease per se, is associated with a high incidence of vitamin D deficiency, and low levels of 25-hydroxyvitamin D correlate with high levels of PTH [44, 45]. These observations suggest that vitamin D deficiency is likely to play an important role in the pathogenesis of secondary hyperparathyroidism in the setting of renal disease, especially since in this patient population calcitriol levels are highly dependent on the availability of substrate [45]. Thus, for the management of secondary hyperparathyroidism in chronic kidney disease, maintenance of adequate 25-hydroxyvitamin D levels are recommended for patients who are not yet on dialysis [46].

D. Measurement of Serum Aluminum Although the definitive diagnosis of aluminum-related osteodystrophy requires a bone biopsy, the use of noninvasive tests may be helpful in this regard. The measurement of basal serum aluminum levels is not a useful test in identifying patients with aluminum bone disease [47]. Increments in serum aluminum following the administration of the metal chelator deferoxamine (DFO) reflect total aluminum accumulation in the body and may be useful to evaluate the tissue burden of aluminum, however, such assays are not specific for bone [48]. The accuracy of this test in the diagnosis of aluminum bone disease increases when it is interpreted in conjunction with PTH values [49]. Thus, an increment in plasma aluminum of 150 µg/L,

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following the administration of DFO (40 mg/kg) in the presence of PTH values of less than 100 pg/ml, is strongly suggestive of significant aluminum accumulation in bone. Lower doses of DFO have also been used and in one study, when the administration of 5 mg/kg was accompanied by an increment of serum aluminum greater than 50 µg/mL and an intact PTH level of less than 150 pg/mL, there was a high risk for aluminum-related bone disease [50]. In addition to PTH, the iron status should also be considered when interpreting the desferoxamine test [51]. Bone biopsy and the use of specific staining are required to show the presence of aluminum at the mineralization front.

E. Measurements of Alkaline Phosphatase in Serum Serum alkaline phosphatase is derived from intestine, liver, kidney, and bone. Nevertheless, measurements of total alkaline phosphatase in blood provide an approximate index of osteoblast activity in patients with renal failure because elevations in alkaline phosphatase are often due to an increase in the bone isoenzyme (an index of osteoblast function) in cases of hyperparathyroidism (see also Chapters 9 and 34). Conversely, alkaline phosphatase values tend to be reduced in cases of low bone turnover osteodystrophy [52]. Serial measurements of alkaline phosphate activity in a given patient may provide useful information regarding the progression of skeletal disease as well as an index of response to therapy for hyperparathyroidism. Most studies of the correlation of alkaline phosphatase with PTH and parameters of bone histomorphometry show wide variation. For these reasons as well as the prevalence of hepatic abnormalities in patients with renal failure, efforts have been made to develop specific measurements of the bone alkaline phosphatase isoenzyme with the hope of improving its diagnostic utility [53–55]. While such measurements eliminate the confusion with alkaline phosphatase from other tissues and show slightly better correlations with PTH and other parameters of bone histology than measurements of total alkaline phosphatase, they remain adjuncts to other parameters of skeletal metabolism. While elevated values are seen in states of high bone turnover, the inability to reliably distinguish normal from abnormally low bone turnover on an individual basis remains.

F. Measurements of Osteocalcin Osteocalcin is a vitamin K-dependent protein produced by the osteoblast and, accordingly, is a potential marker of osteoblast activity (see also Chapters 3 and 34). Levels of osteocalcin may be elevated in patients with renal failure

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because of decreased renal clearance of the protein from the circulation. In addition, the accumulation of osteocalcin fragments of unknown function in renal failure further compounds the difficulties interpreting osteocalcin levels. In general, correlations of osteocalcin values with parameters of bone histomorphometry do not appear to be superior to measurements of bone alkaline phosphatase [56]. The use of newer assay methods that detect only intact osteocalcin may offer a diagnostic advantage to separate patients with high bone turnover from those in whom bone turnover is abnormally low [57].

G. Other Markers of Bone Metabolism There has been increasing interest in measurements of several new markers of bone metabolism, among which are markers of collagen synthesis (such as procollagen type I C-terminal extension peptide [PICP]) and markers of collagen degradation (such as collagen type I C-terminal cross-linked telopeptide [ICTP], pyridinoline [PYD], and deoxypyridinoline [DPD] cross-links) [56, 58, 59]. PICP is a 100-kDa glycoprotein procollagen cleavage product generated after secretion of procollagen by osteoblasts; an additional N-terminal peptide, PINP is also liberated. In basal conditions, PICP in the circulation is mainly derived from bone and is metabolized by the liver; thus, renal impairment does not directly influence PICP levels in serum. ICTP is a 12-kDa peptide which is produced when collagen fibrils undergo degradation. PYD and DPD are also liberated as collagen is degraded. Due to their size, these substances are filtered by the kidney and excreted in the urine and would be expected to accumulate in serum when renal function is reduced. When these markers were studied in patients with renal osteodystrophy who were on dialysis, PICP did not correlate with histological evidence of bone formation. ICTP, PYP and DPD all correlated with the other humoral markers, such as alkaline phosphatase, osteocalcin, and PTH [56, 199]. At present, assays for these collagen synthesis and degradation products remain investigational, but they do not appear to offer any major advantage over the currently utilized markers of bone metabolism.

H. Cytokines Cytokines and growth factors are important regulators of bone turnover and many are regulated by the major calciotropic hormones, PTH and calcitriol (see also Chapter 7), and thus derangements in uremia, may affect the actions of these cytokines in bone [60]. Chronic kidney

disease is associated with abnormalities in cytokine production. There is evidence for increased levels of IL-1, IL-6, and TNF-α as well as other cytokines and growth factors, such as IGF-I, that may affect bone turnover [61–63]. In addition, the levels of cytokine antagonists and soluble receptors, such as IL-1 receptor antagonist and IL-6 soluble receptors, may also be elevated in uremia [64, 65]. Thus, the final effect of these factors in bone turnover represents an integration of the various components involved in the different systems. The role of cytokines in the pathogenesis of renal osteodystrophy is not clear at present. Ferreira et al. [64] found elevated levels of IL-1 and IL-1 receptor antagonist in patients on hemodialysis, and there was an inverse relationship between the plasma levels of IL-1 receptor antagonist and osteoblast surface. In the same study, elevated levels of IL-6 and IL-6 soluble receptors were found, and there was an inverse relationship between the levels of IL-6 receptor and osteoclast surface [64]. This study suggests that the relative imbalance between the different cytokine systems would influence the development of high versus low turnover bone disease. Studies by Langub et al. [66] have demonstrated the presence of IL-6 and IL-6 receptor mRNA in vivo using in situ hybridization in bone biopsies from patients with secondary hyperparathyroidism; the expression of IL-6 receptor paralleled bone-resorbing activity. Levels of IGF-I have been found to correlate with bone formation rate in patients on hemodialysis [63]. Osteoprotegerin is known to inhibit bone resorption, and when circulating levels are examined in conjunction with PTH, it may be helpful in differentiating high versus low turnover renal osteodystrophy [67]. Although considerable understanding of cytokines and growth factors in uremia has been gained, their relationship with the different histologic types of renal osteodystrophy has not been well characterized. Further study will be required to define the roles of these factors in the skeletal abnormalities and to define the utility of the assay for these mediators in serum for the diagnosis and management of renal bone disease.

I. β2 Microglobulin The syndrome of β2 microglobin amyloidosis, characterized by carpal tunnel syndrome, destructive arthropathies, and juxta-articular radiolucent cysts, is related to the accumulation of β2 microglobulin due to impaired elimination of this protein by the diseased kidney. The manifestations of this disorder result from the deposition of β2 microglobulin in the tissues in the form of amyloid fibrils [68]. However, measurements of the serum levels of β2 microglobulin, although universally elevated in

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patients with advanced renal failure, do not correlate with the presence of amyloid deposits, and therefore, are not clinically useful.

III. SKELETAL IMAGING IN RENAL OSTEODYSTROPHY A. X-Ray Subperiosteal erosions are the most consistent findings in secondary hyperparathyroidism. Radiographs, however, are quite insensitive and can appear virtually normal in patients who have histologic evidence of severe hyperparathyroidism on bone biopsy. The radiological features of secondary hyperparathyroidism are best detected on hand X-rays. The use of fine-grain film and magnification techniques facilitates the detection of abnormalities [69]. Subperiosteal bone erosions are initially evident in the hands and involve the phalangeal tufts and the radial aspects of the middle phalanges of the first and second digits. They may also be seen at the distal end of the clavicles, the tibia, the femoral neck, the humerus, and the region of the sacroiliac joints. While subperiosteal erosions usually represent hyperparathyroidism, it is important to emphasize that the findings may not necessarily be current and may represent prior hyperparathyroidism that has now been replaced by osteomalacia, such as might occur with aluminum intoxication. Skull X-rays may show a ground-glass appearance, focal radiolucencies, and areas of sclerosis. Patchy areas of osteosclerosis in the vertebrae lead to the characteristic appearance of the “rugger jersey spine”. In children, there may be evidence of growth retardation and rickets. Cystic lesions and spondyloarthropathy may represent dialysis-related β2 microglobulin amyloidosis. The lesions are usually multiple, juxta-articular in location and may cause pathological fractures and collapse of the articular surface [70, 71]. The presence of aluminum-related bone is characterized by reduced bone density and fractures. Osteomalacia related to aluminum intoxication is suggested by the finding of Looser’s zones or pseudofractures.

B. Bone Scintiscans Bone scanning using technetium-99m methylene diphosphonate (Tc 99m MDP) has been used to evaluate renal osteodystrophy and may also be used to assess the response to therapy [72–74]. MDP adsorbs to bone surfaces and has a higher affinity to sites of new bone formation. Uptake of tracer tends to be diffusely increased in states of high bone

turnover, such as in patients with hyperparathyroidism. Common features of increased bone turnover include a ‘‘tie’’ sternum, beading of the costochondral junctions, a prominent calvarium and mandible, and increases uptake in the long bones and periarticular regions [75]. Patients with low bone turnover and osteomalacia showed decreased uptake, with possible focal areas of increased activity representing microfractures. Pentavalent 99m Tc-dimercaptosuccinic acid (DMSA) scintigraphy has also been studied in patients with renal osteodystrophy and it appears to be useful in assessing response to therapy [76, 77]. Ectopic calcification may also become apparent by bone scan. The sensitivity for bone scan studies to differentiate between the various kinds of renal osteodystrophy is not high and mixed renal osteodystrophy cannot be identified.

C. PET Scan Efforts to characterize bone metabolic activity by positron emission tomography with 18F have been described and appear capable of differentiating low bone turnover from high bone turnover lesions of renal osteodystrophy [78]. Excellent correlations of fluoride uptake with values of PTH and alkaline phosphatase were obtained. Although this technique has potential value for serial studies of bone metabolism, it is not widely utilized.

D. Measurement of Bone Density While measurement of bone density has been shown to be a reliable indicator of fracture risk in patients without renal disease, the use of such techniques in patients with renal disease is less well defined. Single- and dual-photon absorptiometry have largely been replaced by dual-energy X-ray absorptiometry (DEXA) [79]. A variety of studies have reported low mineral density obtained by DEXA in patients requiring dialysis [80, 81]. Bone mineral density has been found to correlate inversely with alkaline phosphatase, osteocalcin, and PTH levels [82, 83]. DEXA has also been evaluated as a tool to discriminate between the different types of renal osteodystrophy and, in those studies, bone mineral density was lower in the presence of hyperparathyroid bone disease as compared to adynamic bone [83]. One major limitation of DEXA in the diagnosis of renal osteodystrophy is that it only measures bone mineral density and offers no information regarding bone structure. Quantitative CT is less commonly used although this technique can separate cortical from trabecular bone [84]. This method uses considerably more radiation

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than DEXA. MRI offers high-resolution images and is radiation free. Preliminary MRI data suggest that this non-invasive modality may have the potential to directly visualize and quantify trabecular and cortical bone structural parameters, a characterization that is not possible with DEXA. Recently, studies on the potential utility of magnetic resonance imaging in patients with renal osteodystrophy have started to emerge [85]. The use of Micro MRI to obtain a set of high-resolution contiguous slices allows the reconstruction of three-dimensional representation of trabecular bone architecture in a volume of interest which can be analyzed as virtual bone biopsy [86]. The clinical utility of this modality needs to be verified in studies comparing bone biopsies with virtual bone biopsies. Early studies of bone density, in general, showed that renal osteodystrophy was associated with a decrease in cortical bone. Axial bone mass measurements were more heterogeneous [53, 79, 87]. Some studies demonstrated a gradual loss or no bone loss over time, whereas other studies actually demonstrated a gain of bone. Other investigators found some patients who had a relatively rapid loss of bone. From the heterogeneity of bone histology discussed above, it is perhaps not surprising that a composite measurement of bone density would not have a specific diagnostic value in this patient population. In addition, there are many technical difficulties in this group of patients because many may have vascular and ligament calcifications which may affect the results. Data on fracture risk in this patient population are limited and prospective data would be useful [88–90]. Thus, the clinical utility of measurements of bone mineral density in renal osteodystrophy is limited at the present time.

IV. SUMMARY Renal osteodystrophy represents a complex disorder of bone metabolism resulting from the many derangements that occur during the course of chronic kidney disease. The patterns of bone disease revealed by histological examination of bone lead to useful classifications of the disorder and provide specific diagnostic information. The many non-invasive tools that may be utilized for the assessment of bone metabolism are clinically useful when considered together, especially when serial measurements are available. An understanding of the pathogenesis of renal osteodystrophy and early therapy directed at these pathogenetic factors are important to minimize the significant morbidity that occurs as a result of this complication of chronic kidney disease.

References 1. Martin, K. J., Gonzalez, E. A., and Slatopolsky, E. (2004). Renal Osteodystrophy. In: The Kidney (B. M. Brenner, ed.), pp. 2255–2304. W.B. Saunders Company, Philadelphia, PA. 2. Malluche, H., and Faugere, M. C. (1990). Renal bone disease 1990: an unmet challenge for the nephrologist. Kidney Int. 38, 193–211. 3. Sherrard, D. J., Hercz, G., Pei, Y., Maloney, N. A., Greenwood, C., Manuel, A., Saiphoo, C., Fenton, S. S., and Segre, G. V. (1993). The spectrum of bone disease in end-stage renal failure – an evolving disorder. Kidney Int. 43, 436–442. 4. Malluche, H., Ritz, E., and Lange, H. (1976). Bone histology in incipient and advanced renal failure. Kidney Int. 9, 355–362. 5. Coen, G., Ballanti, P., Bonucci, E., Calabria, S., Costantini, S., Ferrannini, M., Giustini, M., Giordano, R., Nicolai, G., Manni, M., Sardella, D., and Taggi, F. (2002). Renal osteodystrophy in predialysis and hemodialysis patients: Comparison of histologic patterns and diagnostic predictivity of intact PTH. Nephron 91, 103–111. 6. Slatopolsky, E., Brown, A., and Dusso, A. (1999). Pathogenesis of secondary hyperparathyroidism. Kidney Int. Suppl. 73, S14–19. 7. González, E. A., and Martin, K. J. (1995). Renal osteodystrophy: pathogenesis and management. Nephrol. Dial. Transplant. 10 (Suppl 3), 13–21. 8. González, E., and Martin, K. (1992). Aluminum and renal osteodystrophy: a diminishing clinical problem. Trends Endocrinol. Metab. 3, 371–375. 9. Goodman, W., Henry, D., Horst, R., Nudelman, R., Alfrey, A., and Coburn, J. (1984). Parenteral aluminum administration in the dog. II. Induction of osteomalacia and effect on vitamin D metabolism. Kidney Int. 25, 370–375. 10. Cournot-Witmer, G., Zingraff, J., Plachot, J. J., Escaig, F., Lefevre, R., Boumati, P., Bourdeau, A., Garabedian, M., Galle, P., Bourdon, R., Drueke, T., and Balsan, S. (1981). Aluminum localization in bone from hemodialyzed patients: relationship to matrix mineralization. Kidney Int. 20, 375–378. 11. Hutchison, A. J., Whitehouse, R. W., Freemont, A. J., Adams, J. E., Mawer, E. B., and Gokal, R. (1994). Histological, radiological, and biochemical features of the adynamic bone lesion in continuous ambulatory peritoneal dialysis patients. Am. J. Nephrol. 14, 19–29. 12. Sherrard, D. J., Hercz, G., Pei, Y., and Segre, G. (1996). The aplastic form of renal osteodystrophy. Nephrol. Dial. Transplant. 11 (Suppl 3), 29–31. 13. Couttenye, M. M., D’Haese, P. C., Verschoren, W. J., Behets, G. J., Schrooten, I., and De Broe, M. E. (1999). Low bone turnover in patients with renal failure. Kidney Int. 56, S70–S76. 14. Malluche, H., and Monier-Faugere, M. (1994). The role of bone biopsy in the management of patients with renal osteodystrophy. J. Am. Soc. Nephrol. 4, 1631–1642. 15. Kates, D., Sherrard, D., and Andress, D. (1997). Evidence that serum phosphate is independently associated with serum PTH in patients with chronic renal failure. Am. J. Kidney Dis. 30, 809–813. 16. Martinez, I., Saracho, R., Montenegro, J., and Llach, F. (1997). The importance of dietary calcium and phosphorous in the secondary hyperparathyroidism of patients with early renal failure. Am. J. Kidney Dis. 29, 496–502. 17. Block, G. A., Martin, K. J., de Francisco, A. L., Turner, S. A., Avram, M. M., Suranyi, M. G., Hercz, G., Cunningham, J., Abu-Alfa, A. K., Messa, P., Coyne, D. W., Locatelli, F., Cohen, R. M., Evenepoel, P., Moe, S. M., Fournier, A., Braun, J., McCary, L. C., Zani, V. J., Olson, K. A., Drueke, T. B., and Goodman, W. G. (2004). Cinacalcet for secondary hyperparathyroidism in patients receiving hemodialysis. N. Engl. J. Med. 350, 1516–1525. 18. Hercz, G., Kraut, J. A., Andress, D. A., Howard, N., Roberts, C., Shinaberger, J. H., Sherrard, D. J., and Coburn, J. W. (1986). Use of calcium carbonate as a phosphate binder in dialysis patients. Min. Electrolyte Metab. 12, 314–319.

Chapter 45 Assessment of Bone and Joint Diseases: Renal Osteodystrophy 19. Slatopolsky, E., Weerts, C., Lopez-Hilker, S., Norwood, K., Zink, M., Windus, D., and Delmez, J. (1986). Calcium carbonate as a phosphate binder in patients with chronic renal failure undergoing dialysis. N. Engl. J. Med. 315, 157–161. 20. Delmez, J. A., Dusso, A. S., Slatopolsky, E., and Teitelbaum, S. L. (1995). Modulation of renal osteodystrophy by extrarenal production of calcitriol. Am. J. Nephrol. 15, 85–89. 21. Barbour, G. L., Coburn, J. W., Slatopolsky, E., Norman, A. W., and Horst, R. L. (1981). Hypercalcemia in an anephric patient with sarcoidosis: evidence for extrarenal generation of 1,25-dihydroxyvitamin D. N. Eng. J. Med. 305, 440–443. 22. Slatopolsky, E., Caglar, S., Gradowska, L., Canterbury, J., Reiss, E., and Bricker, N. S. (1972). On the prevention of secondary hyperparathyroidism in experimental chronic renal disease using “proportional reduction” of dietary phosphorus intake. Kidney Int. 2, 147–151. 23. Slatopolsky, E., and Bricker, N. S. (1973). The role of phosphorus restriction in the prevention of secondary hyperparathyroidism in chronic renal disease. Kidney Int. 4, 141–145. 24. Slatopolsky, E., Finch, J., Denda, M., Ritter, C., Zhong, M., Dusso, A., MacDonald, P. N., and Brown, A. J. (1996). Phosphorus restriction prevents parathyroid gland growth. High phosphorus directly stimulates PTH secretion in vitro. J. Clin. Invest. 97, 2534–2540. 25. Almaden, Y., Canalejo, A., Hernandez, A., Ballesteros, E., Garcia-Navarro, S., Torres, A., and Rodriguez, M. (1996). Direct effect of phosphorus on PTH secretion from whole rat parathyroid glands in vitro. J. Bone Min. Res. 11, 970–976. 26. Tanaka, Y., and Deluca, H. F. (1973). The control of 25-hydroxyvitamin D metabolism by inorganic phosphorus. Arch. Biochem. Biophys. 154, 566–574. 27. Block, G. A., Hulbert-Shearon, T. E., Levin, N. W., and Port, F. K. (1998). Association of serum phosphorus and calcium x phosphate product with mortality risk in chronic hemodialysis patients: a national study. Am. J. Kidney Dis. 31, 607–617. 28. Block, G. A., Klassen, P. S., Lazarus, J. M., Ofsthun, N., Lowrie, E. G., and Chertow, G. M. (2004). Mineral metabolism, mortality, and morbidity in maintenance hemodialysis. J. Am. Soc. Nephrol. 15, 2208–2218. 29. Martin, K. J., Hruska, K. A., Lewis, J., Anderson, C., and Slatopolsky, E. (1977). The renal handling of parathyroid hormone. Role of peritubular uptake and glomerular filtration. J. Clin. Invest. 60, 808–814. 30. Brown, R. C., Aston, J. P., Weeks, I., and Woodhead, J. S. (1987). Circulating intact parathyroid hormone measured by a two-site immunochemiluminometric assay. J. Clin. Endocrinol. Metab. 65, 407–414. 31. Nussbaum, S. R., Zahradnik, R. J., Lavigne, J. R., Brennan, G. L., Nozawa-Ung, K., Kim, L. Y., Keutmann, H. T., Wang, C. A., Potts, J. T., Jr., and Segre, G. V. (1987). Highly sensitive two-site immunoradiometric assay of parathyrin, and its clinical utility in evaluating patients with hypercalcemia. Clin. Chem. 33, 1364–1367. 32. Lepage, R., Roy, L., Brossard, J. H., Rousseau, L., Dorais, C., Lazure, C., and D’Amour, P. (1998). A non-(1–84) circulating parathyroid hormone (PTH) fragment interferes significantly with intact PTH commercial assay measurements in uremic samples. Clin. Chem. 44, 805–809. 33. Gao, P., Scheibel, S., D’Amour, P., John, M. R., Rao, S. D., Schmidt-Gayk, H., and Cantor, T. L. (2001). Development of a novel immunoradiometric assay exclusively for biologically active whole parathyroid hormone 1–84: implications for improvement of accurate assessment of parathyroid function. J. Bone Miner. Res. 16, 605–614. 34. Martin, K. J., Akhtar, I., and Gonzalez, E. A. (2004). Parathyroid hormone: New assays, new receptors. Semin. Nephrol. 24, 3–9. 35. Coburn, J., and Salusky, I. B. (1994). Hyperparathyroidism in renal failure: clinical features, diagnosis and management. In: The Parathyroids: Basic and Clinical Concepts (J. P. Bilezikian, M. Levine, and R. Marcus, eds.), pp. 721–745. Raven Press, New York. 36. Qi, Q., Monier-Faugere, M. C., Geng, Z., and Malluche, H. H. (1995). Predictive value of serum parathyroid hormone levels for bone

37.

38.

39.

40.

41.

42.

43.

44.

45.

46. 47.

48.

49.

50.

51.

52.

763

turnover in patients on chronic maintenance dialysis. Am. J. Kidney Dis. 26, 622–631. Slatopolsky, E., Finch, J., Clay, P., Martin, D., Sicard, G., Singer, G., Gao, P., Cantor, T., and Dusso, A. (2000). A novel mechanism for skeletal resistance in uremia. Kidney Int. 58, 753–761. Nguyen-Yamamoto, L., Rousseau, L., Brossard, J. H., Lepage, R., and D’Amour, P. (2001). Synthetic carboxyl-terminal fragments of parathyroid hormone (pth) decrease ionized calcium concentration in rats by acting on a receptor different from the pth/pth-related peptide receptor. Endocrinology 142, 1386–1392. Monier-Faugere, M. C., Geng, Z., Mawad, H., Friedler, R. M., Gao, P., Cantor, T. L., and Malluche, H. H. (2001). Improved assessment of bone turnover by the PTH-(1–84)/large C-PTH fragments ratio in ESRD patients. Kidney Int. 60, 1460–1468. Coen, G., Bonucci, E., Ballanti, P., Balducci, A., Calabria, S., Nicolai, G. A., Fischer, M. S., Lifrieri, F., Manni, M., Morosetti, M., Moscaritolo, E., and Sardella, D. (2002). PTH 1-84 and PTH “7-84” in the noninvasive diagnosis of renal bone disease. Am. J. Kidney Dis. 40, 348–354. Salusky, I. B., Goodman, W. G., Kuizon, B. D., Lavigne, J. R., Zahranik, R. J., Gales, B., Wang, H. J., Elashoff, R. M., and Juppner, H. (2003). Similar predictive value of bone turnover using first- and second-generation immunometric PTH assays in pediatric patients treated with peritoneal dialysis. Kidney Int. 63, 1801–1808. Slatopolsky, E., Lopez-Hilker, S., Delmez, J., Dusso, A., Brown, A., and Martin, K. J. (1990). The parathyroid-calcitriol axis in health and chronic renal failure. Kidney Int. Suppl. 29, S41–47. Nykjaer, A., Dragun, D., Walther, D., Vorum, H., Jacobsen, C., Herz, J., Melsen, F., Christensen, E. I., and Willnow, T. E. (1999). An endocytic pathway essential for renal uptake and activation of the steroid 25-(OH) vitamin D3. Cell 96, 507–515. Thomas, M. K., Lloyd-Jones, D. M., Thadhani, R. I., Shaw, A. C., Deraska, D. J., Kitch, B. T., Vamvakas, E. C., Dick, I. M., Prince, R. L., and Finkelstein, J. S. (1998). Hypovitaminosis D in medical inpatients. N. Engl. J. Med. 338, 777–783. Gonzalez, E. A., Sachdeva, A., Oliver, D. A., and Martin, K. J. (2004). Vitamin D insufficiency and deficiency in chronic kidney disease. A single center observational study. Am. J. Nephrol. 24, 503–510. Eknoyan, G., Levin, A., and Levin, N. W. (2003). Bone metabolism and disease in chronic kidney disease. Am. J. Kidney Dis. 42, 1–201. Malluche, H. H., Smith, A. J., Abreo, K., and Faugere, M. C. (1984). The use of deferoxamine in the management of aluminium accumulation in bone in patients with renal failure. N. Eng. J. Med. 311, 140–144. Milliner, D. S., Nebeker, H. G., Ott, S. M., Andress, D. L., Sherrard, D. J., Alfrey, A. C., Slatopolsky, E. A., and Coburn, J. W. (1984). Use of the deferoxamine infusion test in the diagnosis of aluminum-related osteodystrophy. Ann. Intern. Med. 101, 775–779. Pei, Y., Hercz, G., Greenwood, C., Sherrard, D., Segre, G., Manuel, A., Saiphoo, C., and Fenton, S. (1992). Non-invasive prediction of aluminum bone disease in hemo- and peritoneal dialysis patients. Kidney Int. 41, 1374–1382. D’Haese, P. C., Couttenye, M. M., Goodman, W. G., Lemoniatou, E., Digenis, P., Sotornik, I., Fagalde, A., Barsoum, R. S., Lamberts, L. V., and De Broe, M. E. (1995). Use of the low-dose desferrioxamine test to diagnose and differentiate between patients with aluminium-related bone disease, increased risk for aluminium toxicity, or aluminium overload. Nephrol. Dial. Transplant. 10, 1874–1884. D’Haese, P. C., Couttenye, M. M., and De Broe, M. E. (1996). Diagnosis and treatment of aluminium bone disease. Nephrol. Dial. Transplant. 11, 74–79. Couttenye, M. M., D’Haese, P. C., Van Hoof, V. O., Lemoniatou, E., Goodman, W., Verpooten, G. A., and De Broe, M. E. (1996). Low serum levels of alkaline phosphatase of bone origin: a good marker of adynamic bone disease in haemodialysis patients. Nephrol. Dial. Transplant. 11, 1065–1072.

764 53. Fletcher, S., Jones, R. G., Rayner, H. C., Harnden, P., Hordon, L. D., Aaron, J. E., Oldroyd, B., Brownjohn, A. M., Turney, J. H., and Smith, M. A. (1997). Assessment of renal osteodystrophy in dialysis patients: use of bone alkaline phosphatase, bone mineral density and parathyroid ultrasound in comparison with bone histology. Nephron 75, 412–419. 54. Jarava, C., Armas, J. R., Salgueira, M., and Palma, A. (1996). Bone alkaline phosphatase isoenzyme in renal osteodystrophy. Nephrol. Dial. Transplant. 11, 43–46. 55. Urena, P., Hruby, M., Ferreira, A., Ang, K. S., and de Vernejoul, M. C. (1996). Plasma total versus bone alkaline phosphatase as markers of bone turnover in hemodialysis patients. J. Am. Soc. Nephrol. 7, 506–512. 56. Mazzaferro, S., Pasquali, M., Ballanti, P., Bonucci, E., Costantini, S., Chicca, S., De Meo, S., Perruzza, I., Sardella, D., Taggi, F., and Coen, G. (1995). Diagnostic value of serum peptides of collagen synthesis and degradation in dialysis renal osteodystrophy. Nephrol. Dial. Transplant. 10, 52–58. 57. Morishita, T., Nomura, M., Hanaoka, M., Saruta, T., Matsuo, T., and Tsukamoto, Y. (2000). A new assay method that detects only intact osteocalcin. Two-step non-invasive diagnosis to predict adynamic bone disease in haemodialysed patients. Nephrol. Dial. Transplant. 15, 659–667. 58. De la Piedra, C., Diaz Martin, M. A., Diaz Diego, E. M., Lopez Gavilanes, E., Gonzalez Parra, E., Caramelo, C., and Rapado, A. (1994). Serum concentrations of carboxyterminal cross-linked telopeptide of type I collagen (ICTP), serum tartrate resistant acid phosphatase, and serum levels of intact parathyroid hormone in parathyroid hyperfunction. Scand. J. Clin. Lab. Invest. 54, 11–15. 59. Ibrahim, S., Mojiminiyi, S., and Barron, J. L. (1995). Pyridinium crosslinks in patients on haemodialysis and continuous ambulatory peritoneal dialysis. Nephrol. Dial. Transplant. 10, 2290–2294. 60. Gonzalez, E. A. (2000). The role of cytokines in skeletal remodelling: possible consequences for renal osteodystrophy. Nephrol. Dial. Transplant. 15, 945–950. 61. Herbelin, A., Ureña, P., Nguyen, A. T., Zingraff, J., and DescampsLatscha, B. (1991). Elevated circulating levels of interleukin-6 in patients with chronic renal failure. Kidney Int. 39, 954–960. 62. Herbelin, A., Nguyen, A. T., Zingraff, J., Urena, P., and DescampsLatscha, B. (1990). Influence of uremia and hemodialysis on circulating interleukin-1 and tumor necrosis factor alpha. Kidney Int. 37, 116–125. 63. Andress, D. L., Pandian, M. R., Endres, D. B., and Kopp, J. B. (1989). Plasma insulin-like growth factors and bone formation in uremic hyperparathyroidism. Kidney Int. 36, 471–477. 64. Ferreira, A., Simon, P., Drueke, T. B., and Deschamps-Latscha, B. (1996). Potential role of cytokines in renal osteodystrophy [letter]. Nephrol. Dial. Transplant. 11, 399–400. 65. Moutabarrik, A., Nakanishi, I., Namiki, M., and Tsubakihara, Y. (1995). Interleukin-1 and its naturally occurring antagonist in peritoneal dialysis patients. Clin. Nephrol. 43, 243–248. 66. Langub, M. C., Jr., Koszewski, N. J., Turner, H. V., MonierFaugere, M. C., Geng, Z., and Malluche, H. H. (1996). Bone resorption and mRNA expression of IL-6 and IL-6 receptor in patients with renal osteodystrophy. Kidney Int. 50, 515–520. 67. Haas, M., Leko-Mohr, Z., Roschger, P., Kletzmayr, J., Schwarz, C., Domenig, C., Zsontsich, T., Klaushofer, K., Delling, G., and Oberbauer, R. (2002). Osteoprotegerin and parathyroid hormone as markers of high-turnover osteodystrophy and decreased bone mineralization in hemodialysis patients. Am. J. Kidney Dis. 39, 580–586. 68. Gejyo, F., Yamada, T., Odani, S., Nakagawa, Y., Arakawa, M., Kunitomo, T., Kataoka, H., Suzuki, M., Hirasawa, Y., Shirahama, T., Cohen, A. S., and Schmid, K. (1985). A new form of amyloid protein associated with chronic hemodialysis was identified as beta 2-microglobulin. Biochem. Biophys. Res. Commun. 129, 701–706.

Esther A. González ET AL. 69. Meema, H. E., Rabinovich, S., Meema, S., Lloyd, G. J., and Oreopoulos, D. G. (1972). Improved radiological diagnosis of azotemic osteodystrophy. Radiology 102, 1–10. 70. Murphey, M. D., Sartoris, D. J., Quale, J. L., Pathria, M. N., and Martin, N. L. (1993). Musculoskeletal manifestations of chronic renal insufficiency. Radiographics 13, 357–379. 71. Moriniere, P., Marie, A., el Esper, N., Fardellone, P., Deramond, H., Remond, A., Sebert, J. L., and Fournier, A. (1991). Destructive spondyloarthropathy with beta 2-microglobulin amyloid deposits in a uremic patient before chronic hemodialysis. Nephron. 59, 654–657. 72. Karsenty, G., Vigneron, N., Jorgetti, V., Fauchet, M., Zingraff, J., Drueke, T., and Cournot-Witmer, G. (1986). Value of the 99mTcmethylene diphosphonate bone scan in renal osteodystrophy. Kidney Int. 29, 1058–1065. 73. Vanherweghem, J. L., Schoutens, A., Bergman, P., Dhaene, M., Goldman, M., Fuss, M., and Kinnaert, P. (1985). Usefulness of 99mTc pyrophosphate bone scintigraphy in the survey of dialysis osteodystrophy. Proceedings of the European Dialysis & Transplant Association. European Renal Association 21, 431–434. 74. Olgaard, K., Heerfordt, J., and Madsen, S. (1976). Scintigraphic skeletal changes in uremic patients on regular hemodialysis. Nephron 17, 325–334. 75. Hain, S. F., and Fogelman, I. (2002). Nuclear medicine studies in metabolic bone disease. Semin. Musculoskelet. Radiol. 6, 323–329. 76. Higuchi, T., Hirano, T., Inoue, T., Aoki, J., Ueki, K., Wakamatsu, R., Yano, S., Naruse, T., and Endo, K. (1998). Pentavalent technetium99m-dimercaptosuccinic acid scintigraphy in renal osteodystrophy. J. Nucl. Med. 39, 541–543. 77. Sarikaya, A., Sen, S., Hacimahmutoglu, S., and Pekindil, G. (2002). 99mTc(V)-DMSA scintigraphy in monitoring the response of bone disease to vitamin D3 therapy in renal osteodystrophy. Ann. Nucl. Med. 16, 19–23. 78. Messa, C., Goodman, W. G., Hoh, C. K., Choi, Y., Nissenson, A. R., Salusky, I. B., Phelps, M. E., and Hawkins, R. A. (1993). Bone metabolic activity measured with positron emission tomography and [18F]fluoride ion in renal osteodystrophy: correlation with bone histomorphometry. J. Clin. Endocrinol. & Metab. 77, 949–955. 79. Erlichman, M., and Holohan, T. V. (1996). Bone densitometry: patients with end-stage renal disease. Health Tech. Assess, 1–27. 80. Stein, M. S., Packham, D. K., Ebeling, P. R., Wark, J. D., and Becker, G. J. (1996). Prevalence and risk factors for osteopenia in dialysis patients. Am. J. Kidney Diseases 28, 515–522. 81. Gabay, C., Ruedin, P., Slosman, D., Bonjour, J. P., Leski, M., and Rizzoli, R. (1993). Bone mineral density in patients with end-stage renal failure. Am. J. Nephrol. 13, 115–123. 82. Ha, S. K., Park, C. H., Seo, J. K., Park, S. H., Kang, S. W., Choi, K. H., Lee, H. Y., and Han, D. S. (1996). Studies on bone markers and bone mineral density in patients with chronic renal failure. Yonsei Med. J. 37, 350–356. 83. Gerakis, A., Hadjidakis, D., Kokkinakis, E., Apostolou, T., Raptis, S., and Billis, A. (2000). Correlation of bone mineral density with the histological findings of renal osteodystrophy in patients on hemodialysis. J. Nephrol. 13, 437–443. 84. Hutchison, A. J., Whitehouse, R. W., Boulton, H. F., Adams, J. E., Mawer, E. B., Freemont, T. J., and Gokal, R. (1993). Correlation of bone histology with parathyroid hormone, vitamin D3, and radiology in end-stage renal disease. Kidney Int. 44, 1071–1077. 85. Wehrli, F. W., Leonard, M. B., Saha, P. K., and Gomberg, B. R. (2004). Quantitative high-resolution magnetic resonance imaging reveals structural implications of renal osteodystrophy on trabecular and cortical bone. J. Magn. Reson. Imag. 20, 83–89. 86. Wehrli, F. W., Saha, P. K., Gomberg, B. R., Song, H. K., Snyder, P. J., Benito, M., Wright, A., and Weening, R. (2002). Role of magnetic resonance for assessing structure and function of trabecular bone. Top. Magn. Reson. Imag. 13, 335–355.

Chapter 45 Assessment of Bone and Joint Diseases: Renal Osteodystrophy 87. Asaka, M., Iida, H., Entani, C., Fujita, M., Izumino, K., Takata, M., Seto, H., and Sasayama, S. (1992). Total and regional bone mineral density by dual photon absorptiometry in patients on maintenance hemodialysis. Clin. Nephrol. 38, 149–153. 88. Fontaine, M. A., Albert, A., Dubois, B., Saint-Remy, A., and Rorive, G. (2000). Fracture and bone mineral density in hemodialysis patients. Clin. Nephrol. 54, 218–226.

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89. Atsumi, K., Kushida, K., Yamazaki, K., Shimizu, S., Ohmura, A., and Inoue, T. (1999). Risk factors for vertebral fractures in renal osteodystrophy. Am. J. Kidney Dis. 33, 287–293. 90. Jamal, S. A., Chase, C., Goh, Y. I., Richardson, R., and Hawker, G. A. (2002). Bone density and heel ultrasound testing do not identify patients with dialysis-dependent renal failure who have had fractures. Am. J. Kidney Dis. 39, 843–849.

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Chapter 46

Primary Hyperparathyroidism

I. II. III. IV.

Shonni J. Silverberg

Department of Medicine, College of Physicians and Surgeons, Columbia University, New York, USA

John P. Bilezikian

Departments of Medicine and Pharmacology, College of Physicians and Surgeons, Columbia University, New York, USA

V. Cytokines in Primary Hyperparathyroidism VI. Treatment of Primary Hyperparathyroidism VII. Summary References

Introduction Etiology Clinical Presentation Bone Markers in Primary Hyperparathyroidism

I. INTRODUCTION

have no signs or symptoms attributable to their primary hyperparathyroidism. Nephrolithiasis is still seen, although less frequently than in the past. However, radiologically evident bone disease is rare. This chapter presents the clinical picture of primary hyperparathyroidism as it presents today and the current status of bone markers in the investigation and management of this disorder.

Primary hyperparathyroidism, one of the most common endocrine disorders, is characterized by hypercalcemia and elevated parathyroid hormone levels. The disease today bears little resemblance to the symptomatic disorder of “stones, bones, and groans” described by Fuller Albright and others in the 1930s [1–6]. Nephrocalcinosis was present in most patients, and neuromuscular dysfunction with muscle weakness was common. Osteitis fibrosa cystica was the skeletal hallmark of classic primary hyperparathyroidism. This condition was characterized by brown tumors of the long bones, subperiosteal bone resorption, distal tapering of the clavicles and phalanges, and “salt and pepper” appearance of the skull on radiograph [7]. With the advent of the automated serum chemistry autoanalyzer in the 1970s, diagnosis of primary hyperparathyroidism in the absence of typical features became common [8–10]. In the United States today, more than three-quarters of patients Dynamics of Bone and Cartilage Metabolism

II. ETIOLOGY The most common lesion found in patients with primary hyperparathyroidism is the solitary parathyroid adenoma, which accounts for 80% of cases [11]. Identifiable risk factors in the development of parathyroid adenoma include a history of neck irradiation [12] and prolonged use of lithium therapy for affective disorders [13–16]. Research has helped to elucidate the molecular pathogenesis of the parathyroid adenoma. Most parathyroid adenomas are 767

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Shonni J. Silverberg and John P. Bilezikian

monoclonal in origin with a variety of molecular defects proposed to account for abnormal parathyroid cell function in this disease [17–25]. Even with advances in understanding of the molecular pathogenesis of primary hyperparathyroidism, the vast majority of cases are idiopathic. In approximately 15% of cases, all four parathyroid glands are involved in a process characterized by hyperplasia of all four glands. There are no clinical features that differentiate primary hyperparathyroidism due to an adenoma vs. hyperplasia. The etiology of parathyroid hyperplasia is multifactorial. In nearly one-half of cases, it is associated with a familial hereditary syndrome, such as multiple endocrine neoplasia types 1 and 2a.

III. CLINICAL PRESENTATION When the diagnosis of primary hyperparathyroidism is made, approximately 80% of patients have neither symptoms nor signs referable to their disease [7]. A history of previous neck irradiation is obtained in 10–15% of cases. Diseases epidemiologically associated with primary hyperparathyroidism include hypertension [8, 26–28], peptic ulcer disease, pancreatitis, gout, or pseudogout [29, 30], but no causal relationship has been established between primary hyperparathyroidism and these other disorders. Non-specific complaints include weakness, easy fatigability, depression, and a sense of intellectual weariness. Neuropsychiatric features have been described, but quantification of nonspecific symptomatology has been difficult [31–34]. Physical examination is generally unremarkable.

A. Biochemical Profile The diagnosis of primary hyperparathyroidism is made by demonstrating hypercalcemia in the presence of elevated parathyroid hormone levels. Improved means of measuring intact parathyroid hormone, by immunoradiometric and immunochemiluminometric assays, offer far greater utility than previously available assays (see Chapter 32). Even with the immunoradiometric assays, however, parathyroid hormone is frankly elevated in only about 80% of patients at the time of diagnosis [7]. In the remaining 15–20%, the parathyroid hormone concentration is in the upper range of normal, a situation that generally does not present a diagnostic problem because it is inappropriately high given the patient’s hypercalcemia. It should be noted also that in younger individuals with primary hyperparathyroidism (less than 45 years old), the normal range is actually lower than the given commercial laboratory normal range of 10–65 pg/ml. In those younger individuals,

the normal range is more accurately defined as approximately 10–45 pg/ml. Thus, a parathyroid hormone level of 50 pg/ml in a 40-year-old woman with hypercalcemia should be regarded as elevated. Any patient with hypercalcemia should have suppressed levels of parathyroid hormone unless they have primary hyperparathyroidism. The only exceptions to this rule are those with familial hypocalciuric hypercalcemia (FHH) and those who are taking thiazide diuretics or lithium. Finally, it has become clear that the IRMA assays in common use today measure not only PTH(1-84) but also some large carboxyterminal fragments of PTH, such as PTH(7-84). A newer IRMA assay that detects only the full-length molecule, PTH(1-84), has been documented to be more sensitive in the diagnosis of primary hyperparathyroidism than previously available assays [35], but whether it will be shown to be more clinically useful remains to be seen (see also Chapter 32). Other biochemical features of primary hyperparathyroidism include a serum phosphorus concentration in the low-normal range. Frankly low levels in primary hyperparathyroidism are present in less than one-quarter of patients. While mean values of 1,25-dihydroxyvitamin D3 are usually in the high-normal range, approximately onethird of patients have frankly elevated levels [36]. Serum 25-hydroxyvitamin D levels tend to be in the lower end of the normal range. Although frank vitamin D deficiency is not common in patients with primary hyperparathyroidism in the United States, levels of 25-hydroxyvitamin D in the “insufficiency” range (<20 ng/ml) have been reported in over half of all patients with mild disease [37]. More profound vitamin D deficiency co-exists with primary hyperparathyroidism in those areas of the world where vitamin D deficiency is seen regularly. In those regions, the simultaneous presence of vitamin D deficiency may account for the more severe skeletal phenotype seen in those countries. Indeed, osteitis fibrosa cystica is commonly seen in such patients [38].

B. Extraskeletal Manifestations of Primary Hyperparathyroidism Although the incidence of nephrolithiasis has declined since the 1960s, kidney stones remain the most common manifestation of symptomatic primary hyperparathyroidism. Klugman et al. place the incidence of kidney stones at 15–20% of all patients [39], and more recent studies confirm that frequency to be unchanged. Other renal manifestations of primary hyperparathyroidism include hypercalciuria, which is seen in approximately 40% of patients, and nephrocalcinosis, the frequency of

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which is unknown [40]. Classical primary hyperparathyroidism used to be associated with a distinct neuromuscular syndrome, characterized by type II muscle cell atrophy [41]. This is no longer seen [42]. The more non-specific neurobehavioral manifestations noted above, to be distinguished from the specific neuromuscular syndrome, remain under active investigation in an effort to determine whether an extra-skeletal component of primary hyperparathyroidism incorporates these more diffuse elements [31–34].

C. The Skeleton in Primary Hyperparathyroidism Osteitis fibrosa cystica is rarely seen. However, newer, more sensitive techniques have demonstrated that skeletal involvement in primary hyperparathyroidism is common. In this section, we review the profile of the skeleton in primary hyperparathyroidism as it is reflected by bone densitometry and bone histomorphometry. 1. Bone Densitometry

Bone densitometry is particularly well suited to the investigation of the skeleton in primary hyperparathyroidism. In addition to features such as sensitivity, accuracy, and precision, it is a non-invasive test that can be applied to sites containing different amounts of cortical and cancellous bone. Parathyroid hormone has long been known to be catabolic in cortical bone. Consistent with this idea, measurement of bone mass in primary hyperparathyroidism typically shows reduced bone density at the distal third of the radius, a site composed primarily of cortical bone [43]. Studies have also shown bone density to be relatively preserved at the lumbar spine, a site containing a preponderance of cancellous bone. The femoral neck, a more even mixture of cortical and cancellous bone, is intermediate in bone density between the lumbar spine and the distal radius (see Fig. 1). These data support two different actions of parathyroid hormone: a catabolic effect in cortical bone and an anabolic effect in cancellous bone. It should be noted that this pattern, also seen when bone mass is measured in postmenopausal women with primary hyperparathyroidism, represents a reversal of the pattern usually associated with postmenopausal bone loss, in which cancellous bone is lost first. Although the pattern of decreased bone mass at the radius and preserved bone mass at the spine is typical, there are those whose pattern of bone loss presents exceptions to the rule. We have patients with vertebral osteopenia when they are evaluated for primary hyperparathyroidism. In our natural history study, approximately 15% of patients had a lumbar spine Z score < – 1.5s [44]. Not all vertebral bone loss in this group could be attributed to estrogen

Figure 1 Bone densitometry in primary hyperparathyroidism. Data are shown in comparison to age- and sex-matched normal subjects. Divergence from expected values is different at each site (p = 0.0001). Reproduced from J. Bone Miner. Res. 1989; 4, 283–291 [43] with permission of the American Society for Bone and Mineral Research.

deficiency, as this group included both men and premenopausal women. It should also be noted that in the rare cases of severe primary hyperparathyroidism, seen today, more generalized skeletal demineralization is the rule, and the lumbar spine is not protected. The importance of bone densitometry in the evaluation of all patients with primary hyperparathyroidism has been formally recognized. The Consensus Development Conference on Asymptomatic Primary Hyperparathyroidism in 1990 [45], and the NIH Workshop on Asymptomatic Primary Hyperparathyroidism in 2002 [46] both included guidelines for surgical intervention based upon bone density measurements. 2. Bone Histomorphometry

Investigations at a histological level have supported and extended the data obtained by bone densitometry. Histomorphometric analysis of an iliac crest bone biopsy specimen showed cortical thinning consistent with the reduction of cortical bone suggested by densitometric evaluation [47–48]. Histomorphometric studies, however, have contributed most by elucidating the nature of cancellous bone in this disease. Cancellous bone volume is not only relatively preserved in primary hyperparathyroidism, it is actually increased as compared to normal subjects [47, 49]. In primary hyperparathyroidism, there is an increase in trabecular number and a decrease in trabecular separation [47]. Using a histomorphometric technique called “strut analysis,” it has been determined further that cancellous bone is preserved in primary hyperparathyroidism

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through the maintenance of well-connected trabecular plates [47, 50]. In order to determine the mechanism of cancellous bone preservation in primary hyperparathyroidism, static and dynamic histomorphometric indices were compared between normal and hyperparathyroid postmenopausal women. In normal postmenopausal women there is an imbalance in bone formation and resorption which favors excess bone resorption. In postmenopausal women with primary hyperparathyroidism, on the other hand, the adjusted apposition rate is increased. Bone formation, thus favored, may explain the efficacy of PTH at cancellous sites in patients with osteoporosis [51, 52]. Assessment of bone remodeling variables in patients with primary hyperparathyroidism shows increases in the active bone formation period [49]. The increased bone formation rate and total formation period may explain the preservation of cancellous bone seen in this disease. 3. Fractures

Whether or not mild primary hyperparathyroidism increases the risk of fracture, a well-described consequence of skeletal involvement in the classical disease, is unclear. If one could safely extrapolate from epidemiologic data correlating bone mass measurements with fracture risk, one would predict that patients with primary hyperparathyroidism would have increased risk of cortical bone fracture, and possibly a reduced risk of fracture in cancellous bone. In contrast, Khosla et al. [53] have reported a large retrospective series in which fracture risk at all sites was increased in primary hyperparathyroidism. Available data are uncertain because the numbers of patients studied in this regard are small, and existing studies have produced conflicting results [53–58]. What is needed at this time is a prospective study of fracture incidence in primary hyperparathyroidism.

IV. BONE MARKERS IN PRIMARY HYPERPARATHYROIDISM Histomorphometric evidence indicates that even in asymptomatic primary hyperparathyroidism, bone turnover is increased. Indices reflecting increased bone formation and bone resorption are readily demonstrable. Such direct observations would be expected to be demonstrable in the measurement of circulating and urinary markers of bone turnover. Both bone resorption and bone formation are increased by parathyroid hormone. Bone markers would be expected to accurately reflect skeletal metabolism in patients with primary hyperparathyroidism. Bone markers should also be helpful in the longitudinal assessment of

bone turnover in those patients who undergo parathyroidectomy as well as those who are monitored without surgery.

A. Bone Formation Markers Bone formation is reflected by products of osteoblast activity, including alkaline phosphatase, osteocalcin, and the procollagen peptide of type I collagen [59] (see also Chapter 34). Total alkaline phosphatase activity, readily available on routine multichannel panels, continues to be a useful marker in primary hyperparathyroidism [60–62]. Concentrations of alkaline phosphatase are typically elevated, but in asymptomatic individuals they are usually no more than 15–20% above normal. It seemed promising to consider the potential usefulness of the isoenzyme of alkaline phosphatase specific for osteoblastic action because total alkaline phosphatase reflects not only bone activity but also activity from other tissues such as the liver. Bonespecific alkaline phosphatase activity is clearly elevated in patients with mild primary hyperparathyroidism. In a limited study, bone-specific alkaline phosphatase correlated with parathyroid hormone levels and bone mineral density at the lumbar spine and femoral neck [61], but it was no more useful than the total alkaline phosphatase level. Osteocalcin is also generally increased in patients with primary hyperparathyroidism [62–65]. Osteocalcin correlates with other indices of bone formation. However, in our patients, we found no correlation of this marker with parathyroid hormone levels or with bone mineral density at any site. Although carboxy- and aminoterminal extension peptides of procollagen reflect osteoblast activation and bone formation, they have not been shown to have significant predictive or clinical utility in primary hyperparathyroidism [66]. In a small study of patients with primary hyperparathyroidism, carboxyterminal propeptide of human type 1 procollagen (PICP) levels were higher than in control subjects, but the deviations from the norm were much less impressive than those seen for alkaline phosphatase, osteocalcin, or even hydroxyproline [66].

B. Bone Resorption Markers Bone resorption is reflected in a product of osteoclast activity, tartrate-resistant acid phosphatase (TRAP), and in collagen breakdown products such as hydroxyproline, hydroxypyridinium crosslinks of collagen, and the N- and C-telopeptides of collagen metabolism [59] (see also Chapter 35). The oldest marker of bone resorption, urinary hydroxyproline excretion, is frankly elevated in patients

Chapter 46 Primary Hyperparathyroidism

with osteitis fibrosa cystica. However, in asymptomatic patients with mild, asymptomatic primary hyperparathyroidism, values of urinary hydroxyproline are often entirely normal. In addition, hydroxyproline does not reflect bone resorption exclusively, because its measurement can be influenced by diet, and assays for this amino acid have never been simplified for clinical convenience. The hydroxypyridinium crosslinks of collagen have replaced hydroxyproline excretion as markers of bone resorption in primary hyperparathyroidism. These crosslinks of collagen (pyridinoline and deoxypyridinoline) are clearly elevated in primary hyperparathyroidism [68]. The increase in deoxypyridinoline excretion in primary hyperparathyroidism is greater than that of pyridinoline, yet levels of both correlate with parathyroid hormone concentrations. The fact that urinary concentrations of pyridinoline and deoxypyridinoline seem to correlate closely with disease activity, coupled with their usefulness in longitudinal follow-up, make the hydroxypyridinium crosslinks of collagen an attractive choice as a bone resorptive marker in primary hyperparathyroidism. Assays for collagen crosslinked N-telopeptide levels have yet to be systematically explored in this disease. In the case of pyridinoline crosslinked telopeptide domain of type 1 collagen (ICTP), pooled data from patients with high turnover diseases (i.e., primary hyperparathyroidism as well as hyperthyroidism) suggest that this marker may reflect calcium kinetics [69] and histomorphometric indices [70]. Unfortunately, data on primary hyperparathyroidism alone are not available. Other reports of newer bone resorption markers in primary hyperparathyroidism are available. Studies of the TRAP in primary hyperparathyroidism are limited, although levels have been shown to be elevated [71]. One study reported a correlation between levels of parathyroid hormone and tartrate-resistant acid phosphatase in primary hyperparathyroidism [72]. Bone sialoprotein is a phosphorylated glycoprotein that makes up approximately 5–10% of the non-collagenous protein of bone (see Chapters 4, 8, and 35). It is elevated in high turnover states and reflects, in part, bone resorption. In primary hyperparathyroidism, bone sialoprotein levels are elevated and correlate with levels of urinary pyridinoline and deoxypyridinoline [73]. It is clear from the reports summarized in this section that markers of bone formation and resorption tend to be elevated, even in asymptomatic primary hyperparathyroidism. Thus, sensitive assays reflecting bone formation and bone resorption show both processes to be increased in mild primary hyperparathyroidism. It is not certain, however, whether quantification of these markers is useful

771 to predict the extent of disease severity generally or whether they are useful predictors of the course of the skeletal involvement.

C. Longitudinal Bone Markers in Prospective Studies of Primary Hyperparathyroidism Measurements of bone markers hold promise in primary hyperparathyroidism not only as indices of disease activity but also as a means by which patients can be followed after surgery. It could be expected that after successful surgery, these indices of disease activity would decline. Studies of bone markers in the longitudinal follow-up of patients with primary hyperparathyroidism are limited. However, those that have been done offer interesting insights. Seibel et al. [68], Guo et al. [74], and Tanaka et al. [75] all report declining levels of bone markers following surgery. The kinetics of change in bone resorption versus bone formation markers following parathyroidectomy have provided insight into the nature of postoperative anabolic skeletal effects observed by a number of investigators. Seibel et al. [68] have found that bone resorption markers decline rapidly following successful surgery, whereas indices of bone formation follow a slower decline. For example, urinary pyridinoline and deoxypyridinoline levels decline significantly as early as 2 weeks following parathyroidectomy, preceding the onset of reductions in alkaline phosphatase (see Fig. 2). Tanaka et al. [75] reported similar data, which showed a discrepancy between changes in urinary N-telopeptide excretion and osteocalcin following parathyroidectomy. Minisola et al. [67] reported a drop in bone resorptive markers but no significant change in alkaline phosphatase activity or osteocalcin. Several short-term studies reported a brief increase in PICP immediately following parathyroidectomy, whereas bone resorptive markers fell promptly [67, 76]. Such differences in the time course between the postoperative activities in bone resorption markers (rapid) and bone formation markers (slow) shift the dynamics of bone remodeling toward anabolic results. Ultimately, all indices of bone turnover return towards or into the normal range. The return of resorption and formation markers to normal favors a restoration of bone metabolism to normal. Elsewhere in this book, it is shown that reductions in bone turnover markers are independent predictors of gains in bone mass. With respect to the postoperative changes in primary hyperparathyroidism, it is likely that the reduction in bone turnover is associated with the mineralization of the enlarged remodeling space seen so commonly in the preoperative state.

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Figure 2

Bone turnover markers following parathyroidectomy. Data are presented as percentage change from preoperative baseline measurement. PYD is pyridinoline, DPD is deoxypyridinoline, ^ denotes significant change at p <0.05. Reprinted with permission from Seibel et al. [68], J. Clin. Endocrinol. Metab. 74, 481–486, 1992; © The Endocrine Society.

V. CYTOKINES IN PRIMARY HYPERPARATHYROIDISM In addition to studies of bone markers that have helped to clarify the skeletal dynamics of primary hyperparathyroidism, other investigations have sought to elucidate the mechanisms underlying these changes in bone markers. To this end, potential mediators of parathyroid hormone action in this disease have been identified. Candidate cytokine molecules such as interleukin-1 (IL-1), IL-6, and IL-II, TGF-alpha, EOF, and tumor necrosis factor, all stimulate bone resorption. Other factors, including IL-4, IGF-1, TGF-beta, and interferons, may be anabolic for bone (see Chapters 6 and 7). Measuring changes in the activity of some, or all, of these cytokines, may help elucidate the mechanism of bone loss in primary hyperparathyroidism. Attention has focused on interleukin-6 and tumor necrosis factor-alpha as possible mediators of bone resorption in primary hyperparathyroidism. In vitro and in vivo studies support the hypothesis that parathyroid hormone induces production of IL-6, which leads to increased osteoclastogenesis [77–79] (see Chapters 7 and 16). Antibodies to IL-6 prevent parathyroid hormone-mediated bone resorption. Reports from Rusinko et al. [80] and Grey et al. [77] have shown IL-6 and TNF-alpha levels to be elevated in primary hyperparathyroidism, to correlate well with parathyroid hormone levels, and to fall following parathyroidectomy. Although cytokine levels did not correlate with bone density measurements in this report, it should be noted that bone density was not assessed at the site containing most cortical bone (the radius), where the catabolic effects of excess parathyroid hormone would be expected

to be seen. These observations require confirmation with appropriate control subjects. Furthermore, little is known about the effect of primary hyperparathyroidism on other bone resorptive cytokines. Ongoing research efforts are also considering which cytokines might mediate, or potentiate, the anabolic effect of primary hyperparathyroidism on bone. One such factor is insulin-like growth factor-1 (IGF-1), which is known to be a direct mediator of parathyroid hormone action in bone [81–84]. It is possible that the anabolic properties of PTH at cancellous sites (e.g., lumbar spine) can be explained by IGF-1. IGF-1 and IGFBP3 (the major binding protein for IGF-1) have been documented to change after parathyroidectomy in primary hyperparathyroidism. The alteration in the ratios of these cytokines supports enhanced delivery of IGF-1 to the tissues following surgery, resulting in an increase in IGF-1 that is inversely proportional to the observed rise in lumbar spine and femoral neck bone density. Further work is needed to document the relationship between these changes and the effects of parathyroid hormone excess in the hyperparathyroid state, and to document the effect of its withdrawal after parathyroidectomy.

VI. TREATMENT OF PRIMARY HYPERPARATHYROIDISM A. Surgery Parathyroidectomy remains the only option for cure of primary hyperparathyroidism. With the change in clinical profile to a largely asymptomatic disorder in the latter

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half of the last century, reasonable questions were raised concerning the need for surgery in all patients. The National Institutes of Health Consensus Development Conference on the Management of Asymptomatic Primary Hyperparathyroidism, in 1991, developed guidelines for parathyroidectomy [45], which were revised a decade later (National Institutes of Health Workshop on Primary Hyperparathyroidism, 2002 [46]) based upon data accumulated in the intervening years. The current guidelines support parathyroidectomy in asymptomatic patients meeting the following criteria: (1) serum calcium greater than 1 mg/dl above the upper limits of normal; (2) marked hypercalciuria (>400 mg/day) or reduction in creatinine clearance by more than 30% below age- and sex-matched reference values; (3) markedly reduced bone density (T-score < –2.5 at any site); (4) age less than 50 years. A comparison between the older guidelines and the newer guidelines is provided in Table I. After parathyroidectomy, hypercalcemia resolves rapidly. Surgery is also of clear benefit in reducing the incidence of recurrent nephrolithiasis [39, 56]. Parathyroidectomy also leads to a major improvement in bone mineral density [86], averaging 12% at the lumbar spine and femoral neck (see Fig. 3). This increase is sustained over a decade after surgery. In the subset of patients with low vertebral bone density at the time of diagnosis, an even more dramatic improvement can be anticipated following parathyroidectomy [44]. For this reason, surgery is recommended in patients with low bone density regardless of the severity of other manifestations of primary hyperparathyroidism. The increase in bone density after successful parathyroidectomy might be expected to reduce fracture incidence. This would be the case if improvements in bone mass are associated with reduction in fracture incidence in this disease, as they are in some therapies for postmenopausal osteoporosis. However, no studies of fracture incidence after successful parathyroid surgery are large enough to provide this information in a definitive way.

Table I. A Comparison of New and Old NIH Recommended Guidelines for Surgery in Asymptomatic Primary Hyperparathyroidism Measurement Serum calcium (above upper limit of normal) 24-hour urinary calcium Creatinine clearance Bone mineral density Age

Guidelines, 1990 (45)

Guidelines, 2002 (46)

1–1.6 mg/dl

1.0 mg/dl

>400 mg Reduced by 30% Z-score <–2.0 (forearm) <50 years

>400 mg Reduced by 30% T-score < –2.5 (any sites) <50 years

B. Medical Management of Primary Hyperparathyroidism Many patients with asymptomatic hypercalcemia do not meet any of the National Institutes of Health Consensus Conference Guidelines for surgery and choose to be followed without parathyroidectomy. Data from our group and others have shown stability in patients with mild primary hyperparathyroidism followed over a decade of observation [33, 86, 87]. In a large, prospective study of patients followed without surgery, annual measurements showed no change in serum calcium, phosphorus, parathyroid hormone, vitamin D, or alkaline phosphatase levels and no change in urinary calcium, hydroxyproline, or hydroxypyridinium crosslink excretion. In sharp contrast to the change in bone mineral density seen in patients after parathyroidectomy, lumbar spine, femoral neck, and radius bone mineral density also showed stability in this group. Some patients did have a progressive worsening of their disease indices. Approximately 25% of patients developed a new indication for surgery while under observation. The only demographic, biochemical or bone densitometric predictor of this progressive course was young age (less than 50 years [88]). This fact confirmed the importance of age as a surgical guideline in this disease, and emphasizes the importance of close follow-up of biochemical and bone density studies in all patients who do not have surgery.

C. Drug Treatment of Primary Hyperparathyroidism 1. Phosphate

Oral phosphate can lower the serum calcium by up to 1 mg/dl [89, 90]. Concerns regarding limited gastrointestinal tolerance, possible further increases in PTH levels, and the possibility of soft tissue calcifications have limited the chronic use of oral parathyroid in primary hyperparathyroidism. 2. Bisphosphonates

Bisphosphonates, antiresorptive agents that reduce bone turnover, do not affect PTH secretion directly. Although they could theoretically reduce serum and urinary calcium levels, studies to date do not show a sustained effect on calcium levels [91–96]. Two controlled, double-blinded randomized trials of alendronate in primary hyperparathyroidism have demonstrated substantial gains in lumbar spine and hip bone density without any changes in bone density of the distal radius [95, 96]. Risedronate, another bisphosphonate used in the treatment of osteoporosis,

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lowered serum and urinary calcium, as well as hydroxyproline excretion significantly, while the PTH concentration rose in a short 7-day study of 19 patients [97]. 3. Estrogens and Serums

Estrogen replacement therapy has been used for postmenopausal women with primary hyperparathyroidism since the early 1970s. Treatment is associated with a 0.5–1.0 mg/dl reduction in total serum calcium levels but no change in PTH levels [98–100]. Various approaches to skeletal dynamics have been used in studies of metabolic and skeletal responses to estrogen replacement therapy in women with primary hyperparathyroidism. Markers of bone turnover, including alkaline phosphatase activity, urinary hydroxyproline, and urinary N-telopeptide, have been uniformly documented to decline. The role of cytokines in the skeletal response to estrogen therapy in primary hyperparathyroidism is unknown. While the local anabolic actions of PTH may be mediated in part by IGF-1, estrogens play an important independent role in regulating the synthesis of this cytokine. Estrogen treatment is associated with a beneficial effect on BMD at the femoral neck and lumbar spine [100]. Recently, there has been an increasing appreciation of the risks associated with estrogen use. In view of the fact that doses higher than the current standard have been most consistently associated with beneficial effects in primary hyperparathyroidism it is no longer advisable to begin HRT for the purpose of lowering calcium levels in primary hyperparathyroidism. Raloxifene, a selective estrogen receptor modulator, may be an alternative. Preliminary data come from a short-term (8-week) trial of 18 postmenopausal women, in which raloxifene (60 mg/day) was associated with a statistically significant although small (0.5 mg/dl) reduction in the serum calcium concentration and in markers of bone turnover [101]. 4. Calcimimetic Agents

Figure 3

Bone density by site over 10 years in patients with primary hyperparathyroidism. Data are for patients who underwent parathyroidectomy (O) or who were followed with no intervention (O). Data are presented as percentage change from baseline bone density measurement. *denotes significant change at p < 0.05. Reprinted with permission from Silverberg et al. (86), N Engl J Med. 341: 1249–1255.

Compounds that act on the parathyroid cell calciumsensing receptor, activated by increased extracellular calcium, can lead to inhibition of PTH secretion. The phenylalkylamine (R)-N(3-methoxy-α-phenylethyl)-3(2-chlorophenyl)-1-propylamine (R-568) is one such calcimimetic compound that has been shown to inhibit PTH secretion from postmenopausal women with primary hyperparathyroidism [102]. A second-generation ligand, cinacalcet, has been the subject of recent more extensive investigation in primary hyperparathyroidism. These studies have shown clearly that cinacalcet normalizes the serum calcium in most patients with mild hyperparathyroidism [103]. Interestingly, the PTH concentration falls but not to normal levels. These data suggest that a drug of this type

Chapter 46 Primary Hyperparathyroidism

may become a useful alternative to parathyroidectomy in patients with primary hyperparathyroidism.

VII. SUMMARY Despite the absence of radiologically evident bone disease, modern-day primary hyperparathyroidism has a clearly defined pattern of skeletal involvement. The diminution of cortical bone and preservation of cancellous bone documented in bone mineral density studies and histomorphometry are associated with increases in bone turnover. Increases in both bone formation and resorption are reflected in increased levels of bone turnover markers. Data on the role of certain cytokines as potential mediators of skeletal effect in primary hyperparathyroidism continue to provide insight into the pathogenesis of this important metabolic bone disease.

Acknowledgments This work was supported in part by National Institutes of Health Grants NlDDK 32333 and RR 00645.

References 1. Albright, H., and Reifenstein, E. C. (1948). “The Parathyroid Glands and Metabolic Bone Disease.” Williams & Wilkins, Baltimore, MD. 2. Cope, O. (1966). The story of hyperparathyroidism at the Massachusetts General Hospital. N. Engl. J. Med. 21, 1174–1182. 3. Bauer, W. (1933). Hyperparathyroidism: Distinct disease entity. J. Bone J. Surg. 15, 135–141. 4. Bauer, W., and Federman, D. D. (1962). Hyperparathyroidism epitomized: Case of Captain Charles E. Martell. Metab. Clin. Exp. 11, 21–22. 5. Mandl, F. (1925). Therapeutischer Versuch bei Ostitis fibrosa generalisata mittels Extirpation eines Epithelkoerperchentumors. Wien. Klin. Wochenschr. 50, 1343–1344. 6. Albright, P., Aub, J. C., and Bauer, W. (1934). Hyperparathyroidism common and polymorphic condition as illustrated by seventeen proved cases from one clinic. JAMA 102, 1276–1287. 7. Silverberg, S. J., and Bilezikian, J. P. (2001). Primary hyperparathyroidism. In: “Principles and Practice of Endocrinology and Metabolism” (K. L Becker, ed), pp. 564–573. Lippincott-Williams & Wilkins, New York. 8. Heath, H., Hodgson, S. F., and Kennedy, M. A. (1980). Primary hyperparathyroidism: Incidence, morbidity, and economic impact in a community. N. Engl. J. Med. 302, 189–193. 9. Mundy, G. R., Cove, D. H., and Friskin, R. (1980). Primary hyperparathyroidism: Changes in the pattern of clinical presentation. Lancet 1, 1317–1320. 10. Schulz, D. A., and Pummel, D. C. (1981). Asymptomatic primary hyperparathyroidism. Mayo Clin. Proc. 56, 473–478. 11. Silverberg, S. J., and Bilezikian, J. P. (2001). Clinical presentation of primary hyperparthyroidism in the United States. In: “The Parathyroids” (J. P Bilezikian, R. Marcus, M. A. Levine, eds), 2nd edn, pp. 349–360. Academic Press, New York.

775 12. Rio, S. D., Frame, B., Miller, M- J., Kleerekoper, M., Block, M. A., and Parfitt, A. M. (1980). Hyperparathyroidism following head and neck irradiation. Arch. Intern. Med. 140, 205–207. 13. Seely, E. W., Moore, T. J., LeBoff, M. S., and Brown, E. M. (1989). A single dose of lithium carbonate acutely elevates intact parathyroid hormone levels in humans. Acta Endocrinol. 121, 174–176. 14. Nordenstrom, J., Strigard, K., Perbeck, L., Willems, J., BagedahlStrindlund, M., and Under, J. (1992). Hyperparathyroidism associated with treatment of manic-depressive disorders by lithium. Ear. J. Surg. 158, 207–211. 15. McHenry, C. R., Rosen, I. B., Rotstein, L. E., Forbath, N., and Walfish, P. G. (1989). Lithiumogenic disorders of the thyroid and parathyroid glands as surgical disease. Surgery 108, 1001–1005. 16. Krivitzky, A., Bentata-Pessayre, M., Sarfati, E., Gardin, J. P., Callard, P., and Delzant, G. (1986). Multiple hypersecreting lesions of the parathyroid glands during treatment with lithium. Ann. Med. Intern. 137, 118–122. 17. Arnold, A., Staunton, C. E., Kirn, H. G., Gaz, R. D., and Kronenberg, H. M. (1988). Monoclonality and abnormal parathyroid hormone genes in parathyroid adenomas. N. Engl. J. Med. 318, 658–662. 18. Arnold, A., and Kirn, H. G. (1989). Molecular cloning and chromosomal mapping of DNA rearranged with the parathyroid hormone gene in a parathyroid adenoma. J. Clin. Invest. 83, 2034–2040. 19. Friedman, E., Bale, A. E., Marx, S. J. et al. (1990). Genetic abnormalities in sporadic parathyroid adenoma. J. Clin. Endocrinol. Metab. 71, 293–297. 20. Arnold, A., and Kim, H. G. (1989). Clonal loss of one chromosome 11 in a parathyroid adenoma. Clin. Endocrinol. Metab. 69, 496–499. 21. Cryns, V. L., Yi, S. M., Tahara, H., Gaz, R. D., and Arnold, A. (1995). Frequent loss of chromosome arm I p in parathyroid adenomas. Genes Chromosomes Cancer 13, 9–17. 22. Tahara, H., Smith, A. P., Gaz, R. D., Cryns, V. L., and Arnold, A. (1996). Genomic localization of novel candidate tumor suppressor gene loci in human parathyroid adenomas. Cancer Res. 56, 599–605. 23. Carling, T., Correa, P., Hessman, O., Hedberg, J., Skogseid, B., Lindberg, D., Rastad, J., Westin, G., and Akerstrom, G. (1999). Parathyroid MEN1 gene mutations in relation to clinical characteristics of non-familial primary hyperparathyroidism. J. Clin. Endocrinol. Metab. 83, 2960–2963. 24. Brown, E. M., Gardner, D. G., Brennan, M. F. et al. (1979). Calciumregulated parathyroid hormone release in primary hyperparathyroidism. Studies in vitro with dispersed parathyroid cells. Am. J. Med. 66, 923–931. 25. Lloyd, H. M., Parfitt, A. M., Jacobi, J. M. et al. (1989). The parathyroid glands in chronic renal failure: A study of their growth and other properties made on the basis of findings in patients with hypercalcemia. Lab. Clin. Med. 114, 358–367. 26. Ringe, J. D. (1984). Reversible hypertension in primary hyperparathyroidism: Pre- and postoperative blood pressure in 75 cases. Klin. Wochenschr. 62, 465–469. 27. Broulik, P. D., Horky, K., and Pacovsky, V. (1985). Blood pressure in patients with primary hyperparathyroidism before and after parathyroidectomy. Exp. Clin. Endocrinol. 86, 346–352. 28. Rapado, A. (1986). Arterial hypertension and primary hyperparathyroidism. Am. J. Nephrol. 6 (Suppl. 1), 46–50. 29. Bilezikian, J. P., Aurbach, G. D., Connor, T. B. et al. (1973). Pseudogout following parathyroidectomy. Lancet 1, 445–447. 30. Geelhoed, G. W., and Kelly, T. R. (1989). Pseudogout as a clue and complication in primary hyperparathyroidism. Surgery 106, 1036–1041. 31. Joborn C., Hetta, J., Johansson, H. et al. (1988). Psychiatric morbidity in primary hyperparathyroidism. World J. Surg. 12, 476–481. 32. Brown, G. G., Preisman, R. C., and Kleerekoper, M. D. (1987). Neurobehavioral symptoms in mild primary hyperparathyroidism: Related to hypercalcemia but not improved by parathyroidectomy. Henry Ford Med. 7, 211–215.

776 33. Roam, D. S., Phillips, E. R., Divine, G. W., and Talpos, G. B. (2004). Randomized controlled clinical trail of surgery vs no surgery in patients with mild asymptomatic primary hyperparathyroidism. J. Clin. Endocrinol. Metab. 89, 5414–5422. 34. Silverberg, S. J. (2002). Non-classical target organs in primary hyperparathyroidism. J. Bone Min. Res. 17:(suppl 2), N117–N125. 35. Silverberg, S. J., Brown, I., LoGerfo, P., Gao, P., Cantor, T., and Bilezikian, J. P. (2003). Clinical utility of an immunoradiometric assay for whole PTH (1–84) in primary hyperparathyroidism. J. Clin. Endocrinol. Metab. 88, 4725–4730. 36. Vieth, R., Bayley, T. A., Walfish, P. G., Rosen, I. B., and Pollard, A. (1991). Relevance of vitamin D metabolite concentrations in supporting the diagnosis of primary hyperparathyroidism. Surgery 110, 1043–1046. 37. Silverberg, S. J., Shane, E., Dempster, D. W., and Bilezikian, J. P. (1999). Vitamin D deficiency in primary hyperparathyroidism. Am. J. Med. 107, 561–567. 38. Mithal, A., Bandeira, F., Meng, X., Silverberg, S. J., Shi, Y., Mishra, S. K., Griz, L., Macedo, G., Celdas, G., Bandeira, C., Bilezikian, J. P., Rao, S. D. (2001). Clinical presentation of primary hyperparthyroidism: India, Brazil and China. In: The Parathyroids (Bilezikian JP, Marcus R, Levine MA, eds 2nd edition). New York: Academic Press, pp. 375–386. 39. Klugman, V. A., Favus, M., and Pak, C. Y. C. (1994). Nephrolithiasis in primary hyperparathyroidism. In “The Parathyroids: Basic and Clinical Concepts” (J. P. Bilezikian, ed.), pp. 505–518. Raven Press, New York. 40. Silverberg, S. J., Shane, E., Jacobs, T. P. et al. (1990). Nephrolithiasis and bone involvement in primary hyperparathyroidism. Am. J. Med. 89, 327–334. 41. Patten, B. M., Bilezikian, J. P., Mallette, L. E-, Prince, A., Engel, W. K., and Aurbach, G. D. (1989). The neuromuscular disease of hyperparathyroidism. Ann. Intern. Med. 80, 182–194. 42. Turken, S. A., Cafferty, M., Silverberg, S. J. et al. (1989). Neuromuscular involvement in mild, asymptomatic primary hyperparathyroidism. Am. J. Med. 87, 553–557. 43. Silverberg, S. J., Shane, E., De La Cruz, L. et al. (1989). Skeletal disease in primary hyperparathyroidism. J. Bone Miner. Rex. 4, 283–291. 44. Bilezikian, J. P., Potts, J. T., El-Hajj Fuleihan, G., Kleerekoper, M., Neer, R., Peacock, M., Rastad, J., Silverberg, S. J., Udelsman, R., and Wells, S. A. (2002). Summary statement from a workshop on asymptomatic primary hyperparathyroidism: a perspective for the 21st century. J. Clin. Endocrinol. Metab. 87, 5353–5361. 45. National Institutes of Health. (1991). Consensus development conference statement on primary hyperparathyroidism. J. Bone Miner. Res. 6, s9–s13. 46. Silverberg, S. J., Locker, F. G., and Bilezikian, J. P. (1996). Vertebral osteopenia: A new indication for surgery in primary hyperparathyroidism. J. Clin. Endocrinol. Metab. 81, 4007–4012. 47. Parisien, M., Silverberg, S. J., Shane, E. et al. (1990). The histormorphometry of bone in primary hyperparathyroidism: Preservation of cancellous bone structure. Clin. Endocrinol. Metab. 70, 930–938. 48. Parfitt, A. M. (1989). Surface specific bone remodeling in health and disease. In “Clinical Disorders of Bone and Mineral Metabolism” (M. Kleerekoper, ed.), pp. 7–14. Mary Ann Liebert, New York. 49. Dempster, D. W., Parisien, M., Silverberg, S. J., et al. (1999). On the mechanism of cancellous bone preservation in postmenopausal women with mild primary hyperparathyroidism. J. Clin. Endocrinol. Metab. 84, 1562–1566. 50. Parisien, M., Cosman, F., Mellish, R. W. E., et al. (1995). Bone structure in postmenopausal hyperparathyroid, osteoporotic and normal women. J. Bone Miner. Res. 10, 1393–1399. 51. Dempster, D. W., Cosman, F., Kurland, E. S., Zhou, H., Nieves, J., Woelfert, L., Shane, E., Plavetic, K., Muller, R., Bilezikian, J., and Lindsay, R. (2001). Effects of daily treatment with parathyroid

Shonni J. Silverberg and John P. Bilezikian

52.

53.

54.

55.

56.

57.

58.

59.

60. 61.

62.

63.

64.

65.

66.

67.

68.

69.

hormone on bone microarchitecture and turnover in patients with osteoporosis: a paired biopsy study. J. Bone Min. Res. 16, 1846–1853. Neer, R. M., Arnaud, C. D., Zanchetta, J. R., Prince, R., Gaich, G. A., Reginster, J. Y., Hodsma, A. B., Eriksen, E. F., Ish-Shalom, S., Genant, H. K., Wang, O., and Mitalk, B. H. (2001). Effect of parathyroid hormone (1–34) on fractures and bone mineral density in postmenopausal women with osteoporosis. N. Engl. J. Med. 344, 1434–1441. Khosla, S., Melton, L. J., Wermers, R. A., et al. (1999). Primary hyperparathyroidism and the risk of fracture: A population-based study. J. Bone Miner. Res. 14, 1700–1707. Kochesberger, G., Buckley, N. J., Leight, G. S., et al. (1987). What is the clinical significance of bone loss in primary hyperparathyroidism. Arch. Intern. Med. 147, 1951–1953. Wilson, R. J., Rao, S., Ellis, B., et al. (1988). Mild asymptomatic primary hyperparathyroidism is not a risk factor for vertebral fractures. Ann. Intern. Med. 109, 959–962. Larsson, K., Ljunghall, S., Krusemo, U. B., et al. (1993). The risk of hip fractures in patients with primary hyperparathyroidism: A populationbased cohort study with a follow-up of 19 years. J. Intern. Med. 234, 585–593. Kenny, A. M., MacGillivray, D. C., Pilbeam, C. C., et al. (1995). Fracture incidence in postmenopausal women with primary hyperparathyroidism. Surgery 118, 109–114. Dauphine, R. T., Riggs, B. L., and Scholz, D. A. (1975). Back pain and vertebral crush fractures: An unemphasized mode of presentation for primary hyperparathyroidism. Ann. Intern. Med. 83, 365–367. Deftos, L. J. (1994). Markers of bone turnover in primary hyperparathyroidism. In “The Parathyroids: Basic and Clinical Concepts” (J- P. Bilezikian, ed.), pp. 485–492. Raven Press, New York. Moss, D. W. (1992). Perspectives in alkaline phosphatase research. Clin. Chem. 38, 2486–2492. Silverberg, S. J., Deftos, L, J., Kim, T., and Hill, C. S. (1991). Bone alkaline phosphatase in primary hyperparathyroidism. J. Bone Miner. Res. 6, A624. Duda, R. J., O’Brien, J. K., Katzman, J. A., Paterson, J. M., Mann, K. G., and Riggs, B. L. (1988). Concurrent assays of circulating bone Gla-protein and bone alkaline phosphatase: Effects of sex, age, and metabolic bone disease. J. Clin. Endocrinol. Metab. 5, 1–7. Price, P. A., Parthemore, J. G., and Deftos, L. J. (1980). New biochemical marker for bone metabolism. Measurement by radioimmunoassay of bone Gla-protein in the plasma of normal subjects and patients with bone disease. J. Clin. Invest. 66, 878–883. Deftos, L. J., Parthemore, J. G., and Price, P. A. (1982). Changes in plasma bone Gla-protein during treatment of bone disease. Calcif. Tissue Int. 34, 121–124. Eastell, R., Delmas, P. D., Hodgson, S., Eriksen, E. E, Mann, K. M., and Riggs, B. L. (1988). Bone formation rate in older normal women: Concurrent assessment with bone histomorphometry, calcium kinetics, and biochemical markers. J. Clin. Endocrinol. Metab. 67, 741–748. Ebeling, P. R., Peterson, J. M., and Riggs, B. L. (1992). Utility of type 1 procollagen propeptide assays for assessing abnormalities in metabolic bone diseases. J. Bone Miner. Res. 7, 1243–1250. Minisola, S., Romagnoli, E., Scarnecchia, L., Rosso, R., Pacitti, M. T.-, Scarda, A., and Mazzuoli, G. (1994). Scrum CITP in patients with primary hyperparathyroidism: Studies in basal conditions and after parathyroid surgery. Eur. J. Endocrinol. 130, 587–591. Seibel, M. J., Gartenberg, F., Silverberg, S. J., Ratcliffe, A., Robins, S. P., and Bilezikian, J. P. (1992). Urinary hydroxypyridinium crosslinks of collagen in primary hyperparathyroidism. Clin. Endocrinol. Metab. 74, 481–486. Charles, P., Mosekilde, L., Risteli, L., and Eriksen, E. F. (1994). Assessment of bone remodeling using biochemical indicators of type 1 collagen synthesis and degredation: Relation to calcium kinetics. Bone Miner. 24, 81–94.

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Chapter 46 Primary Hyperparathyroidism 70. Eriksen, E. F., Charles, P., Melsen, K., Mosekilde, L., Ristei, L., and Risteli, J. (1993). Serum markers of type 1 collagen formation and degredation in metabolic bone diseases: Correlation with bone histomorphometry. J. Bone Miner. Res. 8, 127–132. 71. Kraenzlin, M. E., Lau, K. H. W., Liang, L. et al. (1990). Development of an immunoassay for human serum osteoclastic tartrate-resistant acid phosphatase. Clin. Endocrinol. Metab. 71, 442. 72. De La Piedra, C., Diaz Martin, M. A., Diaz Diego, E. M., Lopez Gavilanes, E., Gonzalez Parra, E., Carmelo, C., and Rapado, A. (1994). Serum 1CTP, serum tartrate resistant acid phosphatase, and serum levels of intact FIH in parathyroid hyperfunction. Scand. J. Clin. Lab. Invest. 54, 11–15. 73. Seibel, M. J., Woigte, H. W., Pecherstorfer, M., Karmatschek, M., Horn, E., Ludwig, M., Armbruster, F. P., and Ziegler, R. (1996). Serum immunoreactive bone sialoprotein as a new marker of bone turnover in metabolic and malignant diseases. J. Clin. Endocrinol. Metab. 81, 3289–3294. 74. Guo, C. Y., Thomas, W. E. R., Al-Dehaimi, A. W., Assiri, A. M. A., and Eastell, R. (1996). Longitudinal changes in bone mineral density and bone turnover in women with primary hyperparathyroidism. J. Clin. Endocrinol. Metab. 81, 3487–3491. 75. Tanaka, Y., Funahashi, H., Imai, T., Tominga, Y., and Takagi, H. (1997). Parathyroid function and bone metabolic markers in primary and secondary hyperparathyroidism. Semin. Surg. Oncol. 13, 125–133. 76. Hoshino, H., Kushida, K., Takahashi, M., Denda, M., Yamazaki, K., Yamanashi, A., and Inoue, T. (1997). Short term effect of parathyroidectomy on biochemical markers in primary and secondary hyperparathyroidism. Miner. Electrolyte Metab. 23, 93–99. 77. Grey, A., Mitnick, M., Shapses, S., Gundberg, C., and Insogna, K. (1996). Circulating levels of IL-6 and TNF-alpha are elevated in primary hyperparathyroidism and correlate with markers of bone resorption. J. Clin. Endocrinol. Metab. 81, 3450–3454. 78. Roodman, G. D. (1992). Interleukin-6: An osteotropic factor? Bone Miner. Res. 1, 475–478. 79. Greenfield, E. M., Shaw, S. M., Gomik, S. A., and Banks, M. A. (1995). Adenyl cyclase and interleukin 6 are downstream effectors of PTH resulting in stimulation of bone resorption. J. Clin. Invest. 96, 1238–1244. 80. Rusinko, R., Yin, J. J., Yee, J., and Saad, G. R. (1995). PTH excess is associated with increased IL-6 and soluble IL-6 receptor production. Bone Miner. Res. 10 (Suppl. 1), T585A. 81. Jones, J. L., and Clemmons, D. R. (1995). Insulin-like growth factors and their binding proteins. Endocr. Rev. 15, 3–34. 82. Spencer, E. M., Si, E. C., Liu, C- C., and Howard, G. A. (1989). PTH potentiates the effect of IGF-1 on bone formation. Acta Endocrinol. 121, 435–442. 83. McCarthy, T. L., Centrella, M. C., and Canalis, E. (1989). PTH enhances the transcript and polypetide levels of IGF-1 in osteoblast enriched cultures from fetal rat bone. J. Clin. Endocrinol. Metab. 124, 1247–1253. 84. Pfeilschifter, J., Laukhuf, P., Muller-Beckmann, B., Blum, W. F, Pfister, T., and Ziegler, R. (1995). PTH increases the concentration of IGF-1 and TGF-B1 in rat bone. J. Clin. Invest. 96, 767–774. 85. Deaconson, T. E, Wilson, S. D., and Lemann, J. (1987). The effect of parathyroidectomy on the recurrence of nephrolithiasis. Surgery 215, 241–251. 86. Silverberg, S. J., Shane, E., Jacobs, T. P., Siris, E., and Bilezikian, J. P. (1999). The natural history of treated and untreated asymptomatic primary hyperparathyroidism: a ten-year prospective study. N. Engl. J. Med. 341, 1249–1255. 87. Rao, D. S., Wilson, R. J., Kleerekoper, M., and Parfitt, A. M. (1988). Lack of biochemical progression or continuation of accelerated bone

88.

89. 90.

91.

92.

93.

94.

94a.

95.

96.

97.

98.

99.

100.

101.

102.

103.

loss in mild asymptomatic primary hyperparathyroidism. J. Clin. Endocrinol. Metab. 67, 1294–1298. Silverberg, S. J., Brown, I., and Bilezikian, J. P. (2002). Youthfulness as a criterion for surgery in primary hyperparathyroidism. Am. J. Med. 113, 681–684. Purnell, D. C., Scholz, D. A., Smith, L. M. et al. (1984). Treatment of primary hyperparathyroidism. Am. J. Med. 56, 800–809. Broadus, A. E., Magee, J. S., Mallette, L. E. et al. (1983). A detailed evaluation of oral phosphate therapy in selected patients with primary hyperparathyroidism. J. Clin. Endocrinol. Metab. 56, 953–961. Kaplan, R. A., Geho, W. B., Poindexter, C. et al. (1977). Metabolic effects of diphosphonate in primary hyperparathyroidism. J. Clin. Pharmacol. 17, 410–419. Shane, E., Baquiran, D. C., and Bilezikian, J. P. (1981). Effects of dichloromethylene diphosphonate on serum and urinary calcium in primary hyperparathyroidism. Ann. Intern. Med. 95, 23–27. Hassani, S., Braunstein, G. D., Seibel, M. J., Brickman, A. S., Geola, F. L., Pekary, A. E., and Hershman, J. M. (2001). Alendronate therapy of primary hyperparathyroidism. Endocrinologist 11, 459–464. Rossini, M., Gatti, D., Isaia, G., Sartori, L., Braga, V., and Adami, S. (2001). Effects of oral alendronate in elderly patients with osteoporosis and mild primary hyperparathyroidism. J. Bone Miner. Res. 16, 113–119. Parker, C. R., Blackwell, P. J., Fairbairn, K. J., and Hosking, D. J. (2002). Hyperparathyroid-Related osteoporosis: A 2-Year Study. J. Clin. Endocrinol. Metab. 87, 4482–4489. Chow, C. C., Chan, W. B., Li, J. K. Y., Chan, N. N., Chan, M. H. M., Ko, G. T. C., Lo, K. W., and Cockram, C. S. (2003). Oral alendronate increases bone mineral density in postmenopausal women with primary hyperparathyroidism. J. Clin. Endocrinol. Metab. 88, 581–587. Kahn, A. A., Bilezikiam, J. P., Kung, A. W. C., Ahmed, M. M., Dubois, S. J., Ho, A. Y. Y., Schussheim, D., Rubin, M. R., Shaikh, A. M., Silverberg, S. J., Standish, T. I., Syed, Z., and Syed, Z. A. (2004). Alendronate in primary hyperparathyroidism: a double-blind, randomized, placebo-controlled trial. J. Clin. Endocrinol. Metab. Gallagher, J. C., and Nordin, B. E. C. (1972). Treatment with oestrogens of primary hyperparathyroidism in postmenopausal women. Lancet 1, 503–507. Marcus, R. (2002). The role of estrogens and related compounds in the management of primary hyperparathyroidism. J. Bone Min. Res. 17 (Suppl 2), N146–N149. Selby, P. L., and Peacock, M. (1986). Ethinyl estradiol and norethinedrone in the treatment of primary hyperparathyroidism in postmenopausal women. N. Engl. J. Med. 314, 1481–1485. Grey, A. B., Stapleton, J. P., Evans, M. C. et al. (1996). Effect of hormone replacement therapy on BMD in postmenopausal women with primary hyperparathyroidism. Ann. Intern. Med. 125, 360–368. Rubin, M. A., Lee, K. H., McMahon, D. J., and Silverberg, S. J. (2003). Raloxifene lowers serum calcium and markers of bone turnover in postmenopausal women with primary hyperparathyroidism. J. Clin. Endocrinol. Metab. 88, 1174–1178. Silverberg, S. J., Marriott, T. B., Bone, H. G. III, et al. (1997). Short term inhibition of parathyroid hormone secretion by a calcium receptor agonist in primary hyperparathyroidism. N. Engl. J. Med. 307, 1506–1510. Shoback, D. M., Bilezikian, J. P., Turner, S. A., McCary, L. C., Guo, M. D., and Peacock, M. (2003). The calcimimetic AMG 073 normalizes serum calcium in patients with primary hyperparathyroidism. J. Clin. Endocrinol. Metab. 88, 5644–5649.

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Chapter 47

Paget’s Disease of Bone Andreas Grauer Ethel Siris Stuart Ralston

Procter & Gamble Pharmaceuticals Mason, OH, USA Department of Medicine, Columbia University College of P&S, New York, NY, USA Rheumatic Diseases Unit, University of Edinburgh, Edinburgh, UK

I. Introduction II. Etiology and Pathogenesis

III. Treatment References

I. INTRODUCTION

incidence and severity of the disease at presentation are declining [6, 7].

Paget’s disease of bone is a chronic, localized disease, which can be monostotic or polyostotic. It is characterized by increased bone remodeling, bone hypertrophy, and abnormal bone structure. Patients may suffer from bone pain and deformity. Complications involve fractures, osteoarthritis, nerve compression syndromes, and neoplastic transformation [1]. The disorder rarely presents before the age of 40. Thereafter it is found with increasing prevalence with advancing age. Large autopsy [2] or radiographic studies [3] suggest a prevalence of approximately 3% in the population over 40 years, which increases up to 10% by the age of 95. Interestingly there is a marked geographic variation. The disorder is particularly common in Great Britain, where the prevalence of the disease among hospital patients is 5% of the population over 50 [4]. In other parts of western and central Europe, and regions of the world influenced by European migration including the United States, Australia and New Zealand, Argentina, and South Africa, the prevalence of the disease is estimated to be 1–3% of the population over 50 [5]. The disorder seems to be extremely rare in Nordic countries [4], the Middle East, China, and Japan, and most of sub-Saharan Africa. Studies suggest that both Dynamics of Bone and Cartilage Metabolism

II. ETIOLOGY AND PATHOGENESIS The marked ethnic differences in prevalence of Paget’s disease of bone, coupled with strong evidence of familial aggregation and changes in incidence of the disease over the past 30 years indicate that genetic and environmental factors both contribute to the pathogenesis of Paget’s disease. It is currently believed that, whilst genetic susceptibility clearly plays a critical role, there must be additional local factors that trigger, where and when Paget’s disease occurs in individual patients [8].

A. Role of Genetic Factors in Paget’s Disease Genetic factors have long been suspected to play a role in the etiology of Paget’s disease [9]. In an epidemiologic study of patients with Paget’s disease of bone from the USA, Siris reported that the rate of Paget’s disease was approximately seven times higher in relatives of cases as compared with relatives of controls [10]. The risk was 779

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increased to approximately 20 times greater than controls for relatives of cases that had deforming disease with an early age at diagnosis [10]. An earlier study by Sofaer in the UK came to essentially the same conclusion, in showing a 10-fold higher incidence of Paget’s disease amongst parents and siblings of index cases, when compared with parents and siblings of spouses [11]. In most families that have been reported, the disease is inherited in an autosomal dominant manner, with high penetrance by the sixth decade [12–16]. 1. Paget’s Disease of Bone and Related Syndromes

Several rare bone disorders have also been described which exhibit clinical, radiological, or histological features in common with Paget’s disease (Table I). These include; familial expansile osteolysis (FEO) [17], expansile skeletal hyperplasia (ESH) [18], early-onset Paget’s disease [19], juvenile Paget’s disease (also known as idiopathic hyperphosphatasia) [20], and the syndrome of hereditary inclusion body myopathy, Paget disease of bone, and frontotemporal dementia (IBMPFD) [21]. With the exception of juvenile Paget’s disease, which is an autosomal recessive condition, all of these disorders are inherited in an autosomal dominant manner. 2. Susceptibility Loci for Paget’s Disease of Bone and Related Syndromes

Several susceptibility loci for Paget’s disease and related syndromes have now been identified by genome-wide searches and candidate locus linkage studies (Table II). These include the PDB2 locus on chromosome 18q21 [13, 15, 22]; the PDB3 locus on chromosome 5q35 [16, 23]; the PDB4 locus on chromosome 5q31 [16]; the PDB5 locus on chromosome 2p36 [23]; the PDB6 locus on chromosome 10p13 [23]; and the PDB7 locus on chromosome 18q23 [24]. Previous reports of linkage of Paget’s disease to the HLA region on chromosome 6 (PDB1) [25] have not been confirmed in any of the genome scans performed over recent years, indicating Table I. Rare Bone Disorders which have Clinical, Radiological, or Histological Features in Common with Paget’s Disease Autosomal dominant Familial expansile osteolysis (FEO) [17] Expansile skeletal hyperplasia (ESH) [18] Early-onset Paget’s disease [19, 20] IBMPFD (syndrome of hereditary inclusion body myopathy, Paget’s disease of bone, and frontotemporal dementia) [21] Autosomal recessive Juvenile Paget’s disease (also known as idiopathic hyperphosphatasia) [20]

Table II.

Susceptibility Loci for Paget’s Disease and Related Syndromes*

PDB2 locus PDB3 locus PDB4 locus PDB5 locus PDB6 locus PDB7 locus Juvenile Paget’s disease IBMPFD Familial expansile osteolysis to chromosome

18q21 [13, 15, 22] 5q35 [16, 23] 5q31 [16] 2p36 [23] 10p13 [23] 18q23 [24] 8q24 [26] 9p21 [21] 18q21 (PDB2) [22]

*Previous reports of linkage of Paget’s disease to the HLA region on chromosome 6 (PDB1) [25] have not been confirmed in any of the genome scans performed over recent years, indicating that this result is likely a false positive.

that this result is likely a false positive. Juvenile Paget’s disease has been localized to chromosome 8q24 by genome-wide search [26]; IBMPFD to chromosome 9p21 [21], and familial expansile osteolysis to chromosome 18q21 (PDB2) [22]. 3. Genes Involved in Paget’s Disease of Bone and Related Syndromes

Positional cloning studies have now identified many of the genes responsible for Paget’s disease and related disorders and the emerging picture is that the causal genes are either involved in the receptor activator of nuclear factor kappa B (RANK) signaling pathway, and/or in the ubiquitin pathway (Fig. 1). Hughes and colleagues identified mutations in the TNFRSF11A gene encoding RANK as the cause of familial expansile osteolysis and early-onset Paget’s disease using a positional candidate approach. They described an 18 bp insertion mutation affecting the signal peptide of RANK, as the cause of FEO and 27 bp mutation in the same region as the cause of early-onset PDB [27, 28]. Both mutations resulted in failure of signal peptide cleavage and were found to cause NFκB activation in a promoter-reporter assay system in vitro [27]. Subsequently, Hughes and Whyte reported that expansile skeletal hyperplasia was caused by a 15 bp insertion mutation affecting the same region of RANK [29]. Other groups have also reported 18 bp insertion mutations affecting the RANK signal peptide which are similar [30] or identical [31] to those described by Hughes in patients with FEO. The gene responsible for juvenile Paget’s disease was discovered using a candidate gene approach. Promoted by the observation that RANK mutations cause FEO, Whyte and colleagues screened the TNFRSF11B gene encoding osteoprotegerin (OPG) for mutations in two apparently unrelated Navajo individuals with juvenile Paget’s disease and found that both patients were

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Figure 1

Proteins and signaling pathways affected by the known gene defects in Paget’s disease of bone and related syndromes. Positional cloning studies have now identified many of the genes responsible for Paget’s disease and related disorders and the emerging picture is that the causal genes are either involved in the receptor activator of nuclear factor kappa B (RANK) signaling pathway, and/or in the ubiquitin pathway.

homozygous for an identical 100 Kb deletion involving TNFRSF11B on chromosome 8q24 [32]. In an independent study, Cundy and colleagues reported a homozygous mutation affecting exon 3 of TNFRSF11B as the cause of juvenile Paget’s disease in three affected siblings with the disease [26]. Functional studies showed that the mutant OPG protein was unable to inhibit bone resorption when expressed in vitro, indicating that it was an inactivating mutation. The gene for IBMPFD (frontotemporal dementia) was also discovered by a positional candidate gene approach by mutation screening of several genes in the 9p21 locus identified by Kovach [21]. Analysis of individuals from 16 families identified several different missense mutations in exons 3, 5, and 6 of the valosin containing protein (VCP) gene which segregated with the disease in affected family members [33]. All of the mutations affected highly conserved amino acid residues clustered in the N-terminal domain of the VCP gene product [34], which is known to be involved in ubiquitin binding. This finding is of interest in relation to the fact the VCP is known to regulate degradation of the IKB alpha protein, which is involved in NFκB signaling [35]. Mutations of RANK and OPG have been excluded as a cause of late-onset familial or sporadic Paget’s disease [36–38] and three groups of investigators failed to detect evidence of an allelic association between common polymorphisms of RANK and sporadic Paget’s disease [27, 39, 40]. There is evidence to suggest that polymorphic

variation at the TNFRSF11B locus might predispose to Paget’s disease however. In a small study of Belgian patients, Wuyts found evidence of an association between a common polymorphism in intron 2 of TNFRSF11B and Paget’s disease [40]. Whilst another study by Daroszewska failed to confirm this specific finding in British patients, strong evidence of allelic association between a common polymorphism causing an lysineasparagine amino acid change at codon 3 of the OPG gene was reported in a large case-control study and a family based study, indicating that this common polymorphism might influence susceptibility to both familial and sporadic Paget’s disease [41]. Positional cloning studies of the chromosome 5q35 (PBD3) locus by both Laurin [42] and Hocking [43] resulted in the identification of mutations in the sequestosome 1 (SQSTM1) gene as a common cause of familial and sporadic Paget’s disease. A proline-leucine mutation affecting codon 392 (P392L) in the ubiquitin-associated (UBA) domain of SQSTM1 was initially identified as a cause of familial and sporadic Paget’s disease by Laurin in the French-Canadian population [42] and subsequently, this and other mutations affecting the UBA domain of SQSTM1 were found in patients with familial and sporadic Paget’s disease of predominantly British descent [43]. Over recent months, eight different mutations of SQSTM1 have been identified in Paget’s disease and all of these cluster in the UBA domain [44–47]. On the basis of current information, it appears that 30–50% of patients

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with familial PDB and 10–30% of patients with “sporadic” PDB harbor mutations in SQSTM1 [42, 44, 47]. The SQSTM1 gene encodes p62 which is a scaffold protein that plays an important role in NFκB signaling through the IL-1 receptor, TNF receptor, and nerve growth factor receptor [48]. Whilst all of the mutations identified so far affect the so-called ubiquitin-associated (UBA) domain of the molecule, there does not appear to be a clear correlation between the ubiquitin-binding properties of the isolated mutant UBA domains and the propensity to cause Paget’s. For example, the P392L mutation, which is the commonest cause of Paget’s, appears to be able to bind ubiquitin normally [49], as does the G411S mutation [44]. It should be noted however, that these ubiquitin-binding studies were done with the isolated UBA domain and it could be that the mutations cause a subtle conformational change in the whole protein that affects the ubiquitin-binding properties. Whilst the exact mechanisms by which these UBA mutations of SQSTM1 cause Paget’s disease are unclear, studies in mice with targeted inactivation of SQSTM1 have shown impaired osteoclastogenesis in response to RANKL in vitro, and impaired osteoclastogenesis in response to PTHrP injection in vivo, indicating that p62 plays a role in regulating osteoclast function and activity in response to at least two osteoclastogenic stimuli [50].

B. Role of Viral Infection in Paget’s Disease Interest in the possible role of viruses in the pathogenesis of Paget’s disease was stimulated some 30 years ago by the observation that osteoclast nuclei in Paget’s disease contained characteristic nuclear inclusions which were thought to resemble the nucleocapsids of paramyxoviruses [51, 52]. These observations led to speculation that Paget’s disease may be due to a slow virus infection affecting osteoclasts. Although these are very typical of Paget’s disease, they are not totally specific since they have also been found in the osteoclasts of patients with benign osteopetrosis [53] and pycnodysostosis [54] in macrophages from patients with primary oxalosis [55] and in osteoblasts from patients with otosclerosis [56]. Many attempts have been made to detect evidence of paramyxovirus protein and/or nucleic acids in cells and bone tissue from patients with Paget’s disease, with mixed results. The earliest studies employed immunohistochemistry and showed positive staining with antibodies directed against measles virus (MV) and respiratory syncytial virus (RSV) in cultured osteoclast-like cells from Paget’s patients [57–59]. In two studies, antibody staining for both viruses was observed in the same samples [57, 60].

Other groups have focused on the possibility that canine distemper virus (CDV), rather than measles virus, might be responsible for Paget’s disease, based on the observation that patients with Paget’s disease were more likely to have owned a dog than non-pagetic controls [61]. Whilst this epidemiologic association has not been replicated in other studies in the UK or USA [62, 63], proponents of the CDV hypothesis have found evidence of canine distemper virus (CDV) mRNA in pagetic bone samples from British patients with PDB using in situ hybridization [64]; reverse-transcription/PCR-in situ hybridization (RT/PCR-ISH) [65] and RT/PCR [66]. Positive in situ hybridization for MV has also been found in pagetic bone [67] and in a subset of peripheral blood cells from pagetic patients [68]. Other workers have failed to detect MV or CDV in pagetic bone, peripheral blood, or cultured bone marrow using RT/PCR [69–72] or using RT/PCR-ISH [73]. The conflicting results between laboratories in their ability to detect paramyxovirus transcripts in bone, tissue, and blood cells from patients with Paget’s disease, has been a source of much debate. Proponents of the viral hypothesis have argued that the presence of mutations in PCR generated DNA products from pagetic samples is consistent with the fact that the disease is caused by a persistent virus infection [66, 74, 75]. Opponents have argued that the detection of viral transcripts could simply be the result of PCR contamination and that the apparent mutations could represent the result of PCR incorporation errors [76]. This issue remains unresolved at present. In the most comprehensive study performed to date, however, Helfrich and colleagues found no evidence of paramyxovirus protein or nucleic acids in pagetic bone samples or peripheral blood using a wide range of techniques including nested RT/PCR, immunohistochemistry, and in situ hybridization [72]. Helfrich also noted that the nuclear inclusions in osteoclast nuclei in Paget’s disease were different from those in subacute sclerosing panencephalitis, the prototypical slow virus disease caused by measles [72]. Although there is lack of agreement about whether paramyxovirus is present in pagetic bone or not, there is good evidence to suggest that paramyxoviruses and viral proteins can modulate differentiation and function of osteoclasts. Infection of canine bone marrow cells with CDV in vitro has been shown to increase formation of osteoclast-like cells [77]. Similarly, measles virus has been found to stimulate osteoclast formation and activity in vitro, when the virus was used to infect bone marrow cells from transgenic mice which expressed the CD46 cellular receptor for measles virus [78]. Moreover, retroviralinduced expression of the MV nucleocapsid protein in cultured bone marrow cells has been shown to stimulate

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osteoclast formation, multinuclearity, and resorptive activity [79].

C. Cellular and Molecular Mediators of Paget’s Disease Whilst Paget’s disease is characterized by increased osteoclastic bone resorption and osteoblastic formation, the primary cellular abnormality in Paget’s disease is generally considered to reside within the osteoclast. This stems from the observation that osteoclast precursor cells obtained from pagetic marrow aspirates and peripheral blood, form multinucleated cells in culture at lower concentrations of 1,25(OH)2D3 and RANKL than comparable cells of normal subjects [80–83]. Previous work has suggested that these abnormalities of osteoclast formation were driven by the cytokine IL-6 which was reported to be increased concentrations in plasma and conditioned media from pagetic bone marrow cultures when compared with control cultures [84]. Recent studies do not particularly support a role for IL-6 as a primary factor in the pathogenesis of Paget’s disease however. Studies by Neale [83] and Natale [85] found no difference in circulating IL-6 levels in Paget’s patients when compared with controls and Neal further reported that neutralizing antibodies to IL-6 had no effect on osteoclast formation or activity in human osteoclasts generated from Paget’s patients [83]. The c-fos oncogene has been implicated in the pathogenesis of Paget’s disease in view of its association with osteosarcoma and because of the fact that expression of c-fos is increased in pagetic osteoclasts compared with non-pagetic osteoclasts [86]. In light of recent advances in understanding the genetic basis of Paget’s disease, it seems likely that the hyperresponsiveness to RANKL, 1,25(OH)2D3, and increased c-fos expression noted above could be explained on the basis of upregulation of the RANK–NFκB signaling pathway. Vitamin D analogs, which bind to the vitamin D receptor with high affinity without intrinsic activity could therefore be interesting targets for a specific therapy of Paget’s disease of bone [8, 87, 88].

D. Diagnosis of Paget’s Disease of Bone Besides the typical clinical features of Paget’s disease (Table III), which are evident only in a relatively small subset of patients with the disease, the diagnosis is based on the typical radiographic abnormalities, linked to areas of increased bone turnover in the bone scan, on the characteristic histopathological changes and on the elevation

Table III. Typical Clinical Features of Paget’s Disease of Bone Bone pain Hypervascularity due to disease activity Bowing due to mechanical imcompetence Joint dysfunction due to secondary osteoarthritis Enlarging bones Danger of nerve compression symptoms, if the base of the skull or the vertebral column is affected Impaired hearing Predominantly due to sensineural hearing loss Rarely due to involvement of the internal auditory canal

of bone turnover markers, typically, the elevation of the formation marker total serum alkaline phosphatase (tAP). 1. Radiological Changes

Characteristic features include a localized enlargement of bone, cortical thickening, sclerotic changes, and osteolytic areas, like V-shaped lesions in long bones and osteoporosis circumscripta in the skull [89]. Radionuclide bone scanning is a very sensitive method to identify lesions with increased bone turnover, typical for Paget’s disease and should be performed at the time of diagnosis to screen clinically unaffected parts of skeleton for previously undetected manifestations [90]. 2. Role of Biochemical Markers of Bone Metabolism in Paget’s Disease of Bone

Paget’s disease of bone is usually characterized by excessive bone turnover and has therefore often been used as a model disease to evaluate markers of bone metabolism, reflecting either bone resorption of formation. The markers have been used with varying degree of success for both diagnosis and monitoring of disease activity. 3. Role of Biochemical Markers in the Diagnosis of Paget’s Disease

Of the biochemical markers of bone metabolism, tAP is usually, but not always, elevated. In approximately 15% of the symptomatic patients, tAP values are within the statistical limits of the normal range. There is a strong relation between the extent of the disease, measured by bone scan and the tAP activity [91]. It is important to note, however, that even a tAP activity within the upper region of the normal laboratory range may be a sign of disease activity in the individual patient and can be lowered by successful treatment [92]. In patients with concomitant liver or biliary disease an elevation of the bone-specific isoenzyme of the AP (bAP) may be masked, when total tAP is measured, which is one of the clinical situations, when the determination of the bAP is more

784 useful [93]. An increase in diagnostic sensitivity and specificity is the main criterion in the evaluations of other formation markers to establish superiority over tAP. Using samples from 51 patients with Paget’s disease, Alvarez et al. determined the levels of a variety of markers of bone formation: total alkaline phosphatase (tAP), bone-specific alkaline phosphatase (bAP), carboxyterminal propeptide of type I procollagen (PICP), aminoterminal propeptide of type I procollagen (PINP), osteocalcin, and a group of markers of bone resorption: hydroxyproline (OHP), pyridinoline (PYD), deoxypyridinoline (DPD), C-terminal telopeptide of type I collagen (CTx), N-terminal telopeptide of type I collagen (NTx), serumtartrate-resistant acid phosphatase (TRAP), and serumcarboxyterminal telopeptide of type I collagen (TCIP). All biochemical markers except osteocalcin correlated with the disease activity and the disease extent. Serum PINP, bAP, and total tAP showed the highest proportions of increased values among the bone formation markers (94%, 82%, and 76%, respectively). Among the bone resorption markers, urinary NTx showed the highest proportion of increased values in patients with Paget’s disease (96%), compared with PYD (69%), DPD (71%), CTx (65%), and OHP (64%). In patients with mild disease activity, serum PINP was the marker with the highest proportion of increased values (71%) [94]. When the activity of the disease is high (AP values > 500 U/L) most biochemical markers reveal values above the upper limit of normal [95]. However, when the disease activity is low, markers like PINP or BAP as formation markers and NTx as a resorption marker may improve the diagnostic sensitivity over the standard markers AP and DPD crosslinks. One study found elevated bone sialoprotein (BSP) levels in all patients, which was comparable to the performance of tAP and bAP in that study and superior to PYD, DPD, and osteocalcin [96]. BSP is a phosphorylated glycoprotein with a M(r) of 70–80 kDa that accounts for approximately 5–10% of the non-collagenous proteins of bone. Due to its relatively restricted distribution to mineralized tissues, BSP has the potential to serve as a marker of bone metabolism. Compared to healthy controls, serum BSP was significantly higher in patients with Paget’s disease and other bone disorders with high turnover like pHPT or metastatic bone disease [97]. Osteoprotegerin (OPG), but not RANKL values have been found elevated in patients with Paget’s disease, but no correlation was found between either bone markers or quantitative scintigraphic indices and serum levels of OPG, RANKL, IL-6, and RANKL:OPG ratios. The increase in OPG may therefore reflect a protective mechanism of the

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skeleton to compensate for increased bone resorption, rather than a marker of disease activity [98].

III. TREATMENT The increasing potency of newer bisphosphonates has shifted the emphasis of treatment from simply reducing bone turnover towards a complete suppression of the pathologically increased (i.e. excess) bone turnover for prolonged periods of time [99, 100]. The aim of this strategy is to relieve symptoms and to prevent long-term complications such as deformity, fracture, and neurological sequelae [101]. Up to know, it has not been proven, that long-term disease-suppressing therapy, will actually prevent those complications. A number of studies have generated circumstantial evidence, showing benefits for radiological or histolological endpoints. In studies with risedronate it has been shown that effective treatment will eventually lead to an improvement in radiological features especially for osteolytic lesions [102] and to normal lamellar bone formation rather than the woven bone characteristic of Paget’s disease [103]. In an observational study 41 patients with Paget’s disease of bone were followed over 12 years of treatment. In this study only four of the 12 patients, whose bone metabolism could be normalized during treatment, but 13 of 29 patients, where no complete normalization of the disease activity was possible, developed long-term complications [104]. How do we assess response to bisphosphonate treatment and the determinants of the duration of the response? The biochemical markers of bone turnover appear to be clearly superior to other established parameters (Tables IV and V). They are more objective and reproducible than history and clinical changes, they are readily available and more practical for clinical use than serial radiographs, bone scans, or bone biopsies [105]. How can we use the information given by the magnitude of change of the biochemical markers in a meaningful way? There are two different issues to address, the first being the assessment of response to treatment, the second the attempt to predict the duration of remission. Table IV.

Potential Parameters to Assess Response to Treatment in Paget’s Disease

Symptoms Findings at examination Biochemical markers of bone turnover Radiological features Histological features

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Chapter 47 Paget’s Disease of Bone Table V. Biochemical Markers in the Assessment of Paget’s Disease Bone Formation Formation markers of choice bAP Bone-specific alkaline phosphatase Serum PINP Aminoterminal propeptide of type I Serum procollagen tAP Total alkaline phosphatase Serum Rarely used, not superior to better validated markers PICP Carboxyterminal propeptide of type I Serum procollagen Well studied, does not correlate well with disease activity and response to therapy OC Osteocalcin Serum Bone resorption Current resorption marker of choice NTX Degradation fragments from the N-telopeptide Urine of alpha 1 chain of type I collagen Not widely validated, but potentially very useful αCTX Non-isomerized degradation fragments Urine from the C-telopeptide of alpha 1 chain of type I collagen Well studied, not superior to better validated markers CTX Degradation fragments from the Urine/serum C-telopeptide of alpha 1 chain of type I collagen DPD Deoxypyridinolin Urine/serum PYD Pyridinolin Urine tDPD Total DPD Urine fDPD Free DPD Urine Not widely validated BSP Bone sialoprotein Serum Well studied, do not correlate well with disease activity and response to therapy ICTP C-terminal telopeptide of type I collagen Serum TRAcP Tartrate-resistant acid phosphatase Serum

A. Assessing the Response to Treatment The determination of the total AP as mentioned is inexpensive, accurate and readiliy available. In daily practice the determination of the AP before and 3–6 months after initiation of any kind of antiresorptive treatment will provide sufficient information to assess the treatment response in an individual patient [106, 107]. During treatment monitoring there are a few specific situations, when other markers can become useful and superior to tAP. Two areas, which will be elaborated on in the next section, are the assessment of onset of response, where modern markers of bone resorption seem superior and the monitoring of patients with relatively low disease activity, where modern formation markers like bAP and PINP seem superior at least in some studies (Fig. 2). 1. Onset of Response

Antiresorptive agents lead to a significant reduction of biochemical markers of bone resorption, within days

Figure 2 Bone resorption and bone formation marker after bisphosphonate treatment of Paget’s disease. Serum total alkaline phosphatase (AP) activity and urinary hydroxyproline excretion (OHP) in precent of pretreatment values (mean ± S.E.M) of 22 patients with Paget’s disease treated with 9 × 20 mg pamidronate i.v. The first data point represents the values before treatment, the second on day 10 after initiation of treatment. Note the sharp decline of the resorption marker OHP within a few days of treatment, whereas the formation marker AP reaches its minimum after 3 months. (Modified after [108].)

after initiation of treatment [108].The reduction is maximal after 10 days [109]. Bone formation markers, such as AP take longer to reveal the treatment response, usually 3–4 weeks or longer, reaching a minimum activity approximately 3–6 months after treatment (Fig. 1) [108]. Whether this difference in onset of change can be utilized for a better prediction of treatment success has been addressed in a study where 21 patients were treated with daily infusions of two different aminobisphosphonates. Four different markers of bone resorption, OHP, total deoxypyridinoline crosslinks (tDPD), free deoxypyridinoline crosslinks (fDPD), and the crosslinked N-telopeptide of collagen type I (NTx) were compared. All markers showed a rapid decrease in urinary excretion within a few days. The reductions in urinary excretion of NTx and of the excess in OHP were nearly identical and more pronounced than the reduction of tDPD and fDPD. All patients who had a reduction in NTx > 75% after 10 days achieved a total AP within the normal range after 1 year. This preliminary study suggests that there may be a role in determining biochemical markers of bone resorption in the first 10 days after the initiation of the treatment to predict treatment response in an individual patient. This interesting concept needs to be further evaluated [109]. 2. Monitoring Treatment in Patients with Lower Disease Activity

Whether more modern markers of bone turnover are superior to the established markers in regular patient monitoring has been investigated in a number of trials

786 and is still an area of debate. Blumsohn compared the urinary measurements of OHP to total and free PYD (HPLC), immunoreactive free PYDs, immunoreactive total PYDs, and the N- and C-terminal telopeptides of type I collagen (NTX, CTX). Serum measurements included tartrate-resistant acid phosphatase (TRAP) and the C-terminal telopeptide of type I collagen (ICTP). The response was greatest for urinary telopeptides (NTX and CTX; followed by total PYD which showed a somewhat larger change than free PYD) [106]. ICTP and TRAP showed a minimal response to therapy and were, in line with earlier observations [110], considered unreliable indicators of changes in bone turnover in Paget’s disease. A more recent study, monitoring patients after zoledronate treatment found that tAP, bone AP, PINP, and uNTX all showed that >95% of subjects have abnormal values at baseline, reducing to 15–30% of their initial values following treatment. The poorer precision of uNTX reduced its slightly utility in this analysis, whereas already the baseline sensitivity of other markers (osteocalcin 68% of subjects abnormal, fDPD 22%, betaCTX 50%) was lower. The conclusion of these two studies was that the advantage of newer markers over tAP was at best marginal. Alvarez et al. compared marker changes after treatment with 400 mg/day of oral tiludronate for 3 months. In 31 patients that completed treatment, biochemical markers were measured at baseline and at 1 and 6 months after treatment ended, assessing serum values for tAP, bAP, PINP, CTX, and urine values for OHP, CTX, and NTX. The marker changes were compared to changes in quantitative bone scintigraphy. In addition to the relative change of the bone markers, the authors also determined a ratio for monitoring response to treatment by taking analytical and biological variation of the marker into account. Here ratio values of >1 indicated a significant decrease of the marker after therapy. Mean values of all markers of bone turnover decreased significantly after therapy. Serum bAP and PINP and urinary NTX showed the highest percentage reduction (between 58% and 68%). Furthermore, serum bAP and PINP showed the highest ratios for monitoring changes induced by treatment, followed by serum tAP and urinary NTX. Serum CTX and urinary CTX, as well as OHP, showed mean ratios for monitoring changes of <1, indicating a low sensitivity for monitoring treatment. The performance of tAP in this study was comparable to that of BAP and PINP in patients with higher disease activity and only inferior in patients with lower disease activity. Serum and urinary CTX did not perform as well as NTX in this analysis [111]. Initially the relatively poor performance of CTX was somewhat surprising, but can be explained physiologically.

ANDREAS GRAUER ET AL.

Most conventional CTX assays, both in serum and urine measure primarily βCTX In Paget’s disease of bone, the normal lamellar bone is replaced by a woven structure with an irregular arrangement of collagen fibers. The immunohistochemistry of normal and pagetic human bone sections showed a preferential distribution of αCTX within the woven structure, while lamellar bone was intensely stained with an anti-βCTX antibody, suggesting a lower degree of β-isomerization of type I collagen in the woven pagetic bone. This could explain why the urinary excretion of non-isomerized (α) fragments derived from degradation of type I collagen C-telopeptide (CTx) is markedly increased compared with β-isomerized CTx (+13-fold vs. +3.5-fold over controls) resulting in a urinary αCTx/βCTx ratio three-fold higher than in controls. In five pagetic patients in complete remission, as demonstrated by normal total alkaline phosphatase activity, the αCTx/βCTx ratio was normal [112]. Further studies looked into the racemization and isomerization of Asp(1211) in vitro and in vivo, which generates a mixture of four isomers: the native peptide form (alphaL), an isomerized form containing a beta-Asp bond (betaL), a racemized form containing a D-Asp residue (alphaD), and an isomerized/racemized form (betaD). The extent of racemization and isomerization were correlated with the age and turnover of collagen [113]. In Paget’s disease and prostate cancer-induced bone metastases, conditions characterized by focal increased bone turnover, alpha L CTX levels were more elevated than those of age-related CTX forms, resulting in increased ratios between native and age-modified CTX. For example, the ratio alpha L/alpha D was increased seven-fold in Paget’s disease (P < 0.001) and two-fold in prostate cancer-induced bone metastases (P < 0.002). This suggests that in conditions with a localized alteration in bone turnover the ratio between alpha L CTX and the age-modified forms is significantly elevated [114]. The use of a specific alphaalpha CTX assay was shown to improve the ability of αCTX measurement to assess disease activity and monitor treatment efficacy of conventional CTX assays, which measure primarily βCTX. In a recent study, urinary ααCTX showed a marked reduction (−82% and −77% at 1 and 6 months of treatment, respectively) in response to antiresorptive therapy in patients with Paget’s disease. The alpha-alpha CTX marker also provided a high correlation (r = 0.89) to disease activity as assessed by scintigraphic activity index [115]. 3. Serum or Urine Measurements?

Whether serum or urine measurements are preferred is more a question of clinical convenience. Serum markers have the advantage that there is no collection period and

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that a sample can be obtained when the patient visits the clinic. The downside here is that in particular for serum CTX, sampling has to be performed both fasting and in the morning, as there is a large variation due to fasting status and a marked diurnal variation. Urine markers are usually obtained from the second morning void– which again sometimes poses logistical problems for the patients. One major advantage of the newer markers of bone resorption over OHP is that they are largely independent of dietary influences, and do not require compliance to a proline-free diet during and prior to the sampling period. 4. Other Markers Investigated for the Monitoring of Paget’s Disease

a. Bone Sialoprotein (BSP) Potentially suitable for diagnosis, BSP was found significantly reduced early as 24 h after treatment with 2 mg ibandronate compared to pretreatment levels, reaching a minimum after 30 days. This time course suggests that BSP primarily reflects bone resorption [116]. b. ICTP and TRAP As mentioned above the bone resorption markers ICTP [110] and TRAP measured in serum are unreliable indicators of changes in bone turnover in Paget’s disease [106]. c. Osteocalcin Several studies show that there is no significant relation between osteocalcin levels, disease activity in Paget’s disease, or any of the established markers of bone resorption or formation [117–119].

B. Clinical Decision Making in Monitoring Treatment of Paget’s Disease 1. Evaluation of Response

Three months after initiation of a bisphosphonate treatment, the curves of resorption and formation markers synchronize, and in particular in patients with moderate to high disease activity, total AP will be sufficient to monitor treatment. A commonly used method is to express the decrease in AP as a percentage of pretreatment AP (percent decrease in AP), or the percent decrease of the pretreatment AP in relation to the upper limit of the reference range of a normal population (percent decrease in excess AP). Although useful, this approach is limited by the inverse relationship between these percentage changes and the pretreatment AP. The greatest percentage change is usually seen in those with the lowest pretreatment AP. To achieve a 50% reduction in AP activity could either take a reduction from 800 to 400 IU or from

300 to 150 IU [99]. Moreover, the post-treatment AP is also affected by the pretreatment AP. Patients with active disease are less likely to be suppressed into the normal range [100, 120]. Therefore, unless matched for pretreatment AP, these methods are of limited usefulness in the comparison between patients and studies of the efficacy of various bisphosphonates. A possible solution may be to assess the rate of AP decrease. It has been shown to be exponential and be easily expressed as a half life on a log-linear plot [121]. Using this method, the half life of AP has been shown to be independent of pretreatment AP [105] which would suggest the AP half life is a superior expression of response compared with percentage decrease. There is also a marked interindividual variation between patients with similar pretreatment AP, administered the same dose of intravenous bisphosphonate [99]. Also here, the half life of the AP seems to give an indication of bone cell sensitivity within an individual. The AP half life may therefore be a good predictor of fair and poor response to bisphosphonate treatment and provide an opportunity to modify therapy in the hope of a better outcome [105]. This interesting concept needs to be evaluated in prospective clinical trials. The ideal marker would eliminate all those considerations, by showing normalization only if disease is suppressed and elevated values in case of residual disease activity, allowing a clear distinction between a sufficient and a not-yet-sufficient treatment.

C. Predicting the Duration of Remission The goal of a bisphosphonate treatment in Paget’s disease of bone is to achieve prolonged suppression of bone turnover and reduced complications. Patients with a short AP half life after bisphosphonate treatment tend to have a longer duration of remission, possibly indicating their increased sensitivity to the treatment [105]. The determinants of the duration of the response have been examined and shown to be dependent on both pre- and post-treatment AP. The greater the AP value before therapy and the lesser the degree of AP suppression after treatment, the shorter the period during which bone turnover will be suppressed. Interestingly, this seems also to apply when the post-treatment AP is within the population reference range [100, 120]. This highlights the difficulty of applying population ranges to individuals. A post-treatment AP of 150 U/L maybe within the population range of 50–170 U/L but may still be clearly elevated if the individual’s natural AP is 100 U/L. The difficulty of converting this finding into a treatment concept is that the individual’s natural AP is usually not known. The observation suggests, however, that the AP should not only

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be “normalized”, but suppressed well within the population reference range to achieve a prolonged remission [122]. Again, this is a hypothesis, which needs to be evaluated in prospective studies. Preliminary data suggest that both bAP and PINP may be more suitable to discriminate between patients whose pagetic activity is truly normalized and patients, who just have reached the normative range of the tAP.

References 1. Delmas, P. D., and Meunier, P. J. (1997). The management of Paget’s disease of bone. N. Engl. J. Med. 336, 558–566. 2. Schmorl, G. (1932). Über Osteitis deformans Paget. Virchow’s Arch. Path. Anat. 283, 694–751. 3. Pygott, F. (1957). Paget’s disease of bone; the radiological incidence. Lancet 272, 1170–1171. 4. Detheridge, F. M., Guyer, P. B., and Barker, D. J. (1982). European distribution of Paget’s disease of bone. Br. Med. J. (Clin. Res. Ed.) 285, 1005–1008. 5. Siris, E. S. (1996). Seeking the elusive etiology of Paget disease: a progress report [editorial; comment]. J. Bone Miner. Res. 11, 1599–1601. 6. Cundy, H. R., Gamble, G., Wattie, D., Rutland, M., and Cundy, T. (2004). Paget’s disease of bone in New Zealand: continued decline in disease severity. Calcif. Tissue Int. 75, 358–364. 7. van Staa, T. P., Selby, P., Leufkens, H. G., Lyles, K., Sprafka, J. M., and Cooper, C. (2002). Incidence and natural history of Paget’s disease of bone in England and Wales. J. Bone Miner. Res. 17, 465–471. 8. Roodman, G. D., and Windle, J. J. (2005). Paget disease of bone. J. Clin. Invest. 115, 200–208. 9. Montagu, M. F. A. (1949). Paget’s disease (osteitis deformans) and hereditary. Am. J. Hum. Genet. 1, 94–95. 10. Siris, E. S., Ottman, R., Flaster, E., and Kelsey, J. L. (1991). Familial aggregation of Paget’s disease of bone. J. Bone Miner. Res. 6, 495–500. 11. Sofaer, J. A., Holloway, S. M., and Emery, A. E. (1983). A family study of Paget’s disease of bone. J. Epidemiol. Comm. Health 37, 226–231. 12. Morales-Piga, A. A., Rey-Rey, J. S., Corres-Gonzalez, J., GarciaSagredo, J. M., and Lopez-Abente, G. (1995). Frequency and characteristics of familial aggregation of Paget’s disease of bone. J. Bone Miner. Res. 10, 663–670. 13. Haslam, S. I., Van Hul, W., Morales-Piga, A., Balemans, W., San Millan, J. L., Nakatsuka, K., Willems, P., Haites, N. E., and Ralston, S. H. (1998). Paget’s disease of bone: evidence for a susceptibility locus on chromosome 18q and for genetic heterogeneity. J. Bone Miner. Res. 13, 911–917. 14. Hocking, L., Slee, F., Cundy, T., Nicholson, G., Van Hul, W., and Ralston, S. H. (2000). Familial Paget’s disease of Bone: Patterns of inheritance and frequency of linkage to chromosome 18q. Bone 26, 577–580. 15. Cody, J. D., Singer, F. R., Roodman, G. D., Otterund, B., Lewis, T. B., Leppert, M., and Leach, R. J. (1997). Genetic linkage of Paget disease of the bone to chromosome 18q. Am. J. Hum. Genet. 61, 1117–1122. 16. Laurin, N., Brown, J. P., Lemainque, A., Duchesne, A., Huot, D., Lacourciere, Y., Drapeau, G., Verreault, J., Raymond, V., Morissette, J. (2001). Paget disease of bone: mapping of two loci at 5q35-qter and 5q31. Am. J. Hum. Genet. 69, 528–543. 17. Osterberg, P. H., Wallace, R. G., Adams, D. A., Crone, R. S., Dickson, G. R., Kanis, J. A., Mollan, R. A., Nevin, N. C., Sloan, J., and Toner, P. G. (1988). Familial expansile osteolysis. A new dysplasia. J. Bone. Joint Surg. Br. 70, 255–260.

18. Whyte, M. P., Mills, B. G., Reinus, W. R., Podgornik, M. N., Roodman, G. D., Gannon, F. H., Eddy, M. C., McAlister, W. H. (2000). Expansile skeletal hyperphosphatasia: a new familial metabolic bone disease. JBMR 15, 2330–2344. 19. Nakatsuka, K., Nishizawa, Y., and Ralston, S. H. (2003). Phenotypic characterization of early onset Paget’s disease of bone caused by a 27-bp duplication in the TNFRSF11A gene. J. Bone Miner. Res. 18, 1381–1385. 20. Whyte, M. P., Obrecht, S. E., Finnegan, P. M., Jones, J. L., Podgornik, M. N., McAlister, W. H., Mumm, S. (2002). Osteoprotegerin deficiency and juvenile Paget’s disease. N. Engl. J. Med. 347, 175–184. 21. Kovach, M. J., Waggoner, B., Leal, S. M., Gelber, D., Khardori, R., Levenstien, M. A., Shanks, C. A., Gregg, G., Al Lozi, M. T., Miller, T., Rakowicz, W., Lopate, G., Florence, J., Glosser, G., Simmons, Z., Morris, J. C., Whyte, M. P., Pestronk, A., and Kimonis, V. E. (2001). Clinical delineation and localization to chromosome 9p13.3-p12 of a unique dominant disorder in four families: hereditary inclusion body myopathy, Paget disease of bone, and frontotemporal dementia. Mol. Genet. Metab. 74, 458–475. 22. Hughes, A. E., Shearman, A. M., Weber, J. L., Barr, R. J., Wallace, R. G., Osterberg, P. H., Nevin, N. C., Mollan, R. A. (1994). Genetic linkage of familial expansile osteolysis to chromosome 18q. Hum. Mol. Genet. 3, 359–361. 23. Hocking, L. J., Herbert, C. A., Nicholls, R. K., Williams, F., Bennett, S. T., Cundy, T. Nicholson, G. C., Wuyts, W., Van Hul, W., and Ralston, S. H. (2001). Genomewide search in familial paget disease of bone shows evidence of genetic heterogeneity with candidate loci on chromosomes 2q36, 10p13, and 5q35. Am. J. Hum. Genet. 69, 1055–1061. 24. Good, D. A., Busfield, F., Fletcher, B. H., Duffy, D. L., Kesting, J. B., Andersen, J., Shaw, J. T. (2001). Linkage of Paget Disease of bone to a novel region on human chromosome 18q23. Am. J. Hum. Genet. 70, 517–525. 25. Tilyard, M. W., Gardner, R. J., Milligan, L., Cleary, T. A., and Stewart, R. D. (1982). A probable linkage between familial Paget’s disease and the HLA loci. Aus. N. Z. J. Med. 12, 498–500. 26. Cundy, T., Hegde, M., Naot, D., Chong, B., King, A., Wallace, R. Mulley, J., Love, D. R., Seidel, J., Fawkner, M., Banovic, T., Callon, K. E., Grey, A. B., Reid, I. R., Middleton-Hardie, C. A., and Comish, J. (2002). A mutation in the gene TNFRSF11B encoding osteoprotegerin causes an idiopathic hyperphosphatasia phenotype. Hum. Mol. Genet. 11, 2119–2127. 27. Hughes, A. E., Ralston, S. H., Marken, J., Bell, C., MacPherson, H., Wallace, R. G., Van Hul, W., Whyte, M. P., Nakatsuka, K., Hovy, L., and Anderson, D. M. (2000). Mutations in TNFRSF11A, affecting the signal peptide of RANK, cause familial expansile osteolysis. Nat. Genet. 24, 45–48. 28. Nakatsuka, K., Nishizawa, Y., and Ralston, S. H. (2003). Phenotypic characterization of early onset Paget’s disease of bone caused by a 27-bp duplication in the TNFRSF11A gene. J. Bone Miner. Res. 18, 1381–1385. 29. Whyte, M. P., and Hughes, A. E. (2002). Expansile skeletal hyperphosphatasia is caused by a 15-base pair tandem duplication in TNFRSF11A encoding RANK and is allelic to familial expansile osteolysis. J. Bone Miner. Res. 17, 26–29. 30. Johnson-Pais, T. L., Singer, F. R., Bone, H. G., McMurray, C. T., Hansen, M. F., and Leach, R. J. (2003).Identification of a novel tandem duplication in exon 1 of the TNFRSF11A gene in two unrelated patients with familial expansile osteolysis. J. Bone Miner. Res. 18, 376–380. 31. Palenzuela, L., Vives-Bauza, C., Fernandez-Cadenas, I., Meseguer, A., Font, N., Sarret, E. Schwartz, S., and Andreu, A. L. (2002). Familial expansile osteolysis in a large Spanish kindred resulting from an insertion mutation in the TNFRSF11A gene. J. Med. Genet. 39, E67. 32. Whyte, M. P., Obrecht, S. E., Finnegan, P. M., Jones, J. L., Podgornik, M. N., McAlister, W. H., and Mumm, S. (2002). Osteoprotegerin deficiency and juvenile Paget’s disease. N. Engl. J. Med. 347, 175–184.

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Chapter 47 Paget’s Disease of Bone 33. Watts, G. D., Wymer, J., Kovach, M. J., Mehta, S. G., Mumm, S., Darvish, D., Pestronk, A., Whyte, M. P., and Kimonis, V. E. (2004). Inclusion body myopathy associated with Paget disease of bone and frontotemporal dementia is caused by mutant valosin-containing protein. Nat. Genet. 36, 377–381. 34. Schroder, R., Watts, G. D., Mehta, S. G., Evert, B. O., Broich, P., Fliessbach, K., Frassica, F. J., Streenten, E. A., Levine, M. A., Fraser, C. M., Adams, M. D., Broder, S., Venter, J. C., Kinzler, K. W., Vogelstein, B., and Ralston, S. H. (2005). Mutant valosin-containing protein causes a novel type of frontotemporal dementia. Ann. Neurol. 57, 457–461. 35. Dai, R. M., Chen, E., Longo, D. L., Gorbea, C. M., and Li, C. C. (1998). Involvement of valosin-containing protein, an ATPase Co-purified with IkappaBalpha and 26 S proteasome, in ubiquitin-proteasome-mediated degradation of IkappaBalpha. J. Biol. Chem. 273, 3562–3573. 36. Sparks, A. B., Peterson, S. N., Bell, C., Loftus, B. J., Hocking, L., Cahill, D. P., Frassica, F. J., Streenten, E. A., Levine, M. A., Fraser, C. M., Adams, M. D., Broder, S., Venter, J. C., Kinzler, K. W., Vogelstein, B., Ralston, S. H. (2001). Mutation screening of the TNFRSF11A gene encoding receptor activator of NF kappa B (RANK) in familial and sporadic Paget’s disease of bone and osteosarcoma. Calcif. Tissue Int. 68, 151–155. 37. Wuyts, W., Van Wesenbeeck, L., Morales-Piga, A., Ralston, S., Hocking, L., Vanhoenacker, F., Westhovens, R., Verbruggen, L., Anderson, D., Hughes, A., and Van Hul, W. (2001). Evaluation of the role of RANK and OPG genes in Paget’s disease of bone. Bone 28, 104–107. 38. Eekhoff, E. W., Karperien, M., Houtsma, D., Zwinderman, A. H., Dragoiescu, C., Kneppers, A. L., and Papapoulos, S. E. (2004). Familial Paget’s disease in The Netherlands: occurrence, identification of new mutations in the sequestosome 1 gene, and their clinical associations. Arthritis. Rheum. 50, 1650–1654. 39. Sparks, A. B., Peterson, S. N., Bell, C., Loftus, B. J., Hocking, L., Cahill, D. P., Frassica, F. J., Streenten, E. A., Levine, M. A., Fraser, C. M., Adams, M. D., Broder, S., Venter, J. C., Kinzler, K. W., Vogelstein, B., Ralston, S. H. (2001). Mutation screening of the TNFRSF11A gene encoding receptor activator of NF kappa B (RANK) in familial and sporadic Paget’s disease of bone and osteosarcoma. Calcif. Tissue Int. 68, 151–155. 40. Wuyts, W., Van Wesenbeeck, L., Morales-Piga, A., Ralston, S., Hocking, L., Vanhoenacker, F., Westhovens, R., Verbruggen, L., Anderson, D., Hughes, A., and Van Hul, W. (2001). Evaluation of the role of RANK and OPG genes in Paget’s disease of bone. Bone 28, 104–107. 41. Daroszewska, A., Hocking, L. J., McGuigan, F. E. A., Langdahl, B. L., Stone, M. D., Cundy, T., Nicholson, G. C., Fraser, W. D., Ralston, S. H., (2004). Susceptibility to Paget’s disease of bone is influenced by a common polymorphic variant of Osteoprotegerin. J. Bone Min. Res. 19, (in press). 42. Laurin, N., Brown, J. P., Morissette, J., and Raymond, V. (2002). Recurrent mutation of the gene encoding sequestosome 1 (SQSTM1/p62) in Paget Disease of bone. Am. J. Hum. Genet. 70, 1582–1588. 43. Hocking, L. J., Lucas, G. J. A., Daroszewska, A., Mangion, J., Olavesen, M., Nicholson, G. C., Ward, L., Bennett, S. T., Wuyts, W., Van Hul, W., and Ralston, S. H. (2002). Domain specific mutations in Sequestosome 1 (SQSTM1) cause familial and sporadic Paget’s disease. Hum. Mol. Gen. 11, 2735–2739. 44. Hocking, L. J., Lucas, G. J. A., Daroszewska, A., Cundy, T., Nicholson, G. C., Donath, J., Walsh, J. P., Finlayson, C., Cavey, J. R., Ciani, B., Sheppard, P. W., Searle, M. S., Layfield, R., and Ralston, S. H. (2004). UBA domain mutations of SQSTM1 in Paget’s disease of bone: genotype phenotype correlation, functional analysis and structural consequences. J. Bone Miner. Res. 19, 1122–1127. 45. Johnson-Pais, T. L., Wisdom, J. H., Weldon, K. S., Cody, J. D., Hansen, M. F., Singer, F. R., and Leach, R. J. (2003). Three novel

46.

47.

48.

49.

50.

51.

52. 53.

54.

55.

56.

57.

58.

59.

60.

61. 62.

63.

64.

mutations in SQSTM1 identified in familial Paget’s disease of bone. J. Bone Miner. Res. 18, 1748–1753. Falchetti, A., Di Stefano, M., Marini, F., Del Monte, F., Mavilia, C., Strigoli, D., De Feo, M. L., Isaia, G., Masi, L., Amedei, A., Cioppi, F., Ghinoi, V., Maddali Bongi, S., Di Fede, G., Strerrazza, C., Rini, G. B., Melchiorre, D., Matucci-Cerenic, M., and Brandi, M. L. (2004). Two novel mutations at exon 8 of Sequestosome 1 gene (SQSTM1) in an Italian series of patients affected by Paget’s disease of bone (PDB). J. Bone Miner. Res. 19, (in press). Eekhoff, E. W. M., Karperien, M., Houtsma, D., Zwinderman, A. H., Dragoiescu, C., Kneppers, A. L. J., and Papaoulos, S. E. (2004). Familial Paget’s Disease in the Netherlands. Arth. Rheum. 50(5), 1650–1654. Moscat, J., and Diaz-Meco, M. T. (2002). The atypical PKC scaffold protein P62 is a novel target for anti-inflammatory and anti-cancer therapies. Adv. Enzyme. Regul. 42, 173–179. Ciani, B., Layfield, R., Cavey, J. R., Sheppard, P. W., and Searle, M. S. (2003). Structure of the UBA domain of p62 (SQSTM1) and implications for mutations which cause Paget’s disease of bone. J. Biol. Chem. 278, 37409–37412. Duran, A., Serrano, M., Leitges, M., Flores, J. M., Picard, S., Brown, J. P., Moscat, J., and Diaz-Meco, M. T. (2004). The atypical PKCinteracting protein p62 is an important mediator of RANK-activated osteoclastogenesis. Dev. Cell. 6, 303–309. Rebel, A., Malkani, K., Basle, M., and Bregeon, C. H. (1974). Particularites ultrastructurales des osteoclasts de la maladie de Paget. Rev. Rhum. Mal. Osteoartic. 41, 767–771. Mills, B. G., and Singer, F. R. (1976). Nuclear inclusions in Paget’s disease of bone. Science 194, 201–202. Mills, B. G., Yabe, H., and Singer, F. R. (1988). Osteoclasts in human osteopetrosis contain viral-nucleocapsid-like nuclear inclusions. J. Bone Miner. Res. 3, 101–106. Beneton, M. N. C., Harris, S., and Kanis, J. A. (1987). Paramyxoviruslike inclusions in two cases of pycnodysostosis. Metab. Bone Dis. Rel. Res. 8, 211–217. Bianco, P., Silvestrini, G., Ballanti, P., and Bonucci, E. (1992). Paramyxovirus-like nuclear inclusions identical to those of Paget’s disease of bone detected in giant cells of primary oxalosis. Virch. Archiv. Patho. Anat. Histopatho. 421, 427–433. McKenna, M. J., Mills, B. G., Galey, F. R., and Linthicum, F. H. Jr. (1986). Filamentous structures morphologically similar to viral nucleocapsids in otosclerotic lesions in two patients. Am. J. Otol. 7, 25–28. Mills, B. G., Frausto, A., Singer, F. R., Ohsaki, Y., Demulder, A., and Roodman, G. D. (1994). Multinucleated cells formed in vitro from Paget’s bone marrow express viral antigens. Bone 15, 443–448. Rebel, A., Basle, M., Pouplard, A., Malkani, K., Filmon, R., and Lepatezour, A. (1980). Bone tissue in Paget’s disease of bone. Ultrastructure and immunocytology. Arth. Rheum. 23, 1104–1114. Mills, B. G., Singer, F. R., Weiner, L. P., and Holst, P. A. (1981). Immunohistological demonstration of respiratory syncytial virus antigens in Paget’s disease of bone. Proc. Natl Acad. Sci. USA 78, 1209–1212. Mills, B. G., Singer, F. R., Weiner, L. P., Suffin, S. C., Stabile, E., and Holst, P. (1984). Evidence for both respiratory syncytial virus and measles virus antigens in the osteoclasts of patients with Paget’s disease of bone. Clin. Orthop. Rel. Res. 183, 303–311. O’Driscoll, J. B., and Anderson, D. C. (1985). Past pets and Paget’s disease. Lancet 2, 919–921. Khan, S. A., Brennan, P., Newman, J., Gray, R. E., McCloskey, E. V., and Kanis, J. A. (1996). Paget’s disease of bone and unvaccinated dogs. Bone 19, 47–50. Siris, E. S., Kelsey, J. L., Flaster, E., and Parker, S. (1990). Paget’s disease of bone and previous pet ownership in the United States: dogs exonerated. Int. J. Epidemiol. 19, 455–458. Gordon, M. T., Anderson, D. C., and Sharpe, P. T. (1991). Canine distemper virus localised in bone cells of patients with Paget’s disease. Bone 12, 195–201.

790 65. Mee, A. P., Dixon, J. A., Hoyland, J. A., Davies, M., Selby, P. L., and Mawer, E. B. (1998). Detection of Canine Distemper Virus in 100% of Paget’s disease samples by in situ-reverse transcriptase polymerase chain reaction. Bone 23, 171–175. 66. Gordon, M. T., Mee, A. P., Anderson, D. C., and Sharpe, P. T. (1992). Canine distemper transcripts sequenced from pagetic bone. Bone Min. 19, 159–174. 67. Basle, M. F., Fournier, J. G., Rozenblatt, S., Rebel, A., and Bouteille, M. (1986). Measles virus RNA detected in Paget’s disease bone tissue by in situ hybridization. J. Gen. Virol. 67, 907–913. 68. Reddy, S. V., Menaa, C., Singer, F. R., Cundy, T., Cornish, J., Whyte, M. P., Roodman, G. D. (1999). Measles virus nucleocapsid transcript expression is not restricted to the osteoclast lineage in patients with Paget’s disease of bone. Exp. Hematol. 27, 1528–1532. 69. Ralston, S. H., DiGiovine, F. S., Gallacher, S. J., Boyle, I. T., and Duff, G. W. (1991). Failure to detect paramyxovirus sequences in Paget’s disease of bone using the polymerase chain reaction. J. Bone Miner. Res. 6, 1243–1248. 70. Birch, M. A., Taylor, W., Fraser, W. D., Ralston, S. H., Hart, C. A., and Gallagher, J. A. (1994). Absence of paramyxovirus RNA in cultures of pagetic bone cells and in pagetic bone. J. Bone Miner. Res. 9, 11–16. 71. Ooi, C. G., Walsh, C. A., Gallagher, J. A., and Fraser, W. D. (2000). Absence of measles virus and canine distemper virus transcripts in long-term bone marrow cultures from patients with Paget’s disease of bone. Bone 27, 417–421. 72. Helfrich, M. H., Hobson, R. P., Grabowski, P. S., Zurbriggen, A., Cosby, S. L., Dickson, G. R., Fraser, W. D., Ooi, C. G., Selby, P. L., Crisp, A. J., Wallace, R. G., Kahn, S., and Ralston, S. H. (2000). A negative search for a paramyxoviral etiology of Paget’s disease of bone: molecular, immunological, and ultrastructural studies in UK patients. J. Bone Miner. Res. 15, 2315–2329. 73. Nuovo, M. A., Nuovo, G. J., MacConnell, P., Forde, A., and Steiner, G. C. (1992). In situ analysis of Paget’s disease of bone for measles-specific PCR-amplified cDNA. Diag. Mol. Pathol. 1, 256–265. 74. Friedrichs, W. E., Reddy, S. V., Bruder, J. M., Cundy, T., Cornish, J., Singer, F. R., and Roodman, G. D. (2002). Sequence analysis of measles virus nucleocapsid transcripts in patients with Paget’s disease. J. Bone Miner. Res. 17, 145–151. 75. Friedrichs, W. E., Reddy, S. V., Singer, F. R., and Roodman, G. D. (2002). The pro and con of measles virus in Paget’s Disease: Pro. J. Bone Miner. Res. 17, 2293. 76. Rima, B. K., Gassen, U., Helfrich, M. H., and Ralston, S. H. (2002). The pro and con of measles virus in Paget’s disease: con. J. Bone Miner. Res. 17, 2290–2292. 77. Mee, A. P., May, C., Bennett, D., and Sharpe, P. T. (1995). Generation of multinucleated osteoclast-like cells from canine bone marrow: effects of canine distemper virus. Bone 17, 47–55. 78. Reddy, S. V., Kurihara, N., Menaa, C., Landucci, G., Forthal, D., Koop, B. A., Windle, J. J., and Roodman, G. D. (2001). Osteoclasts formed by measles virus-infected osteoclast precursors from hCD46 transgenic mice express characteristics of pagetic osteoclasts. Endocrinology 142, 2898–2905. 79. Kurihara, N., Reddy, S. V., Menaa, C., Anderson, D., and Roodman, G. D. (2000). Osteoclasts expressing the measles virus nucleocapsid gene display a pagetic phenotype. J. Clin. Invest. 105, 607–614. 80. Demulder, A., Takahashi, S., Singer, F. R., Hosking, D. J., and Roodman, G. D. (1993). Abnormalities in osteoclast precursors and marrow accessory cells in Paget’s disease. Endocrinology 133, 1978–1982. 81. Kukita, A., Chenu, C., McManus, L. M., Mundy, G. R., and Roodman, G. D. (1990). Atypical multinucleated cells form in longterm marrow cultures from patients with Paget’s disease. J. Clin. Invest. 85, 1280–1286. 82. Menaa, C., Reddy, S. V., Kurihara, N., Maeda, H., Anderson, D., Cundy, T., Comish, J., Singer, F. R., Bruder, J. M., and Roodman, G. D.

ANDREAS GRAUER ET AL.

83.

84.

85.

86.

87.

88.

89.

90.

91.

92. 93.

94.

95.

96.

97.

98.

(2000). Enhanced RANK ligand expression and responsivity of bone marrow cells in Paget’s disease of bone. J. Clin. Invest. 105, 1833–1838. Neale, S. D., Smith, R., Wass, J. A., and Athanasou, N. A. (2000). Osteoclast differentiation from circulating mononuclear precursors in Paget’s disease is hypersensitive to 1,25-dihydroxyvitamin D(3) and RANKL. Bone 27, 409–416. Roodman, G. D., Kurihara, N., Ohsaki, Y., Kukita, A., Hosking, D., Demulder, A., Smith, J. F., Singer, F. R. (1992). Interleukin 6. A potential autocrine/paracrine factor in Paget’s disease of bone. J. Clin. Invest. 89, 46–52. Natale, V. M., Filho, W. J., and Duarte, A. J. (1997). Cellular immunity aspects in elderly subjects with Paget’s disease of bone. Calcif. Tissue Int. 60, 410–414. Hoyland, J. A., and Sharpe, P. T. (1994). Up-regulation of c-fos protooncogene expression in Pagetic Osteoclasts. J. Bone Miner. Res. 8, 1191–1195. Ishizuka, S., Kurihara, N., Miura, D., Takenouchi, K., Cornish, J., Cundy, T., Reddy, S. V., and Roodman, G. D. (2004). Vitamin D antagonist, TEI-9647, inhibits osteoclast formation induced by 1 alpha, 25-dihydroxyvitamin D-3 from pagetic bone marrow cells. J. Steroid Biochem. Molec. Biol. 89–90, 331–334. Ishizuka, S., Kurihara, N., Reddy, S. V., Cornish, J., Cundy, T., and Roodman, G. D. (2005). (23S)-25-Dehydro-1{alpha}-hydroxyvitamin D3-26, 23-lactone, a vitamin D receptor antagonist that inhibits osteoclast formation and bone resorption in bone marrow cultures from patients with Paget’s disease. Endocrinology 146, 2023–2030. Grauer, A., and Wüster, C. (1996). Morbus Paget. In: “Klinische Endokrinologie: Urban & Schwarzenberg”. Allolio, B., Schulte, H. M. pp. 336–341. München, Germany. Grauer, A. (1997). Seltene Osteopathien: Morbus Paget und Osteogenesis imperfecta. Syllabus, IV. Intensivkurs für Klinische Endokrinologie. Neu-Isenburg: Bundesdruckerei, 67–76. Meunier, P. J., Salson, C., Mathieu, L., Chapuy, M. C., Delmas, P., Alexandre, C., and Charhon, S. (1987). Skeletal distribution and biochemical parameters of Paget’s disease. Clin. Orthop. 217, 37–44. Grauer, A. (1998). Therapie des Morbus Paget: Indikationen und Strategien. Osteologie 7(Suppl. 1), 85–86. Garnero, P., and Delmas, P. D. (1993). Assessment of the serum levels of bone alkaline phosphatase with a new immunoradiometric assay in patients with metabolic bone disease. J. Clin. Endocrinol. Metab. 77, 1046–1053. Alvarez, L., Peris, P., Pons, F., Guanabens, N., Herranz, R., Monegal, A., Bedini, J. L., Deulofeu, R., Martinez de Osaba, M. J., Munoz-Gomez, J., and Ballesta, A. M. (1997). Relationship between biochemical markers of bone turnover and bone scintigraphic indices in assessment of Paget’s disease activity. Arthritis. Rheum. 40, 461–468. Alvarez, L., Guanabens, N., Peris, P., Monegal, A., Bedini, J. L., Deulofeu, R., Martinez de Osaba, M. J., Munoz-Gomez, J., RiveraFillat, F., and Ballesta, A. M. (1995). Discriminative value of biochemical markers of bone turnover in assessing the activity of Paget’s disease. J. Bone Miner. Res. 10, 458–465. Woitge, H. W., Oberwittler, H., Heichel, S., Grauer, A., Ziegler, R., and Seibel, M. J. (2000). Short- and long-term effects of ibandronate treatment on bone turnover in Paget disease of bone. Clin. Chem. 46, 684–690. Seibel, M. J., Woitge, H. W., Pecherstorfer, M., Karmatschek, M., Horn, E., Ludwig, H., Ambruster, F. P., and Ziegler, R. (1996). Serum immunoreactive bone sialoprotein as a new marker of bone turnover in metabolic and malignant bone disease. J. Clin. Endocrinol. Metab. 81, 3289–3294. Alvarez, L., Peris, P., Guanabens, N., Vidal, S., Ros, I., Pons, F., Filella, X., Monegal, A., Munoz-Gomez, J., Ballesta, A. M. (2003). Serum osteoprotegerin and its ligand in Paget’s disease of bone: relationship to disease activity and effect of treatment with bisphosphonates. Arthritis Rheum. 48, 824–828.

Chapter 47 Paget’s Disease of Bone 99. Patel, S., Coupland, C. A., Stone, M. D., and Hosking, D. J. (1995). Comparison of methods of assessing response of Paget’s disease to bisphosphonate therapy. Bone 16, 193–197. 100. Patel, S., Stone, M. D., Coupland, C., and Hosking, D. J. (1993). Determinants of remission of Paget’s disease of bone. J. Bone Miner. Res. 8, 1467–1473. 101. Kanis, J. A., and Gray, R. E. (1987). Long-term follow-up observations on treatment in Paget’s disease of bone. Clin. Orthop. 217, 99–125. 102. Brown, J. P., Chines, A. A., Myers, W. R., Eusebio, R. A., RitterHrncirik, C., and Hayes, C. W. (2000). Improvement of pagetic bone lesions with risedronate treatment: a radiologic study. Bone 26, 263–267. 103. Meunier, P. J. (1994). Bone histomorphometry and skeletal distribution of Paget’s disease of bone. Semin. Arthritis Rheum. 23, 219–221. 104. Meunier, P. J., and Vignot, E. (1995). Therapeutic strategy in Paget’s disease of bone. Bone 17(5 Suppl), 489S–491S. 105. Patel, S. (1995). Treatment response in Paget’s disease. Ann. Rheum. Dis. 54, 783–784. 106. Blumsohn, A., Naylor, K. E., Assiri, A. M., and Eastell, R. (1995). Different responses of biochemical markers of bone resorption to bisphosphonate therapy in Paget disease. Clin. Chem. 41, 1592–1598. 107. Reid, I. R., Davidson, J. S., Wattie, D., Wu, F., Lucas, J., Gamble, G. D., Filella, X., Monegal, A., Munoz-Gomez, J., and Ballesta, A. M. (2004). Comparative responses of bone turnover markers to bisphosphonate therapy in Paget’s disease of bone. Bone 35, 224–230. 108. Grauer, A., Klar, B., Scharla, S. H., and Ziegler, R. (1994). Long-term efficacy of intravenous pamidronate in Paget’s disease of bone. Semin. Arthritis Rheum. 23, 283–284. 109. Papapoulos, S. E., and Frolich, M. (1996). Prediction of the outcome of treatment of Paget’s disease of bone with bisphosphonates from short-term changes in the rate of bone resorption. J. Clin. Endocrinol. Metab. 81, 3993–3997. 110. Filipponi, P., Pedetti, M., Beghe, F., Giovagnini, B., Miam, M., and Cristallini, S. (1994). Effects of two different bisphosphonates on Paget’s disease of bone: ICTP assessed. Bone 15, 261–267. 111. Alvarez, L., Guanabens, N., Peris, P., Vidal, S., Ros, I., Monegal, A., Bedini, J. L., Deulofeu, R., Pons, F., Munoz-Gomez, J., and Ballesta, A. M. (2001). Usefulness of biochemical markers of bone turnover in assessing response to the treatment of Paget’s disease. Bone 29, 447–452. 112. Garnero, P., Fledelius, C., Gineyts, E., Serre, C. M., Vignot, E., and Delmas, P. D. (1997). Decreased beta-isomerization of the C-terminal telopeptide of type I collagen alpha 1 chain in Paget’s disease of bone. J. Bone Miner. Res. 12, 1407–1415.

791 113. Cloos, P. A., and Fledelius, C. (2000). Collagen fragments in urine derived from bone resorption are highly racemized and isomerized: a biological clock of protein aging with clinical potential. Biochem J. 345(Pt 3), 473–480. 114. Cloos, P. A., Fledelius, C., Christgau, S., Christiansen, C., Engsig, M., Delmas, P., Body, J. J., and Gamero, P. (2003). Investigation of bone disease using isomerized and racemized fragments of type I collagen. Calcif. Tissue Int. 72, 8–17. 115. Alexandersen, P., Peris, P., Guanabens, N., Byrjalsen, I., Alvarez, L., Solberg, H., and Cloos, P. A. (2005). Non-isomerized C-telopeptide fragments are highly sensitive markers for monitoring disease activity and treatment efficacy in Paget’s disease of bone. J. Bone Miner. Res. 20, 588–595. 116. Woitge, H. W., Pecherstorfer, M., Li, Y., Keck, A. V., Horn, E., Ziegler, R., Seibel, M. J. (1999). Novel serum markers of bone resorption: clinical assessment and comparison with established urinary indices. J. Bone Miner. Res. 14, 792–801. 117. de la, P. C., Rapado, A., Diaz Diego, E. M., Diaz Martin, M. A., Aguirre, C., Lopez, G. E., and Diaz, C. M. (1996). Variable efficacy of bone remodeling biochemical markers in the management of patients with Paget’s disease of bone treated with tiludronate. Calcif. Tissue Int. 59, 95–99. 118. Gutteridge, D. H., Prince, R. L., Stewart, G., Stuckey, B., Will, R. K., Retallack, R. W., Price, R. I., and Ward, L. (1996). Comparison of biochemical markers of bone turnover in Paget disease treated with pamidronate and a proposed model for the relationships between measurements of the different forms of pyridinoline cross-links. J. Bone Min. Res. 11, 1176–1184. 119. Kaddam, I. M., Iqbal, S. J., Holland, S., Wong, M., and Manning, D. (1994). Comparison of serum osteocalcin with total and bone specific alkaline phosphatase and urinary hydroxyproline:creatinine ratio in patients with Paget’s disease of bone. Ann. Clin. Biochem. 31(Pt 4), 327–330. 120. Grauer, A., Knaus, J., Klar, B., and Ziegler, R. (1995). Relapse free interval after pamidronate treatment in Paget’s disease of bone. J. Bone Min. Res. 10(suppl 1), 510. 121. Fenton, A. J., Gutteridge, D. H., Kent, G. N., Price, R. I., Retallack, R. W., Bhagat, C. I., Worth, G. K., Thompson, R. I., Watson, I. G., and Barry-Walsh, C. (1991). Intravenous aminobisphosphonate in Paget’s disease: clinical, biochemical, histomorphometric and radiological responses. Clin. Endocrinol. (Oxf) 34, 197–204. 122. Grauer, A. (1997). Morbus Paget. In: Metabolische Osteopathien. M. J. Seibel, H. Stracke eds.). pp. 288–300. Schattauer, StuttgartNewYork.

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Chapter 48

Metastatic Bone Disease Jean-Jacques Body

Dept of Internal Medicine and Endocrinology/Bone Diseases Clinic, Institut J. Bordet, Univ. Libre de Bruxelles, Brussels, Belgium

IV. Use of Markers of Bone Turnover for the Monitoring of Tumor Bone Disease V. Prediction of the Development of Bone Metastases References

I. Abstract II. Introduction III. Use of Markers of Bone Turnover for the Diagnosis of Bone Metastases

I. ABSTRACT

with specific tumor markers, such as PSA or CA15.3, the diagnostic sensitivity of bone markers can be improved. Their degree of elevation correlates with the tumor burden and has been shown to be an independent prognostic factor for several tumors. On the other hand, biochemical markers of bone turnover have the unique potential to simplify and to improve the monitoring of metastatic bone disease which remains a continuous challenge for the oncologist. Peptide-bound crosslinks could be quite useful to discriminate between patients progressing early on treatment from those with longer disease control. Also, the diagnostic efficiency of a 50% increase in these markers could identify imminent progression. Patients with elevated baseline NTX-I (>100 nmol/mmol Creat) or BALP (>146 IU/L) suffer more frequently from a SRE than patients with normal levels, and have a poorer prognosis. On the other hand, markers of bone resorption could be especially useful to monitor the effects of bisphosphonate therapy. Future work will have to determine if adjusting bisphosphonate dosage according to the levels of specific bone markers could enhance their clinical usefulness. The level of initial elevation and the degree of suppression after therapy of bone resorption markers appears to correlate with clinical outcome in patients with lytic or blastic bone lesions.

The skeleton is the first and most common site of distant relapse in breast and prostate carcinomas. Tumor bone disease is responsible for a considerable morbidity which also makes major demands on resources for healthcare provision. Increased bone resorption in tumor bone disease appears to be essentially mediated by the osteoclasts, explaining why bisphosphonates have been successfully used for the treatment of malignant osteolysis. Hypercalcemia occurs in 10–20% of the patients with advanced cancer and the uncoupling between bone resorption and bone formation is easily demonstrated by the measurement of bone markers. The differential diagnosis between tumor-induced hypercalcemia and primary hyperparathyroidism is most often easy when using intact PTH assays; moreover, PTHrP determination can be useful in selected cases. The diagnosis of bone metastases is often easy when the patient is symptomatic. The diagnostic usefulness of bone markers is limited and available data indicate that bone markers are so far unsuitable for an early diagnosis of neoplastic skeletal involvement on an individual basis. However, by combining bone-specific alkaline phosphatase (BALP) or modern bone resorption markers Dynamics of Bone and Cartilage Metabolism

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Patients who do not normalize their markers of bone resorption under bisphosphonate therapy are at a much higher risk of bone complications.

II. INTRODUCTION A. Clinical and Pathogenic Aspects of Tumor Bone Disease: a Brief Overview 1. CLINICAL ASPECTS

According to the series, 30–90% of patients with advanced cancer will develop skeletal metastases. Carcinomas of the breast (47–85%), prostate (33–85%), and lung (32–60%) are the tumors most commonly associated with skeletal metastases. At the other end of the spectrum, tumors of the digestive tract are rarely (3–13%) complicated by metastatic involvement of bone [1]. The skeleton is, in fact, the most common site of metastatic disease and the most common site of first distant relapse in breast and prostate carcinomas [2, 3]. These patients have a relatively long survival after the diagnosis of bone metastases compared to patients with extraosseous metastases only. Patients with breast cancer and bone metastases have a median survival usually beyond 20 months and about 10% of them are still alive 5–10 years after the first diagnosis of bone metastases [2]. Tumor bone disease is responsible for a considerable morbidity and can markedly decrease the quality of life. Because of the long clinical course breast cancer may follow, morbidity from bone metastases also makes major demands on resources for health care provision [4]. Besides the complications of bone marrow invasion, pain and functional disability occur in 45–75% of the cases [1, 5]. The term “skeletal-related events” (SREs) refers to the major complications of tumor bone disease, namely pathological fractures, need for radiotherapy, need for bone surgery and hypercalcemia. Major complications will be observed in up to one-third of the patients whose first relapse is in bone [2, 4, 6, 7]. Hypercalcemia classically occurs in 10–15% of the cases and, when long bones are invaded, fractures will occur in 10–20% of the cases [6, 7]. Pathologic fractures occur with a median onset of 11 months from the initial diagnosis of bone involvement and breast carcinoma patients have a mean skeletal morbidity rate of up to four SREs per year [8]. Metastatic bone pain is traditionally attributed to various factors, notably the release of chemical mediators, an increased pressure within the bone, microfractures, the stretching of periosteum, reactive muscle spasm, and nerve root infiltration and /or compression [9]. As suggested by

the clinical and biochemical effects of bisphosphonates, the dramatic increase in bone resorption, which is easily seen with bone turnover markers, probably plays an important contributory role. Pathological fractures constitute a major cause of prolonged disability in breast cancer, whereas they occur less often in prostate cancer, with its predominantly sclerotic picture. The majority of bone metastases from breast cancer are osteolytic, but about one-fifth is purely or essentially osteosclerotic, as classically observed in prostate cancer. It is not rare that metastatic breast cancer remains confined to the skeleton for a long time. Sherry and collaborators studied 86 such patients who developed extra skeletal metastases after a mean delay of 2 years. Compared to patients who initially had metastases to other sites, these patients had a prolonged survival (median of 48 months vs. 17 months) but presented even more often with major complications of their skeletal disease. Thus, 21% developed pathological fractures of the long bones and 15% spinal cord compression [10]. This relatively slow evolution of breast cancer metastasizing in bone suggests some peculiar biological properties of these tumor cells. Indeed, breast cancers metastasizing to the skeleton are more frequently estrogen receptor-positive and well differentiated than the tumors metastasizing to lungs or liver [2, 11]. The presence of sclerotic lesions has been associated with a better prognosis than lytic lesions [12]. 2. PATHOPHYSIOLOGY OF TUMOR-INDUCED OSTEOLYSIS

The osteotropism of breast and prostate neoplasms remains incompletely understood. Preferential access of prostate cancer cells to the axial skeleton has been attributed to a passage through the vertebral venous plexus of Batson which is a low-pressure, high-volume system of vertebral veins running adjacent to the spine. On the other hand, various properties of cancer cells, such as the production of proteolytic enzymes and specific cell adhesion molecules, can enhance their metastatic potential. More specifically, deposits into the skeleton can be due to the attraction of tumor cells by chemotactic factors released by the normal remodeling of bone matrix. These factors include fragments of type I collagen and of osteocalcin, and several growth factors [13, 14]. The propensity of breast cancer cells to metastasize and proliferate in bone is currently explained by a “seed and soil” concept [15]. Breast cancer cells (the “seed”) appear to secrete factors, such as PTHrP, potentiating the development of metastases in the skeleton which constitutes a fertile “soil”, rich in cytokines and growth factors that stimulate breast cancer cell growth. Local production of PTHrP and of other osteolytic factors by cancer cells in bone stimulate osteoclastic bone resorption, essentially

Chapter 48 Metastatic Bone Disease

through the osteoblasts and probably also through the immune cells. Such factors induce osteoclast differentiation from hematopoietic stem cells and could also activate mature osteoclasts already present in bone. PTHrP also alters the ratio between osteoprotegerin (OPG), whose production is decreased, and RANKL (receptor activator for NfκB-ligand), whose production is increased [16]. The net result of this imbalance in these newly discovered and essential regulatory factors of osteoclast-mediated bone resorption is an increase in osteoclast proliferation and activity. Increased osteoclast number and activity would then cause local foci of osteolysis, an enhanced release of growth factors and a further stimulation of cancer cell proliferation [13, 15, 17]. A similar vicious cycle probably exists for metastatic prostate cancer as well [18]. Collagen cross-link metabolites are much increased in patients with bone metastases from prostate cancer, underlining an increase in bone resorption even in blastic disease [19]. Prostate cancer cells stimulate osteoclast activity probably also through the osteoblasts [18]. Multiple myeloma is characterized by a marked increase in osteoclast activity and proliferation. This excessive resorption of bone stimulates the growth of myeloma cells in bone, through the release of growth factors from resorbed bone matrix and the secretion of interleukin-6 (IL-6) by the bone microenvironment. Using established cell lines, it has been shown that through direct cellto-cell contact, myeloma cells can downregulate osteocalcin production but upregulate IL-6 secretion, supporting the concept of the importance of the bone microenvironment in the genesis of myeloma-induced osteolysis [20]. The data summarized here indicate that bone-resorbing cells are a logical target for the treatment and perhaps the prevention of tumor-induced osteolysis [21].

B. Classical Methods for the Monitoring of Metastatic Bone Disease The diagnosis of bone metastases is often easy when patients are symptomatic and a bone biopsy is rarely necessary. However, the assessment of the response to therapy remains a continuous challenge. Quantitative pain assessment is performed too rarely by the oncologist. Pain intensity can, however, be quickly measured by visual analog scales. Similarly, quality of life is rarely assessed. Such subjective changes are likely related to, and maybe precede, objective changes in bone metastases size or number, although a formal and prospective comparative assessment between these two types of evaluation has not yet been performed [22].

795 The radionuclide bone scan remains the most widely used method for diagnosis of bone metastatic involvement. The bone scan is more sensitive than X-rays for the early detection of bone metastases. However, the relative lack of specificity of the bone scan is well known and at least one-third of solitary abnormalities detected in cancer patients are due to benign diseases [23]. In one study, the appearance of one or two new abnormalities on the bone scan in cancer patients without known metastases was later confirmed to be of metastatic origin in only 11% and 24% of cases, respectively [24], but the presence of multiple focal hot spots is much more specific for metastatic bone disease. Besides our relative inability to quantitate these changes, one must be aware of the so-called flare phenomenon. This apparent worsening of the bone scan (increased scintigraphic activity in bone lesions or even the appearance of previously invisible lesions) can actually represent the initial healing of osteolytic lesions and thus be misinterpreted as evidence of tumor progression [25]. Bone markers could be useful in such cases, but this has been little documented. Radiographic methods remain the essential tool for the diagnosis and the characterization of bone metastases. Computed tomography (CT) or magnetic resonance imaging (MRI) are particularly useful to diagnose early metastatic involvement of bone when hot spots are detected on the radionuclide scan but corresponding X-rays are normal. MRI is especially helpful in detecting early bone marrow involvement in asymptomatic disease [7]. The assessment of the response of skeletal metastases to therapy remains a daily challenge for the oncologist. For example, a complete response to antineoplastic therapy of a bone metastasis requires the reversal of abnormal radiographic findings, which is highly subjective and actually almost impossible to achieve. On the other hand, development of new blastic lesions does not necessarily mean progression, because they can appear in previously lytic areas that were not detected on X-rays. Bone scans are most useful for restaging disease recurrence in order to identify new lesions. All of the imaging techniques aim to detect recalcification of lytic areas, which is evidently not an early phenomenon. Symptom evaluation and measurement of biochemical parameters should definitely be further investigated for early assessment of bone response or progression. The comparison between an objective but quite costly evaluation means, such as CT-scan, and a combination of pain assessment (with a score for analgesic consumption), performance status and an assay for a bone resorption marker certainly deserves to be performed! Markers of bone turnover could be used as diagnostic tests and to assess response to treatment. Their role to

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predict skeletal complications has been the subject of recent, but still limited, investigations.

III. USE OF MARKERS OF BONE TURNOVER FOR THE DIAGNOSIS OF BONE METASTASES Available markers are reviewed elsewhere in this book; here they will be briefly reviewed as they pertain to the cancer field. Total alkaline phosphatase (TALP) is still the marker most often used in routine oncological practice. However, to exclude contribution from the liver and other organs, the measurement of the bone isoenzyme of alkaline phosphatase (BALP) is preferable. Reproducible assays for direct BALP determination are now commercially available with a cross-reactivity for liver Alk Phos between 7 and 13%. Osteocalcin (or bone-GLA protein, BGP, or OC) is a protein specifically made by the osteoblasts and its measurement is supposed to reflect the late phases of bone formation, but its usefulness in oncology remains obscure and its meaning complicated by discordant results between different assays. Assays for the peptides released during the extracellular processing of collagen, referred to as the procollagen carboxyterminal propeptide (PICP) and the aminoterminal propeptide (PINP), might represent useful markers of bone formation because collagen is by far the most abundant organic component of bone matrix. However, published results obtained with available assays have been most often disappointing. This may be due to the relative lack of specificity of these assays for the evaluation of collagen synthesis by osteoblasts. On the bone resorption side, the fasting urinary excretion of calcium and of hydroxyproline are nowadays less used in oncology. Urinary excretion of calcium is not a true marker of bone resorption; rather it reflects the net effects of bone formation and resorption. It lacks sensitivity and specificity for diagnostic use. For example, in a series of 153 patients with bone metastases from various cancers, it has been shown that urinary calcium measurement could not differentiate patients with and without bone metastases [26]. Hydroxyproline is derived from the various forms of collagen, influenced by dietary gelatin intake, metabolized by the liver, but remains a conventional and widely available parameter to assess bone resorption. When measured in fasting urinary samples, it is a reasonably valid estimate of the bone resorption rate [27, 28]. New parameters of bone resorption are more helpful because of their increased specificity for bone matrix destruction. The molar ratio of their urinary excretion relative to creatinine excretion provides a reproducible measurement of their rate of excretion that is independent

of body size or urine dilution. The intermolecular crosslinking compounds of mature collagen, pyridinoline (PYD), and deoxypyridinoline (DPD), are particularly well suited to monitor the breakdown of bone matrix by cancer cells, even more that they are independent from dietary intake. Crosslink excretion correlates quite well with osteoclast number, as shown in a rat model of PTHrP-mediated tumor osteolysis [29]. The circadian rhythm of crosslinks excretion, with a peak in the second part of the night, implies that the samples have to be collected at the same time of the day, e.g. the second morning void. Values obtained in late afternoon are for example 50% less than values measured in the early morning [30]. Their measurement requires, however, HPLC technique and they are progressively replaced by direct, commercially available assays. Crosslinks exist in both free (40%) and peptide-bound (60%) forms and enzyme-linked immunoassays (ELISA) have been developed to measure the protein-bound crosslinking molecule at either the N-terminal part (NTX-I) or the C-terminal part (CTX-I) and the free portions of both PYR and DPD. If used after sample hydrolysis, the commercial assay for urinary free DPD can then be used for the determination of total DPD which can be advantageous, notably after bisphosphonate therapy (see below). However, there are no extensive face-to-face comparisons in cancer patients between the classical HPLC assay and these direct telopeptide assays [31]. Each of these immunoassays for free crosslinks and related peptides may reflect different aspects of bone resorption. For example, increased bone turnover is associated with a preferential increase of the peptide bound/total fraction and a decrease of the free/total fraction, at least in benign conditions [32]. A limited comparison in 30 patients with breast cancer and bone metastases suggests that determination of crosslinks by HPLC has a higher sensitivity than either the urinary or the newly developed serum assays for the N- or C-terminal telopeptides of type I collagen [33], but inhibition of bone turnover appears to result in greater changes in serum and urine concentrations of NTX-I and CTX-I compared with PYD and DPD [34]. Serum assays for crosslinks offer evident advantages. The only largely available serum assay was a C-telopeptide crosslinks assay which recognizes elements from the helical part of the collagen chain (ICTP) but the clinical value of this assay is uncertain as it appears to be an index of collagen turnover [35], but not a specific marker of bone resorption, especially in cancer patients. Direct assays for NTX-I or CTX-I are becoming widely available. The average short-term (4 days) and long-term (4 months) variablity in normal men is less than 20% but can be as high as 50% in some subjects [36]. Serum assays for N- or C-terminal telopeptides of type I collagen appear to provide a similar

Chapter 48 Metastatic Bone Disease

performance to the urinary assays, except in patients with renal failure. Limited data obtained in patients treated with intravenous pamidronate even suggest that serum CTX-I shows treatment effects more often than urinary NTX-I [37]. An immunoassay has also recently been developed for the measurement of tartrate-resistant acid phosphatase 5b (TRAcP) which does not contain sialic acid and appears to be highly specific for osteoclast activity. It could be a sensitive way to monitor antiresorptive treatment as the changes observed after 6 months of estrogen replacement therapy appear to be at least as large as the ones measured with an immunoassay for DPD [38]. Bone sialoprotein (BSP) is a non-collagenous protein of the bone extracellular matrix, accounting for 5–10% of the non-collagenous proteins of the bone matrix. It is found mainly in osteoblasts but also in osteoclasts. An assay has been developed for BSP which indicates that BSP measurement is more a marker of bone resorption than of bone formation [39]. Lastly, the clinical value of the measurement of serum concentrations of the secreted components of OPG-RANK-RANKL system starts to be evaluated but data are limited and sometimes contradictory. Markers are quite useful in the context of clinical research but much work remains to be done to prove their interest for the individual management of patients. Moreover, it has been shown that results differ markedly among laboratories, even with identical assays and methods [40]. The field of bone markers is still new, especially for the new markers described above, which can explain the small number of good studies in oncology.

797 much more stable. This has notably been shown with biochemical markers of bone turnover. In TIH there is a marked increase in crosslinks levels but quite variable concentrations of OC that are often very low, whereas both markers are increased in primary hyperparathyroidism [43, 44]. The ratio between DPD and OC is normal in primary hyperparathyroidism but markedly increased in both types of TIH, whether of humoral or local osteolytic origin (Fig. 1) [45]. Body et al. [46] measured BALP and other markers of bone turnover in 46 patients with TIH. Mean (±SD) BALP concentrations were slightly higher in patients with TIH than in healthy subjects (15.5 ± 8.5 versus 12.4 ± 3.5 mg/L; P < 0.05). However, the scatter of the data was quite marked (Fig. 2). Increased values (22%) occurred only in patients with bone metastases [46]. BALP levels also correlated with the extent of bone uptake at scintigraphy (rs = 0.54; P < 0.01) but this was not the case for total Alk Phos or OC. Bone resorption is classically considered to be greatly increased in TIH [27]. However, it is interesting to note that urinary pyridinium crosslinks are not significantly higher in hypercalcemic than in normocalcemic patients with bone metastases (see last two series of data of Fig. 3) [26]. This finding suggests that the inhibition of osteoblast activity has an important causal role in the development of TIH. Several studies have established the essential role of parathyroid hormone-related protein (PTHrP) in most types of cancer hypercalcemia. The kidneys may also contribute to the pathogenesis and maintenance of TIH through a decrease in the glomerular filtration rate and an increase in the tubular reabsorption of calcium which

A. Tumor-induced Hypercalcemia The prevalence of tumor-induced hypercalcemia (TIH) is variable in the literature; this complication can classically affect 8–40% of cancer patients at some time during the course of their disease. TIH can complicate any type of cancer, but breast and lung carcinomas and multiple myeloma represent more than 50% of the cases. Most often, TIH complicates advanced cancer and the median survival varies between 1 and 4.5 months [41]. The frequency of TIH is certainly less nowadays because of an increasing use of bisphosphonates in patients with cancer metastatic to the skeleton, but no precise figures are available. Secretion of humoral and paracrine factors by the tumor markedly stimulates osteoclast activity and proliferation, often with inhibition of osteoblast activity, causing an uncoupling between bone resorption and bone formation, and thus a rapid rise in serum calcium [42]. In contrast, in primary hyperparathyroidism, bone formation is stimulated in parallel to bone resorption and serum calcium is

Figure 1 Relationship between total deoxypyridinoline (DPD) and osteocalcin (OC) in 20 patients with primary hyperparathyroidism ( ), 31 patients with humoral hypercalcemia of malignancy (䊊) and 11 patients with hypercalcemia and bone metastases (∆). Dotted areas indicate the normal ranges. Note the suppressed OC values in many hypercalcemic cancer patients. Adapted with permission from Nakayama et al. [45].

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Figure 2 Serum concentrations of the bone isoenzyme of alkaline phosphatase (BALP) in 46 patients with TIH, divided according to the presence (BM+) or the absence (BM−) of bone metastases. Bars represent the median values. Note that increased values occurred only in the group BM+. Adapted with permission from Body et al. [46].

result from the decreased circulating volume and the specific renal effects of PTHrP. PTHrP effects are autocrine or paracrine in nature and its only known endocrine effect is its role in the pathophysiology of TIH. Animal models of humoral hypercalcemia of malignancy (HHM) constitute a useful means to assess the importance of hypercalcemic factors and, in nude mice transplanted with human

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PTHrP-producing tumors, passive immunization with antiPTHrP monoclonal antibodies can correct hypercalcemia, decrease osteoclastic bone resorption, and markedly prolong the survival time of the tumor-bearing animals [47]. The differential diagnosis between TIH and primary hyperparathyroidism has been much simplified since the availability of assays specifically measuring intact parathyroid hormone (PTH). These assays permit a complete separation of PTH levels in the two conditions. Serum PTH concentrations are indeed elevated, or at least in the upper part of the normal range in hyperparathyroid patients, whereas they are suppressed in patients with TIH [48]. Primary hyperparathyroidism is not a rare disease, however, and both conditions can coexist. An elevated intact PTH level in a hypercalcemic cancer patient is a strong indicator for the presence of hyperparathyroidism and does not imply at all that the tumor is secreting PTH [49]. Several immunoassays have been developed to measure levels of circulating PTHrP. According to the published series, 45–100% of patients with TIH have increased circulating PTHrP concentrations [49–51]. Virtually all hypercalcemic patients without bone metastases (HHM) have elevated PTHrP levels but, with a sensitive assay, Grill et al. showed that this is also the case in two-thirds of the patients with hypercalcemia and bone metastases [51]. Two-site assays, including the measurement of large N-terminal regions, appear to be optimal for TIH evaluation [52]. Circulating PTHrP concentrations do not change after successful therapy of TIH with bisphosphonates, implying that elevated PTHrP levels indeed constitute a primary phenomenon and that PTHrP secretion is not regulated in vivo by serum calcium, at least in hypercalcemic subjects [50]. On the contrary, PTH secretion rapidly recovers when serum calcium falls, and circulating PTH levels are normalized as soon as calcium enters the normal range [53], implying that blood sampling for diagnostic purposes must preferably be done before starting bisphosphonate therapy.

B. Metastatic Bone Disease The diagnostic usefulness of biochemical markers for the individual diagnosis of bone metastases is so far limited even if several markers of bone turnover (BALP, ICTP, DPD) correlate with the extent of metastatic bone disease [54]. Figure 3 Urinary pyridinium crosslinks levels in 153 healthy controls, 55 patients without bone metastases (NBM); 42 of them had a normal serum Ca level (NC) and 13 were hypercalcemic (HC), and 98 patients with bone metastases (BM; 56 of them had a normal serum Ca level (NC) and 42 were hypercalcemic (HC)). Adapted with permission from Pecherstorfer et al. [26].

1. MARKERS OF BONE FORMATION

BALP levels appear to better reflect bone metastatic involvement than total Alk Phos or OC in hypercalcemic patients [46]. In breast cancer, the sensitivity of BALP

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determination (whether by lectin precipitation, EIA, or IRMA) does not appear to be better than that of total Alk Phos [55]. Some data indicate that the diagnostic sensitivity of BALP for the diagnosis of bone metastases from breast cancer could be superior to that of CA15.3 but more data are needed [56]. Compared to total Alk Phos, the main advantage of measuring BALP in prostate cancer appears to be a higher positive predictive value for the diagnosis of bone metastatic involvement [57], probably due to a higher diagnostic specificity, as suggested by the study of Zaninotto et al. [58]. In 65 patients with metastatic prostate cancer, they found specificities of 57% and 90% for total Alk Phos and BALP, respectively [58]. Highest levels are seen in patients with multiple blastic metastases, especially for BALP [59]. BALP appears to have a clearly better sensitivity than total Alk Phos in patients with metastatic prostate cancer when total Alk Phos is in the range of normal to twice normal [60]. Serum BALP levels correlate with the number of radiologically identified blastic lesions and, not surprisingly, BALP is more often elevated in prostate cancer (72%) and in breast cancer (60%) than in metastatic lung cancer (5%) since these patients most often have a purely lytic disease [55]. In a study by Garnero et al., including 39 patients with prostate cancer and bone metastases, the diagnostic sensitivity was 62% for BALP but also for total Alk Phos. The sensitivity of PICP was 60% but only 43% for OC. Interestingly, if one excepts OC, markers of bone resorption were increased approximately in the same proportion, 61% for urinary αCTX-I, 53% for urinary βCTX-I, and 64% for serum CTX-I (Fig. 4) [61]. Limited data indicate that BALP could be superior to PSA as a marker for bone metastases [62] and that the

combination of both markers has a better diagnostic sensitivity than either marker alone [63]. Compared to the upper limit of normal, BALP could detect 88% of the patients with bone metastases with a specificity of 100%. By combining BALP with PSA, the sensitivity to detect bone metastases increased in that study to 96% [63]. PICP levels typically correlate with BALP, especially in patients with blastic metastases [59]. BALP could better reflect the inhibition of bone formation than OC in myeloma patients and values are significantly lower than in patients with MGUS. However, the overlap is considerable and the clinical usefulness for the differential diagnosis is doubtful [64]. On the contrary, others have found that BALP levels did not differ between the various stages of myeloma, whereas OC levels were lower in stages III and II than in stage I. Moreover, in that study, OC, but not BALP, levels were inversely correlated with bone resorption markers [65]. Increased OC levels are not rarely observed in patients without bone metastases [43, 66]. In a series of 60 patients with recurring breast cancer, less than 50% of patients with bone involvement had elevated OC levels [67]. OC also is less sensitive than PSA or prostatic acid phosphatase (PAP) for the detection of bone metastases in prostate cancer [68]. OC levels are nevertheless significantly increased in metastatic bone disease from breast and prostate cancers [69], but not in multiple myeloma. Bataille et al. have shown that a subgroup of patients with multiple myeloma (around 12%) had increased OC levels with a more indolent disease and a lower osteolytic potential, whereas patients with decreased OC values had a more severe disease [70]. More generally, however, a large study showed that OC was a poor marker of the severity of multiple myeloma

Individual concentrations of serum BALP and urinary α-CTX in 39 patients with bone metastases from prostate cancer (PC. M+) as compared to eight patients with benign prostate hyperplasia (BPH) and nine patients with localized prostate cancer (PC. M−). Note that the bone resorption marker (α CTX ) is increased in the same percentage of patients as the classical marker of bone formation (BALP). Adapted with permission from Garnero et al. [61].

Figure 4

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and that it did not correlate with survival; however, the study confirmed the good correlation of OC levels with histomorphometric parameters of bone formation and not bone resorption in this typical condition of bone uncoupling [71]. OC levels also do not correlate with BALP in myeloma [71, 72]. Preliminary results obtained with propeptide assays of type 1 collagen also appear to be disappointing, at least in patients with non-neoplastic diseases, because their sensitivity is lower than that of BGP or BALP [73]. PICP has been claimed to have a greater diagnostic specificity than PSA, prostatic acid phosphatase, or total Alk Phos for the diagnosis of bone metastases in prostate cancer, but the superiority of PICP for diagnostic purposes is not convincing. In the ROC analysis, the areas under the curves were actually quite similar for all four markers, whereas the separation between patients with or without bone metastases was the greatest for PSA [74]. However, limited data suggest that PICP could be of value to separate patients who respond to therapy from patients who progress after therapy [74]. Osteoprotegerin (OPG) is overexpressed in bone metastases from prostate cancer patients and could play a role in the development of osteoblastic bone metastases [75]. Serum OPG levels have been reported to be markedly increased in patients with prostate cancer and bone

Figure 5

metastases. RANKL levels were not different from the control group. Interestingly, OPG had a better diagnostic accuracy than Alk Phos or bone resorption markers [76]. The same group recently confirmed their initial findings when comparing ten serum bone turnover markers in 117 patients with prostate cancer at various stages of the disease. OPG and TRAcP were shown to have the best diagnostic efficiency in a logistic regression model [77]. 2. MARKERS OF BONE RESORPTION

Urinary Ca is a poor diagnostic marker for bone metastases, whereas hydroxyproline is typically increased in patients with breast cancer metastatic to bone, but generally not at an early stage. In patients with bone metastases from breast cancer, hydroxyproline, PYD, and DPD are significantly increased compared to normal values, but the proportionate increase is higher for PYD and DPD (Fig. 3 and 5). In a review of six studies, elevated levels of PYD and DPD were found in 58–88% and 60–80%, respectively [78]. Hou et al. found that they are about twice as high as compared to values in patients without bone metastases [79]. However, Lipton et al. found elevated PYD and DPD in 66% of patients with bone metastatic involvement; however, elevated values were also found in 42% of cancer patients without documented bone metastases [80]. This surprisingly high percentage in patients

Individual concentrations of uCa, hydroxyproline, Pyr, D-Pyr, and Crosslaps (CTX) in 19 normocalcaemic patients with breast cancer metastatic to bone. Crosslinks were measured by HPLC. Values are compared to the upper limit of normal values in premenopausal (------) or postmenopausal (——) women. Note the larger effect of menopause on the direct peptide-bound CTX assay than on the HPLC measurement of crosslinks. Adapted with permission from Body et al. [84].

Chapter 48 Metastatic Bone Disease

without bone metastases could reflect an excellent sensitivity but it could also reflect a poor specificity of the pyridinoline crosslinks. The interesting possibility that this could reflect an early bone involvement has yet to be proven, as acknowledged in an editorial written by these investigators [81]. Similar results have been reported by Pecherstorfer et al. who found significantly higher values in patients with bone metastases, but the discrimination was poor (Fig. 3) [26]. This could again indicate a subclinical metastatic involvement, but more likely it is due to a subclinical osteolysis in advanced cancer patients without overt bone metastases under the influence of premature menopause, antineoplastic therapy, PTHrP, or other unknown bonemobilizing humoral tumor products, or to the release of pyridinium crosslinks from extraosseous tissues, as suggested by the slight increase in the ratio PYD/DPD [26]. Anyway, these data indicate that these markers are unsuitable for an early diagnosis of neoplastic skeletal involvement. NTX-I (urine N-telopeptide assay) has been reported to have the highest discriminatory power and to be the most predictive biochemical marker of bone metastases when comparing groups of patients, but it appears less useful on an individual basis [82]. When cutoff values are chosen to give specificities between 90–100%, the NTX-I shows a sensitivity around 30%, implying that 70% of subjects with bone metastases would be missed. Lipton et al. reported that the sensitivity of NTX-I was higher than the one of crosslinks as measured by HPLC, but the values were compared to the upper limit of normal in premenopausal women which is probably inadequate in metastatic breast cancer patients who are typically postmenopausal [83]. We found that the diagnostic sensitivity of the CTX-I assay was poor when the values were compared to those of healthy postmenopausal women. The diagnostic sensitivity was actually the highest for the collagen crosslinks when measured by the classical HPLC method (see Fig. 5) [84]. Moreover, the biological variability of telopeptide assays is higher than that of the free DPD assay [85]. A direct ELISA for the measurement of urinary free DPD is commercially available and the correlation with HPLC determination appears to be adequate [86]. Patients with bone metastases have significantly higher levels than patients without bone metastases, but DPD excretion was again often elevated in these patients, suggesting a higher rate of bone turnover in breast cancer patients without bone metastases. This result indicates that the diagnostic efficiency for bone metastatic involvement for this marker will be similar to what has been described before [87]. Preliminary data obtained with a new ELISA for TRAcP5b in metastatic breast cancer are quite encouraging and this assay certainly deserves further investigation [88].

801 Limited data obtained with BALP, ICTP, PICP, and BGP measurements in 112 postmenopausal patients with breast cancer, 22% of whom had bone metastases, indicate that these markers are unlikely to replace bone scintigraphy. To achieve a sensitivity of 95%, as compared to a positive bone scan, the combined measurement of BALP and ICTP then had a specificity of less than 10% [89]. Houzé et al. have nevertheless reported a positive relationship between bone scintigraphy (scored 0 to 4) and urinary CTX-I in 84 patients with metastatic breast cancer, 38 of whom had bone metastases [90]. Baseline serum NTX-I levels have been shown to be of prognostic value in a retrospective study of 250 metastatic breast cancer patients with bone-only or bone plus soft tissue metastases starting a second-line hormonal therapy. Values were elevated in 24% of the patients (i.e., above 26 nM BCE corresponding to the mean value + 2 SD in postmenopausal women). The group with increased NTX-I levels had a shorter duration of clinical benefit, a shorter time to progression and a shorter survival. In a multivariate analysis, NTX-I had about the same prognostic value as the performance status and the use of adjuvant chemotherapy. Serum NTX-I thus appears to reflect disease burden in bone and could possibly be used as a stratification factor for clinical trials [91]. New markers of bone turnover could thus help to select patients at particular risk of developing bone complications and who would more likely benefit from an early intervention. Collagen crosslink metabolites are also increased in patients with bone metastases from prostate cancer, underscoring the existence of increased bone resorption even in blastic disease. These metabolites permit a better separation from patients with localized disease than does hydroxyproline, but the correlation with a bone scan score is similar for all three resorption markers [92]. Elevated CTX-I levels also correlate with the degree of skeletal involvement [61]. Increased values of ICTP have been associated with a poor prognosis in prostate cancer, possibly reflecting the lytic component of the metastases [93]. In a study including 112 patients, predictors of skeletal morbidity included more extensive bone lesions, more severe pain, lower performance status, higher serum Alk Phos, and higher urinary DPD. Moreover, in a multivariate analysis, only DPD was associated with an increased risk for skeletal complications. Using the optimal cutoff, the sensitivity to predict skeletal events was nevertheless low (51%) but there was only an 8% false positive rate [94]. One has to be cautious, however, before claiming that increased bone resorption in metastatic prostate cancer is only due to metastatic bone disease. Androgen blockade, which is extensively used in prostate cancer [95], is associated with an increase in bone resorption. It has thus been reported

802 that BMD loss can reach 3–5% yearly during the first few years of androgen deprivation therapy [96]. Bone resorption markers are also elevated in myeloma patients, especially in stage III disease and their degree of elevation correlates with the importance of bone involvement [65]. TRAcP is another sensitive marker of bone resorption in myeloma and has been shown to be higher in patients with more than three lytic lesions and/or fractures than in patients with 1–3 lytic lesions. NTX-I levels were measured in the same study but a meaningful comparison has not been performed between these two markers [97]. Bone sialoprotein (BSP) has been more recently evaluated as a new marker of bone turnover in various conditions. BSP serum levels predominantly reflect processes related to bone resorption. BSP levels decrease in a manner similar to that of DPD after bisphosphonate therapy [33]. The diagnostic sensitivity of BSP for discriminating patients with and without bone metastases appears to be inferior to that of BALP [98]. In a series of 62 patients with newly diagnosed multiple myeloma, BSP determination appeared to be superior to that of crosslinks for the discrimination between myeloma and MGUS or osteoporosis. Serum BSP was also an independent prognostic factor, as was serum monoclonal protein [99].

IV. USE OF MARKERS OF BONE TURNOVER FOR THE MONITORING OF TUMOR BONE DISEASE Biochemical markers of bone turnover could be useful to assess the response assessment of bone metastases to antineoplastic therapy and to monitor the effects of bisphosphonates.

A. Response Assessment of Bone Metastases The biochemical markers of bone turnover have the potential to simplify and to improve the monitoring of metastatic bone disease, especially because it is notoriously difficult to rapidly evaluate the bone response to cancer therapy. There is, however, an urgent need for large-scale studies with the new markers whose usefulness has to be compared with the classical means used to assess the response of bone metastases to therapy. Coleman et al. have shown, several years ago, a decrease in uCa 1 month after therapy in patients subsequently classified as ‘‘partial responders’’. In the same patients, these authors also showed a transient increase in BALP and in OC levels and the diagnostic efficiency of a >10% early rise in BALP was 68% for discriminating response

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from progression [100]. It has then been proposed that a persistent increase in levels of bone formation markers is a sign of progressive metastatic disease, while patients in whom osteoblast activity is enhanced because of a favorable response to systemic therapy, formation markers only show a transient increase, similar to the flare phenomenon well described for bone scan. This attractive hypothesis should be further evaluated, especially as the considered elevations in bone formation markers were low [69]. The few available studies indicate that the BALP assay could be more useful for the follow-up of bone metastases than for an early diagnosis. Cooper et al. have thus reported some patients in whom BALP levels increased before the elevation of a specific tumor marker [101]. Limited data in 73 breast cancer patients with bone metastases under specific antineoplastic treatment, but also receiving pamidronate therapy, indicate that total Alk Phos was the only parameter which normalized significantly more often in the group of patients with responding or with stable disease as compared to patients with progressive disease. The other evaluated parameters in that study were PINP, ICTP, DPD, and the tumor marker CA27.29 [102]. The measurement of osteoblastic products in the serum, such as OC or the newly developed assay for the procollagen extension peptides, is not easy to interpret because changes can signify regression of osteoblastic foci or healing of osteolytic lesions. OC does not seem to increase in case of progression of bone metastases in prostate cancer, as it does in PICP and BALP [103]. Limited results indicate that BALP measurement could be useful to monitor bone metastases in prostate cancer [63] and that BALP levels correlate with pain [58]. However, in prostate cancer, BALP and PSA changes were concordant only in 69% of 49 cases with metastatic disease [60] and the clinical potential of BALP for monitoring tumor bone disease remains uncertain. In myeloma patients, an increase in OC levels during treatment was initially reported to be a good indicator of treatment efficacy, whereas decreasing levels appeared to be correlated with disease progression [70]. However, the clinical interest of OC for follow-up remains unclear, as both falling and rising serum levels have been reported in patients with progressive multiple myeloma [104]. Concerning the bone resorption markers, a preliminary report of PYD and DPD measurement in breast cancer patients receiving endocrine treatment had shown that crosslinks excretion increased during progressive disease but remained stable or decreased in patients with stable disease or in responders [105]. Hou et al. confirmed that DPD levels rose in patients with progressive disease, whether they were receiving bisphosphonates or not [79]. In contrast, Walls et al. reported no significant changes

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in PYD or DPD in responding patients [106]. It has been claimed that NTX-I levels could correlate more closely with evolution of bone metastases than the conventional tumor marker CA 15-3 but, again, more data are needed to substantiate this statement [107]. Another preliminary report of 37 well-evaluated patients suggests that NTX-I could be quite useful to discriminate reliably between patients progressing early on in treatment from those with longer disease control [108]. In patients with progressive disease, NTX-I levels steadily rose and the diagnostic efficiency of a 50% increase in NTX-I for identifying imminent progression was 78% [109]. These preliminary data have been confirmed in a series of 97 patients with bone metastases, 54 of whom had breast cancer. Three markers were sequentially measured, namely NTX-I, ICTP, and BALP. For predicting disease progression in bone, NTX-I had the highest positive predicitive value (71%) as compared to ICTP (59%), BALP (50%) and a specific tumor marker (53%). The average percentage changes were 152% for NTX-I and 144% for ICTP (both P < 0.001) whereas changes in BALP were not significant. Diagnostic accuracy, estimated by ROC curves analysis, was also the highest for NTX-I (Fig. 6). The most likely cutoff point of NTX-I for disease progression was an increase to 130% from previous levels, which had a sensitivity, of 70%, a specificity of 80%, and a positive predictive value of 72% [110]. Tumor markers were significantly increased only in patients with progression of extraskeletal disease; however, one must take into account the heterogeneity of the tumor types in this study before generalizing this statement. Interestingly, the authors did not find any evidence of significant interaction between bisphosphonate

Figure 6

administration and bone metastases progression on NTX-I levels, suggesting that the expected change in bone marker levels after disease progression can be safely assumed to be constant, regardless of bisphosphonate administration [110]. Again, this important finding is original but needs further confirmation. The value of NTX-I as a predictor of “skeletal complications” (classical SREs but including also admission for pain control and death) has also been reported by Brown and collaborators. They evaluated a total of 121 patients, including 91 with breast cancer and 26 with prostate cancers, by monthly measurements of NTX-I. NTX-I levels were correlated with the number of SREs and/or death (r = 0.62 for 0–3 months and r = 0.46 for 4–6 months). All patients received bisphosphonate therapy during the study but 93 had not received any before the first NTX-I measurement. Patients with baseline NTX-I value > 100 nmol/mmol Creat were almost 20 times (95% CI, 7.6–50) more likely to experience a SRE/death during the first 3 months than those with NTX-I < 100 [111]. The prognostic values of NTX-I and BALP have recently been evaluated in 203 patients with bone metastases from prostate cancer and in 238 patients with bone metastases from various tumors (except breast, myeloma, and prostate). These patients had been assigned to the placebo arms of two phase III trials of zoledronic acid. As shown in the two upper panels of Figure 7, patients with elevated baseline NTX-I (> 100 nmol/mmol Creat, corresponding approximately to the upper limit of normal in postmenopausal women and in men receiving androgen deprivation therapy) or BALP (> 146 IU/L) suffered more frequently from a SRE than patients with low (“normal”)

Diagnostic accuracy of bone and tumor marker levels for disease progression in bone (ROC curves) in a series of 97 patients with bone metastases from various tumors (54 had breast cancer). NTX-I levels were corrected for the administration of bisphosphonates and had the best diagnostic accuracy (see text). Adapted with permission from Costa et al. [110].

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Figure 7 Predictive value of bone markers for the proportion of patients with either prostate cancer or lung/other solid tumors who will present at least one SRE. The two upper panels correspond to the first month of follow-up according to baseline NTX-I or BALP levels. The two lower panels indicate the proportions of patients experiencing at least one SRE for all study intervals during the trial according to the most recent NTX-I and BALP measurement. Comparisons are made between patients with low (L) or high (H) values for both markers. More patients with high levels of bone markers will present at least one SRE than patients with low levels. The predictive value of NTX-I was better than the one of BALP. Adapted with permission from Brown et al. [112].

levels of NTX-I or BALP. This was also true when considering SREs during study intervals according to the most recent level of NTX-I or BALP (two lower panels of Fig. 7). NTX-I was a stronger prognostic indicator of negative outcomes than BALP. In recent assessments in prostate cancer, patients with a high NTX-I level had an increased relative risk of SREs (3.25, 95% CI 2.26–4.68), disease progression (2.02, 1.48–2.74) and death (4.59, 2.82–7.46). Although statistically significant, the predictive values of NTX-I for these endpoints were lower in the group of patients with other tumors [112]. A recent review also concluded that type I collagen telopeptides are currently the most promising tools for the assessment of therapeutic response of bone metastases [113]. In prostate cancer, effective endocrine treatment led to normal levels of DPD, and in 13 out of 15 patients with progressive disease, increasing levels of DPD were found [114]. Lastly, ICTP has been reported to be a sensitive and convenient way to monitor osteoclastic activity in multiple myeloma [115] and possibly also in prostate cancer [116]. In a series of 26 patients with newly diagnosed multiple myeloma and normal renal function, ICTP showed the highest predictive value for SREs in a multivariate Cox model including classical prognostic factors and urinary

NTX-I [117]. Limited data suggest that TRAcP could be useful to monitor antimyeloma treatment. During the follow-up of 63 myeloma patients treated by chemotherapy (VAD) and pamidronate, five patients progressed. All five had an increase in TRAcP levels at least 2 months prior to radiological progression and 6 months before the confirmation of progression of myeloma; no information is given on the predictive value of the other markers. During antineoplastic therapy, there was indeed a fall in TRAcP, NTX-I, IL-6, and β2 microglobulin levels and, interestingly, no significant change in bone formation markers (OC, BALP, and OPG) [97]. Thus, biochemical markers of bone turnover appear to be better suited for the monitoring of bone metastases than for the detection of metastatic bone involvement. Recent reports are quite promising but the data are still fragmentary and need confirmation.

B. Monitoring of Bisphosphonate Therapy Bisphosphonates constitute one of the most important therapies for the supportive care of the cancer patient. Bisphosphonates successfully reverse hypercalcemic

Chapter 48 Metastatic Bone Disease

episodes, relieve bone pain, may lead to recalcification of lytic metastases, protect the skeleton from new lesions, and can exert favorable effects on the quality of life. Prolonged use of clodronate, pamidronate, zoledronate (zoledronic acid), and ibandronate also decreases the frequency of skeletal-related events in breast cancer patients with metastatic bone disease. The mean number of skeletal complications of metastatic bone disease is indeed reduced by about 40% when using zoledronate or ibandronate in addition to antineoplastic treatment in breast cancer. The effect could be even larger in patients with multiple myeloma. More recently, it has been shown that zoledronate also reduces the skeletal morbidity rate in a variety of other tumors [118, 119]. However, it is possible that current therapeutic schemes could be improved to achieve better therapeutic results following markers of bone turnover. Markedly increased levels of PYD and DPD, and even more elevated levels of NTX-I and CTX-I, have been found in hypercalcemic cancer patients with a reduction of 80–90% after pamidronate therapy for NTX-I and CTX-I [27, 120]. Parameters of bone matrix resorption markedly fall already after a single bisphosphonate infusion. The fall in CTX-I (or NTX-I) is particularly impressive and larger than that of hydroxyproline or DPD, indicating the potential of telopeptides for delineating the optimal therapeutic schemes of new bisphosphonates, especially the optimal frequency of the infusions (Fig. 8) [84]. Free crosslinks decrease less than total crosslinks after bisphosphonate therapy. Similarly, in physiological conditions such as an oral calcium load, there are significant changes of total DPD but not of free DPD. The degradation

Figure 8 Relative changes in markers of bone resorption after a single infusion of pamidronate in 19 normocalcemic patients with breast cancer metastatic to bone. Changes are shown for urinary DPD (), PYD (), CTX (), hydroxyproline (▲), and calcium (䊊). Adapted with permission from Body et al. [84].

805 process of peptide-bound to free DPD could render the measurement of free DPD insensitive to acute changes of osteoclast activity [121]. Interestingly, NTX-I and BALP appear to fall in a similar manner after prolonged bisphosphonate therapy, whereas ICTP levels are not affected [110]. Moreover, such markers could predict the response to bisphosphonate therapy and the risk of subsequent bone events. The biochemical response can predict for pain relief, as an analgesic response has been reported more frequently in the patients in whom NTX-I, CTX-I, or free DPD remain in the normal range or return to normal values (53–63%, dependent on the marker) than in the patients whose markers do not return to normal values (0–20% of clinical response) [122]. It has been recently shown that a high rate of bone resorption is one of the factors underlying a poor response to bisphosphonate therapy and a good predictor of the likelihood of future skeletal events. Lipton et al. have reported that patients who keep an elevated NTX-I (i.e. above the normal range in premenopausal women) after 6 months of pamidronate therapy have a higher frequence of skeletal-related events. Of 21 patients who had an initially high NTX-I value, 12 went from abnormal to normal and the 9 others still had increased NTX-I levels at the end of the evaluation. There were fewer fractures in the first group than in the second one (5/12 vs. 8/9; P = 0.07) and disease progression in bone was less frequent (3/12 vs. 7/9; P = 0.03) [83]. Also, an increase of >50% in NTX-I excretion was found to be predictive of disease progression in patients treated with oral pamidronate [108]. Limited data suggest that low levels of urinary PYD and serum ICTP are significant predictors of non-progressive bone disease during short-term pamidronate therapy [123]. Findings with ibandronate in multiple myeloma also demonstrate the correlation between the inhibition of bone resorption and the clinical response to bisphosphonates. In a 2-year placebo-controlled trial of monthly 2-mg ibandronate or placebo injections in 198 evaluable patients with multiple myeloma, it was shown that this low dose ibandronate therapy did not significantly reduce skeletal morbidity. However, in the patients who had mean relative decreases, ≥ 30% for BGP or ≥ 50% for CTX-I (these criteria approximate a two-fold variation coefficient of the analytic and biologic variability of the respective marker), ibandronate achieved a highly significant decrease in the skeletal morbidity rate [124]. Theoretically at least, one could thus individualize the therapeutic schemes and optimize the efficiency of bisphosphonate therapy following changes in bone marker levels. More validation is, however, required before bone markers can be used for that purpose on an individual basis.

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V. PREDICTION OF THE DEVELOPMENT OF BONE METASTASES An important future role for bisphosphonate treatment will be the prevention of the development of bone metastases. Although available results are still somewhat contradictory and have only been obtained with clodronate, the results of a large controlled trial are very encouraging. Powles et al. conducted a double-blind trial involving 1079 breast cancer patients after surgery who received for 2 years either 1600 mg clodronate or placebo daily. There was a significant reduction in the incidence of bone metastases in the clodronate-treated group [125]. Moreover, recently updated data after a median follow-up of 5 years have shown an increased survival [126]. One can speculate that current trials testing the efficacy of more potent bisphosphonates will yield even better results. It will be essential to filter out the patients at high risk of developing bone metastases before recommending a general primary preventive use of bisphosphonates. The financial burden of a widespread use of bisphosphonates in the adjuvant setting would be enormous. On the other hand, the safety of long-term treatment with bisphosphonates, probably given at relatively high doses, must still be established in patients without bone metastases. Breast cancer is a heterogeneous disease with a large variation in growth rate and in metastatic potential. The most powerful prognostic factor for long-term survival is the nodal status: patients with positive axillary nodes have a greater likelihood of recurrence and death than node-negative patients. The latter group, however, still has a risk of approximately 30% of death from metastatic disease. The identification of new prognostic factors, that would be independent from these conventional variables, would be of great value. Much work looking for predictive proteins produced by the tumor tissue is ongoing but, so far, no such marker has unequivocally proven its value. As suggested by initial data, bone sialoprotein (BSP) could be an interesting marker [127, 128]. The primary structure of BSP contains an RGD domain that confers to BSP its cell binding activity. The RGD sequence is an integrin-recognition motif and it is known that osteoclasts bind to BSP via an αvβ3 integrin-type receptor. Such an interaction might represent a mechanism by which circulating breast cancer cells bind to bone. Initial data suggesting that serum BSP levels before surgery could be predictive of the future development of bone metastases [129] but these exciting data have not been confirmed in further analyses using BSP02 other markers of bone turnover [130]. It is probably unlikely that a circulating bone marker will ever prove useful for that matter. More fruitful approaches lie in genomic microarray and proteomics techniques.

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References 1. Galasko, C. S. B. (1986). Skeletal metastases. Clin. Orthop. Rel. Res. 210, 18–30. 2. Coleman, R. E., and Rubens, R. D. (1987). The clinical course of bone metastases from breast cancer. Br. J. Cancer 55, 61–66. 3. Kamby, C., Vejborg, I., Daugaard, S., Guldhammer, B., Dirksen, H., Rossing, N., and Mouridsen, H. T. (1987). Clinical and radiologic characteristics of bone metastases in breast cancer. Cancer 60, 2524–2531. 4. Body, J. J. (2001). Bisphosphonates in the treatment of metastatic breast cancer. J. Mam. Gland Biol. Neoplasia 6, 477–485. 5. Body, J. J., Lossignol, D., and Ronson, A. (1997). The concept of rehabilitation of cancer patients. Curr. Opin. Oncol. 9, 332–340. 6. Elte, J. W. F., Bijvoet, O. L. M., Cleton, F. J., van Oosterom, A. T., and Sleeboom, H. P. (1986). Osteolytic bone metastases in breast carcinoma: pathogenesis, morbidity and bisphosphonate treatment. Eur. J. Cancer Clin. Oncol. 22, 493–500. 7. Body, J. J. (1999). Bone metastases. In “Handbook of Supportive Care in Cancer” (J. Klastersky, S. C. Schimpff, H. J. Senn, eds.), pp. 453–481. 2nd ed. Marcel Dekker Inc., New York, Basel, Hong Kong. 8. Lipton, A. (2000). Bisphosphonates and breast carcinoma: present and future. Cancer 88(12 Suppl), 3033–3037. 9. Mercadante, S. (1997). Malignant bone pain: pathophysiology and treatment. Pain 69, 1–18. 10. Sherry, M. M., Greco, F. A., Johnson, D. H., and Hainsworth, J. D. (1986). Metastatic breast cancer confined to the skeletal system. An indolent disease. Am. J. Med. 81, 381–386. 11. Kamby, C., Rasmussen, B. B., and Kristensen, B. (1989). Oestrogen receptor status of primary breast carcinomas and their metastases. Relation to pattern of spread and survival after recurrence. Br. J. Cancer 60, 252–257. 12. Yamashita, K., Koyama, H., and Inaji, H. (1995). Prognostic significance of bone metastasis from breast cancer. Clin. Orthop. Rel. Res. 312, 89–94. 13. Rodan, G. A. (2003). The development and function of the skeleton and bone metastases. Cancer 97(3 Suppl), 726–732. 14. Body, J. J. (2000). Tumor Bone Diseases and Osteoporosis in Cancer Patients. Marcel Dekker Inc, New York & Basel. 15. Mundy, G. R. (1997). Mechanisms of bone metastasis. Cancer 80, 1546–1556. 16. Hofbauer, L. C. (1999). Osteoprotegerin ligand and osteoprotegerin: novel implications for osteoclast biology and bone metabolism. Eur. J. Endo. 141, 195–210. 17. Lacroix, M., Siwek, B., Marie, P. J., and Body, J. J. (1998). Production of interleukin-11 by breast cancer cells. Cancer Lett. 127, 29–35. 18. Goltzman, D., and Rabbani, S. A. (2000). Pathogenesis of osteoblastic metastases. In “Tumor Bone Diseases and Osteoporosis in Cancer Patients” (J. J. Body, ed.), pp. 71–84. Marcel Dekker Inc, New York & Basel. 19. Body, J. J. (2003). Rationale for the use of bisphosphonates in osteoblastic and osteolytic bone lesions. Breast 12, S37–44. 20. Barillé, S., Collettes, M., Batailles, R., and Amiots, M. (1995). Myeloma cells upregulate interleukin-6 secretion in osteoblastic cells through cell-to-cell contact but downregulate osteocalcin. Blood 86, 3151–3159. 21. Body, J. J., and Mancini, I. (2002). Bisphosphonates for cancer patients: why, how and when? Supp. Care Cancer 10, 399–407. 22. Body, J. J., Diel, I. J, Bell, R., Pecherstorfer, M., Lichinitzer, M. R., Lazarev, A. F., Tripathy, D., and Bergström, B. (2004). Oral ibandronate improves bone pain and preserves quality of life in patients with skeletal metastases due to breast cancer. Pain 111, 306–312. 23. Gold, R. I., Seeger, L. L., Bassett, L. W., and Steckel, R. J. (1990). An integrated approach to the evaluation of metastatic bone disease. Radiol. Clin. North Am. 28, 471–483.

807

Chapter 48 Metastatic Bone Disease 24. Jacobson, A. F., Cronin, E. B., Stomper, P. C., and Kaplan, W. D. (1990). Bone scans with one or two new abnormalities in cancer patients with no known metastases: frequency and serial scintigraphic behavior of benign and malignant lesions. Radiology 175, 229–232. 25. Coleman, R. E., Mashiter, G., Whitaker, K. B., Moss, D. W., Rubens, R. D., and Fogelman, I. (1988). Bone scan flare predicts successful systemic therapy for bone metastases. J. Nucl. Med. 29, 1354–1359. 26. Pecherstorfer, M., Zimmer-Roth, I., Schilling, T., Woitge, H. W., Schmidt, H., Baumgartner, G., Thiebaud, D., Ludwig, H., and Seibel, M. J. (1995). The diagnostic value of urinary pyridinium cross-links of collagen, serum total alkaline phosphatase and urinary calcium excretion in neoplastic bone disease. J. Clin. Endocrinol. Metab. 80, 97–103. 27. Body, J. J., and Delmas, P. D. (1992). Urinary pyridinium cross-links as markers of bone resorption in tumor-associated hypercalcemia. J. Clin. Endocrinol. Metab. 74, 471–475. 28. Body, J. J., Dumon. J. C., Piccart, M., and Ford, J. (1995). Intravenous pamidronate in patients with tumor-induced osteolysis: a biochemical dose-response study. J. Bone Miner. Res. 10, 1191–1196. 29. Juraschek, M., Seibel, M. J., Woitge, H. W., Krempien, B., and Bauss, F. (2000). Association between histomorphometry and biochemical markers of bone turnover in a longitudinal rat model of parathyroid hormone-related peptide (PTHrP)-mediated tumor osteolysis. Bone 26, 475–483. 30. Blumsohn, A., Herrington, K., Hannon, R. A., Shao, P., Eyre, D. R., and Eastell, R. (1994). The effect of calcium supplementation on the circadian rhythm of bone resorption. J. Clin. Endocrinol. Metab. 79, 730–735. 31. Fermo, I., Arcelloni, C., Casari, E., and Paroni, R. (1997). Urine pyridinium cross-links determination by Beckman Cross Links kit. Clin. Chem. 43, 2186–2187. 32. Garnero, P., Gineyts, E., Arbault, P., Christiansen, C., and Delmas, P. D. (1995). Different effects of bisphosphonate and estrogen therapy on free and peptide-bound crosslinks expression. J. Bone Miner. Res. 10, 641–649. 33. Woitge, H. W., Pecherstorfer, M., Li, Y., Keck, A. V., Horn, E., Ziegler, R., and Seibel, M. J. (1999). Novel serum markers of bone resorption: clinical assessment and comparison with established urinary indices. J. Bone Miner. Res. 14, 792–801. 34. Coleman, R. E., Purohit, O. P., Vinholes, J. J., and Zekri, J. (1997). High dose pamidronate: clinical and biochemical effects in metastatic bone disease. Cancer 80(8 Suppl), 1686–1690. 35. Hassager, C., Jensen, L. T., Podenphant, J., Thomsen, K., and Christiansen, C. (1994). The carboxy-terminal pyridinoline cross-linked telopeptide of type I collagen in serum as a marker of bone resorption: the effect of Nandrolone decanoate and hormone replacement therapy. Calcif. Tissue Int. 54, 30–33. 36. Orwoll, E. S., Bell, N. H., Nanes, M. S., Flessland, K. A., Pettinger, M. B., Mallinak, N. J., and Cain, D. F. (1998). Collagen N-telopeptide excretion in men: the effects of age and intrasubject variability. J. Clin. Endocrinol. Metab. 83, 3930–3935. 37. Rosen, H. N., Moses, A. C., Garber, J., Iloputaife, I. D., Ross, D. S., Lee, S. L., and Greenspan, S. L. (2000). Serum CTX: a new marker of bone resorption that shows treatment effect more often than other markers because of low coefficient of variability and large changes with bisphosphonate therapy. Calcif. Tissue Int. 66, 100–103. 38. Halleen, J. M., Alatalo, S. L., Suominen, H., Cheng, S., Janckila, A. J., and Vaananen, H. K. (2000). Tartrate-resistant acid phosphatase 5b: a novel serum marker of bone resorption. J. Bone Miner. Res. 15, 1337–1345. 39. Seibel, M. J., Woitge, H. W., Pecherstorfer, M., Karmatschek, M., Horn, E., Ludwig, H., Armbruster, F. P., and Ziegler, R. (1996).

40.

41. 42.

43.

44.

45.

46.

47.

48.

49.

50.

51.

52. 53.

54.

55.

Serum immunoreactive bone sialoprotein as a new marker of bone turnover in metabolic and malignant bone disease. J. Clin. Endocrinol. Metab. 81, 3289–3298. Seibel, M. J., Lang, M., and Geilenkeuser, W. J. (2001). Interlaboratory variation of biochemical markers of bone turnover. Clin. Chem. 47, 1443–1445. Body, J. J., Coleman, R. E., and Piccart, M. (1996). Use of bisphosphonates in cancer patients. Cancer Treat. Rev. 22, 265–287. Stewart, A. F., Vignery, A., Silverglate, A., Ravin, N. D., Livolsi, V., Broadus, A. E., and Baron, R. (1982). Quantitative bone histomorphometry in humoral hypercalcemia of malignancy: uncoupling of bone cell activity. J. Clin. Endocrinol. Metab. 55, 219–227. Dumon, J. C., Wantier, H., Mathieu, F., Mantia, M., and Body, J. J. (1996). Technical and clinical validation of a new immunoradiometric assay for human osteocalcin. Eur. J. Endocrinol. 135, 231–237. Dumon, J. C., and Body, J. J. (1995). Circulating osteocalcin in hypercalcemic cancer patients – A comparative evaluation of two immunoassays. Diagnostic Oncol. 4, 170–173. Nakayama, K., Fukumoto, S., Takeda, S., Takeuchi, Y., Ishikawa, T., Miura, M., Hata, K., Hane, M., Tamura, Y., Tanaka, Y. Kitaoka, M., Obara, T., Ogata, E., and Matsumoto, T. (1996). Differences in bone and vitamin D metabolism between primary hyperparathyroidism and malignancy-associated hypercalcemia. J. Clin. Endocrinol. Metab. 81, 607–611. Body, J. J., Dumon, J. C., Blocklet, D., and Darte, C. (1997). The bone isoenzyme of alkaline phosphatase in hypercalcaemic cancer patients. Eur. J. Cancer 33, 1578–1582. Sato, K., Yamakawa, Y., Shizume, K., Satoh, T., Nohtomi, K., Demura, H., Akatsu, T., Nagata, N., Kasahara, T., Ohkawa, H. and Ohsumi, K. (1993). Passive immunization with anti-parathyroid hormone-related protein monoclonal antibody markedly prolongs survival time of hypercalcemic nude mice bearing transplanted human PTHrP-producing tumors. J. Bone Miner. Res. 8, 849–860. Endres, D. B., Villanueva, R., Sharp Jr, C. F., and Singer, F. R. (1991). Immunochemiluminometric and immunoradiometric determinations of intact and total immunoreactive parathyrin: performance in the differential diagnosis of hypercalcemia and hypoparathyroidism. Clin. Chem. 37, 162–168. Walls, J., Ratcliffe, W. A., Howell, A., and Bundred, N. J. (1994). Parathyroid hormone and parathyroid hormone-related protein in the investigation of hypercalcaemia in two hospital populations. Clin. Endocrinol. 41, 407–413. Body, J. J., Dumon, J. C., Thirion, M., and Cleeren, A. (1993). Circulating PTHrP concentrations in tumor-induced hypercalcemia: influence on the response to bisphosphonate and changes after therapy. J. Bone Miner. Res. 8, 701–706. Grill, V., Ho, P., Body, J. J., Johanson, N., Lee, S. C., Kukreja, S. C., Moseley, J. M. and Martin, T. J. (1991). Parathyroid hormone-related protein: elevated levels in both humoral hypercalcemia of malignancy and hypercalcemia complicating metastatic breast cancer. J. Clin. Endocrinol. Metab. 73, 1309–1315. Burtis, W. J. (1992). Parathyroid hormone-related protein: structure, function and measurement. Clin. Chem. 38, 2171–2183. Body, J. J., Dumon, J. C., Seraj, F., and Cleeren, A. (1992). Recovery of parathyroid hormone secretion during correction of tumor-associated hypercalcemia. J. Clin. Endocrinol. Metab. 74, 1385–1388. Berruti, A., Dogliotti, L., Gorzegno, G., Torta, M., Tampellini, M., Tucci, M., Cerutti, S., Frezet, M. M., Stivanello, M., Sacchetto, G., and Angeli, A. (1999). Differential patterns of bone turnover in relation to bone pain and disease extent in bone in cancer patients with skeletal metastases. Clin. Chem. 45, 1240–1247. Woitge, H. W., Seibel, M. J., and Ziegler, R. (1996). Comparison of total and bone-specific alkaline phosphatase in patients with nonskeletal disorders or metabolic bone diseases. Clin. Chem. 42, 1796–1804.

808 56. Marchei, P., Santini, D., Bianco, V., Chiodini, S., Reale, M. G., Simeoni, F., Marchei, G. G., and Vecchione, A. (1995). Serum Ostase in the follow-up of breast cancer patients. Anticancer Res. 15, 2217–2222. 57. Van Hoof, V. O., Van Oosterom, A. T., Lepoutre, L. G., and De Broe, M. E. (1992). Alkaline phosphatase isoenzyme patterns in malignant disease. Clin. Chem. 38, 2546–2551. 58. Zaninotto, M., Secchiero, S., and Rubin, D. (1995). Serum bone alkaline phosphatase in the follow-up of skeletal metastases. Anticanccer Res. 15, 2223–2228. 59. Berruti, A., Piovesan, A., Torta, M., Raucci, C. A., Gorzegno, G., Paccotti, P., Dogliotti, L., and Angeli, A. (1996). Biochemical evaluation of bone turnover in cancer patients with bone metastases: relationship with radiograph appearances and disease extension. Br. J. Cancer 73, 1581–1587. 60. Cooper, E. H., Whelan, P., and Purves, D. (1994). Bone alkaline phosphatase and prostate-specific antigen in the monitoring of prostate cancer. Prostate 25, 236–242. 61. Garnero, P., Buchs, N., Zekri, J., Rizzoli, R., Coleman, R. E., and Delmas, P. D. (2000). Markers of bone turnover for the management of patients with bone metastases from prostate cancer. Br. J. Cancer 82, 858–864. 62. Wolff, J. M., Ittel, T., Boeckmann, W., Reinike, T., Habib, F. K., and Jakse, G. (1996). Skeletal alkaline phosphatase in the metastatic workup of patients with prostate cancer. Eur. Urol. 30, 302–306. 63. Lorente, J. A., Morote, J., Raventos, C., Encabo, G., and Valenzuela, H. (1996). Clinical efficacy of bone alkaline phosphatase and prostate specific antigen in the diagnosis of bone metastasis in prostate cancer. J. Urology 155, 1348–1351. 64. Woitge, H. W., Horn, E., Keck, A. V., Auler, B., Seibel, M. J., and Pecherstorfer, M. (2001). Biochemical markers of bone formation in patients with plasma cell dyscrasias and benign osteoporosis. Clin. Chem. 47, 686–693. 65. Alexandrakis, M. G., Passam, F. H., Malliaraki, N., Katachanakis, C., Kyriakou, D. S., and Margioris, A. N. (2002). Evaluation of bone disease in multiple myeloma: a correlation between biochemical markers of bone metabolism and other clinical parameters in untreated multiple myeloma patients. Clin. Chim. Acta. 325, 51–57. 66. Body, J. J., Cleeren, A., Pot, M., and Borkowski, A. (1986). Serum osteocalcin (BGP) in tumor-associated hypercalcemia. J. Bone Miner. Res. 1, 523–527. 67. Kamby, C., Egsmose, C., Soletormos, G., Dombernowsky, P. (1993). The diagnostic and prognostic value of serum bone GLA protein (osteocalcin) in patients with recurrent breast cancer. Scand. J. Clin. Lab. Invest. 53, 439–446. 68. Shih, W. J., Wierzbinski, B., Collins, J., Magoun, S., Chen, I. W., and Ryo, U. Y. (1990). Serum osteocalcin measurements in prostate carcinoma patients with skeletal deposits shown by bone scintigram: comparison with serum PSA/PAP measurements. J. Nucl. Med. 31, 1486–1489. 69. Coleman, R. E., Mashiter, G., Fogelman, I., and Rubens, R. D. (1988). Osteocalcin: a marker of metastatic bone disease. Eur. J. Cancer 24, 1211–1217. 70. Bataille, R., Delmas, P. D., Chappard, D., and Sany, J. (1990). Abnormal serum bone Gla protein levels in multiple myeloma. Cancer 66, 167–172. 71. Mejjad, O., Le Loët, X., Basuyau, J. P., Ménard, J. F., Jego, P., Grisot, C., Darangon, A., Grosbois, B., Euller-Ziegler, L., Monconduit, M., le Groupe d’Etude et de Recherche sur le Myélome (GERM) (1996). Osteocalcin is not a marker of progress in multiple myeloma. Eur. J. Hematol. 56, 30–34. 72. Burlina, A., Rubin, D., Secchiero, S., Sciacovelli, L., Zaninotto, M., and Plebani, M. (1994). Monitoring skeletal cancer metastases with the bone isoenzyme of tissue unspecific alkaline phosphatase. Clin. Chim. Acta 226, 151–158.

JEAN-JACQUES BODY 73. Ebeling, P. R., Peterson, J. M., and Riggs, B. L. (1992). Utility of type I procollagen propeptide assays for assessing abnormalities in metabolic bone diseases. J. Bone Miner. Res. 7, 1243–1250. 74. Nakashima, J., Sumitomo, M., Miyajima, A., Jitsukawa, S., Saito, S., Tachibana, M., and Murai, M. (1997). The value of serum carboxyterminal propeptide of type 1 procollagen in predicting bone metastases in prostate cancer. J. Urol. 157, 1736–1739. 75. Brown, J. M., Corey, E., Lee, Z. D., True, L. D., Yun, T. J., Tondravi, M., and Vessella, R. L. (2001). Osteoprotegerin and rank ligand expression in prostate cancer. Urology 57, 611–616. 76. Jung, K., Stephan, C., Semjonow, A., Lein, M., Schnorr, D., and Loening, S. A. (2003). Serum osteoprotegerin and receptor activator of nuclear factor-kappa B ligand as indicators of disturbed osteoclastogenesis in patients with prostate cancer. J. Urol. 170, 2302–2305. 77. Jung, K., Lein, M., Stephan, C., Von Hosslin, K., Semjonow, A., Sinha, P., Loening, S. A., and Schnorr, D. (2004). Comparison of 10 serum bone turnover markers in prostate carcinoma patients with bone metastatic spread: diagnostic and prognostic implications. Int. J. Cancer 111, 783–791. 78. Vinholes, J., Coleman, R., and Eastell, R. (1996). Effects of bone metastases on bone metabolism: implications for diagnosis, imaging and assessment of response to cancer treatment. Cancer Treat. Rev. 22, 289–331. 79. Hou, M. F., Lin, S. B., Yuan, S. S., Tsai, L. Y., Tsai, S. M., Hsieh, J. S., and Huang, T. J. (2003). Diagnostic value of urine deoxypyridinoline for detecting bone metastases in breast cancer patients. Ann. Clin. Lab. Sci. 33, 55–61. 80. Lipton, A., Demers, L., Daniloff, Y., Curley, E., Hamilton, C., Harvey, H., Witters, L., Seaman, J., Van der Giessen, R., and Seyedin, S. (1993). Increased urinary excretion of pyridinium cross-links in cancer patients. Clin. Chem. 39, 614–618. 81. Demers, L. M. (1999). Biochemical markers in the management of patients with metastatic bone disease. Clin. Chem. 45, 1131–1132. 82. Demers, L. M., Costa, L., Chinchilli, V. M., Gaydos, L., Curley, E., and Lipton, A. (1995). Biochemical markers of bone turnover in patients with metastatic bone disease. Clin. Chem. 41, 1489–1494. 83. Lipton, A., Demers, L., Curley, E., Chinchilli, V., Gaydos, L., Hortobagyi, G., Theriault, R., Clemens, D., Costa, L., Seaman, J., and Knight, R. (1998). Markers of bone resorption in patients treated with pamidronate. Eur. J. Cancer 34, 2021–2026. 84. Body, J. J., Dumon, J. C., Gineyts, E., and Delmas, P. D. (1997). Comparative evaluation of markers of bone resorption in patients with breast cancer-induced osteolysis before and after bisphosphonate therapy. Br. J. Cancer 75, 408–412. 85. Ju, H. S. J., Leung, S., Brown, B., Stringer, M. A., Leigh, S., Scherrer, C., Shepard, K., Jenkins, D., Knudsen, J., and Cannon, R. (1997). Comparison of analytical performance and biological variability of three bone resorption assays. Clin. Chem. 43, 1570–1576. 86. Robins, S. P., Woitge, H., Hesley, R., Ju, J., Seyedin, S., and Seibel, M. J. (1994). Direct, enzyme-linked immunoassay for urinary deoxypyridinoline as a specific marker for measuring bone resorption. J. Bone Miner. Res. 9, 1643–1649. 87. Nguyen-Pamart, M., Bonneterre, J., and Hecquet, B. (1995). Urinary excretion of deoxypyridinoline in patients with breast cancer. Anticancer Res. 15, 1601–1604. 88. Halleen, J. M., Alatalo, S. L., Janckila, A. J., Woitge, H. W., Seibel, M. J., and Vaananen, H. K. (2001). Serum tartrate-resistant acid phosphatase 5b is a specific and sensitive marker of bone resorption. Clin. Chem. 47, 597–600. 89. Bombardieri, E., Martinetti, A., Miceli, R., Mariani, L., Castellani, M. R., and Seregni, E. (1997). Can bone metabolism markers be adopted as an alternative to scintigraphic imaging in monitoring bone metastases from breast cancer? Eur. J. Nucl. Med. 24, 1349–1355. 90. Houze, P., Bellik, B., Extra, J. M., Bouro, F., and Bousquet, B. (1999). Urinary carboxyterminal telopeptide of collagen I as a potential marker

Chapter 48 Metastatic Bone Disease

91.

92.

93.

94.

95.

96.

97.

98.

99.

100.

101.

102.

103.

104.

105.

106.

of bone metastases chemotherapy monitoring in breast cancer. Clin. Chim. Acta 281, 77–88. Ali, S. M., Demers, L. M., Leitzel, K., Harvey, H. A., Clemens, D., Mallinak, N., Engle, L., Chinchilli, V., Costa, L., Brady, C., Seaman, J., and Lipton, A. (2004). Baseline serum NTx levels are prognostic in metastatic breast cancer patients with bone-only metastasis. Ann. Oncol. 15, 455–459. Miyamoto, K. K., McSherry, S. A., Robins, S. P., Besterman, J. M., and Mohler, J. L. (1994). Collagen cross-link metabolites in urine as markers of bone metastases in prostatic carcinoma. J. Urol. 151, 909–913. Kylmälä, T., Tammela, T. L. J., Risteli, L., Risteli, J., Kontturi, M., and Elomaa, I. (1995). Type I collagen degradation product (ICTP) gives information about the nature of bone metastases and has prognostic value in prostate cancer. Br. J. Cancer 71, 1061–1064. Berruti, A., Dogliotti, L., Bitossi, R., Fasolis, G., Gorzegno, G., Bellina, M., Torta, M., Porpiglia, F., Fontana, D., and Angeli, A. (2000). Incidence of skeletal complications in patients with bone metastatic prostate cancer and hormone refractory disease: predictive role of bone resorption and formation markers evaluated at baseline. J. Urol. 164, 1248–1253. Dawson, N. A. (2002). Bisphosphonates: their evolving role in the management of prostate cancer-related bone disease. Curr. Opin. Urology 12, 413–418. Ross, R. W., and Small, E. J. (2002). Osteoporosis in men treated with androgen deprivation therapy for prostate cancer. J. Urol. 167, 1952–1956. Terpos, E., de la Fuente, J., Szydlo, R., Hatjiharissi, E., Biniou, N., Meletis, J., Yataganas, X., Goldman, J. M., and Rahemtulla, A. (2003). Tartrate-resistant acid phosphatase isoform 5b: a novel serum marker for monitoring bone disease in multiple myeloma. Int. J. Cancer 106, 455–457. Withold, W., Armbruster, F. P., Karmatschek, M., and Reinauer, H. (1997). Bone sialoprotein in serum of patients with malignant bone diseases. Clin. Chem. 43, 85–91. Woitge, H. W., Pecherstorfer, M., Horn, E., Keck, A. V., Diel, I. J., Bayer, P., Ludwig, H., Ziegler, R., and Seibel, M. J. (2001). Serum bone sialoprotein as a marker of tumour burden and neoplastic bone involvement and as a prognostic factor in multiple myeloma. Br. J. Cancer 84, 344–351. Coleman, R. E., Whitaker, K. D., Moss, D. W., Mashiter, G., Fogelman, I., and Rubens, R. D. (1988). Biochemical monitoring predicts response in bone to systemic treatment. Br. J. Cancer 58, 205–210. Cooper, E. H., Forbes, M. A., Hancock, A. K., Parker, D., and Laurence, V. (1992). Serum bone alkaline phosphatase and CA 549 in breast cancer with bone metastases. Biomed. Pharmacother. 46, 31–36. Luftner, D., Richter, A., Geppert, R., Wernecke, K. D., and Possinger, K. (2003). Normalisation of biochemical markers of bone formation correlates with clinical benefit from therapy in metastatic breast cancer. Anticancer Res. 23, 1017–1026. Koizumi, M., Maeda, H., Yoshimura, K., Yamauchi, T., Kawai, T., and Ogata, E. (1997). Dissociation of bone formation markers in bone metastasis of prostate cancer. Br. J. Cancer 75, 1601–1604. Williams, A. T., Shearer, M. J., Oyeyi, J., Aitchison, R. G., Newland, A. C., and Schey, S. A. (1992). Serum osteocalcin in the management of myeloma. Eur. J. Cancer 29A, 140–142. Downey, S., Walls, J., Ratcliffe, W. A., Assiri, A. M. A., Howell, A., Eastell, R., and Bundred, N. O. (1994). Pyridinoline excretion can monitor bone metastases in breast cancer. Breast Cancer Res. Treat. 32, 83. Walls, J., Assiri, A., Howell, A., Rogers, E., Ratcliffe, W. A., Eastell, R., and Bundred, N. J. (1999). Measurement of urinary collagen crosslinks indicate response to therapy in patients with breast cancer and bone metastases. Br. J. Cancer 80, 1265–1270.

809 107. Vinholes, J., Coleman, R., Lacombe, D., Rose, C., Tubiana-Hulin, M., Bastit, P., Wildiers, J., Michel, J., Leonard, R., Nortier, J., Mignolet, F., and Ford, J. (1999). Assessment of bone response to systemic therapy in an EORTC trial: preliminary experience with the use of collagen cross-link excretion. European Organization for Research and Treatment of Cancer. Br. J. Cancer 80, 221–228. 108. Coleman, R. E. (1998). Monitoring of bone metastases. Eur. J. Cancer 34, 252–259. 109. Vinholes, J., Coleman, R., Lacombe, D., Mignolet, F., Rose, C., Leonard, R., Nortier, J., Tubiana-Hulin, M., EORTC Breast Group. (1996). Assessment of bone response to systemic therapy in an EORTC trial of oral pamidronate. Eur. J. Cancer 32A (suppl. 2), 50. 110. Costa, L., Demers, L. M., Gouveia-Oliveira, A., Schaller, J., Costa, E. B., de Moura, M. C., and Lipton, A. (2002). Prospective evaluation of the peptide-bound collagen type I cross-links N-telopeptide and C-telopeptide in predicting bone metastases status. J. Clin. Oncol. 20, 850–856. 111. Brown, J. E., Thomson, C. S., Ellis, S. P., Gutcher, S. A., Purohit, O. P., and Coleman, R. E. (2003). Bone resorption predicts for skeletal complications in metastatic bone disease. Br. J. Cancer 89, 2031–2037. 112. Brown, J. E., Cook, R. J., Major, P., Lipton, A., Saad, F., Smith, M., Lee, K. A., Zheng, M., Hei, Y. J., and Coleman, R. E. (2005). Bone turnover markers as predictors of skeletal complications in prostate cancer, lung cancer, and other solid tumors. J. Natl Cancer Inst. 97, 59–69. 113. Clamp, A., Danson, S., Nguyen, H., Cole, D., and Clemons, M. (2004). Assessment of therapeutic response in patients with metastatic bone disease. Lancet Oncol. 5, 607–616. 114. Ikeda, I., Miura, T., and Kondo, I. (1996). Pyridinium cross-links as markers of bone metatases in patients with prostate cancer. Br. J. Urol. 77, 102–106. 115. Elooma, I., Virkkunen, P., Risteli, L., and Risteli, J. (1992). Serum concentration of the cross-linked carboxyterminal telopeptide of type I collagen (ICTP) is a useful prognostic indicator in multiple myeloma. Br. J. Cancer 66, 337–341. 116. Kylmälä, T., Tammela, T., Risteli, L., Risteli, J., Taube, T., and Elomaa, I. (1993). Evaluation of the effect of oral clodronate on skeletal metastases with type 1 collagen metabolites. A controlled trial of the Finnish Prostate Cancer Group. Eur. J. Cancer 29A, 821–825. 117. Abildgaard, N., Brixen, K., Eriksen, E. F., Kristensen, J. E., Nielsen, J. L., and Heickendorff, L. (2004). Sequential analysis of biochemical markers of bone resorption and bone densitometry in multiple myeloma. Haematologica 89, 567–577. 118. Body, J. J. (2003). Effectiveness and cost of bisphosphonate therapy in tumor bone disease. Cancer 97, 859–865. 119. Body, J. J. (2003). Zoledronic acid: an advance in tumour bone disease and a new hope for osteoporosis. Expert Opin. Pharmacotherapy 4, 567–580. 120. Vinholes, J., Guo, C. Y., Purohit, O. P., Eastell, R., Coleman, R. E. (1997). Randomised double-blind comparison of pamidronate or clodronate for hypercalcemia of malignancy: effects on bone metabolism markers. J. Clin. Oncol. 15, 131–138. 121. Rubinacci, A., Melzi, R., Zampino, M., Soldarini, A., and Villa, I. (1999). Total and free deoxypyridinoline after acute osteoclast activity inhibition. Clin. Chem. 45, 1510–1516. 122. Vinholes, J. J. F., Purohit, O. P., Abbey, M. E., Eastell, R., and Coleman, R. E. (1997). Relationships between biochemical and symptomatic response in a double-blind randomised trial of pamidronate for metastatic bone disease. Ann. Oncol. 8, 1243–1250. 123. Engler, H., Koeberle, D., Thuerlimann, B., Senn, H. J., and Riesen, W. F. (1998). Diagnostic and prognostic value of biochemical markers in malignant bone disease: a prospective study on the effect of

810 bisphosphonate on pain intensity and progression of malignant bone disease. Clin. Chem. Lab. Med. 36, 879–885. 124. Menssen, H. D., Sakalova, A., Fontana, A., Herrmann, Z., Boewer, C., Facon, T., Lichinitser, M. R., Singer, C. R., Euller-Ziegler, L., Wetterwald, M., Fiere, D., Hrubisko, M., Thiel, E., and Delmas, P. D. (2002). Effects of long-term intravenous ibandronate therapy on skeletal-related events, survival, and bone resorption markers in patients with advanced multiple myeloma. J. Clin. Oncol. 20, 2353–2359. 125. Powles, T., Paterson, S., Kanis, J. A., McCloskey, E., Ashley, S., Tidy, A., Rosenqvist, K., Smith, I., Ottestad, L., Legault, S., Pajunen, M., Nevantaus, A., Mannisto, E., Suovuori, A., Atula, S., Nevalainen, J., Pylkkanen, L. (2002). Randomized, placebo-controlled trial of clodronate in patients with primary operable breast cancer. J. Clin. Oncol. 20, 3219–3224. 126. Powles, T., Paterson, A., McCloskey, E., Kurkilahti, M., Kanis, J. (2004). Oral clodronate for adjuvant treatment of operable breast cancer: results of a randomized, double-blind, placebo-controlled multicenter trial. Proc ASCO 23, 9(Abstract 528).

JEAN-JACQUES BODY 127. Bellahcène, A., Kroll, M., Liebens, F., and Castronovo, V. (1996). Bone sialoprotein expression in primary human breast cancer is associated with bone metastases development. J. Bone Miner. Res. 11, 665–670. 128. Bellahcène, A., Menard, S., Bufalino, R., Moreau, L., and Castronovo, V. (1996). Expression of bone sialoprotein in primary human breast cancer is associated with poor survival. Int. J. Cancer 69, 350–353. 129. Diel, I. J., Solomayer, E. F., Seibel, M. J., Pfeilschifter, J., Maisenbacher, H., Gollan, C., Pecherstorfer, M., Conradi, R., Kehr, G., Boehm, E., Armbruster, F. P., and Bastert, G. (1999). Serum bone sialoprotein in patients with primary breast cancer is a prognostic marker for subsequent bone metastasis. Clin. Cancer. Res. 5, 3914–3919. 130. Seibel, M. J., Koeller, M., Van der Velden, B., and Diel, I. (2002). Markers of bone turnover do not predict bone metastases in breast cancer. Clin. Lab. 48, 583–588.

Chapter 49

Rare Bone Diseases Michael P. Whyte, M.D.

Center for Metabolic Bone Disease and Molecular Research, Shriners Hospitals for Children; and Division of Bone and Mineral Diseases, Washington University School of Medicine At Barnes-Jewish Hospital; St. Louis, Missouri

I. Introduction II. Osteopenia III. Osteosclerosis and Hyperostosis

IV. Ectopic Calcification V. Other Disorders References

I. INTRODUCTION

This chapter summarizes information concerning genetic as well as local and systemic factors involved in the etiology and pathogenesis of rare bone diseases where data from assays of biochemical markers of skeletal remodeling are accumulating. For such disorders, bone turnover assessed biochemically can shed light on the nature of the skeletal pertubation and help to design, evaluate, and monitor treatment. Here, however, systemic or focal derangements in bone growth, modeling, and remodeling and cartilage formation and degradation can be occurring and lead to generalized or regional osteopenia (bone density below the normal) or increases in skeletal mass. Biochemical markers of skeletal formation or resorption provide summative values reflecting the impact of a global disturbance or a partly localized lesion. Hence, the type and extent of the pathologic process in individual patients merits consideration when interpreting results from such assays. This is best illustrated by the common focal skeletal disorder, Paget’s bone disease [3]. Such reflection has typically not been applied to rare bone disorders – some of which are discussed below grouped as osteopenia, osteosclerosis and hyperostosis, ectopic calcification, and “other.”

Rare bone diseases are often challenging for physicians. Lack of familiarity with their myriad clinical, radiological, biochemical, and histopathological features will make diagnosing difficult. Little personal or cumulative experience with their management will render treatment uncertain or ineffective. In fact, new skeletal disorders continue to be discovered [1, 2]. Understanding rare bone diseases is important for several reasons. First, there are many such entities, and their cumulative prevalence is not negligible. Second, although a few are mere radiographic curiosities, most pose significant lifelong problems and some have useful medical and surgical treatments. Third, a sizeable number are heritable, representing an “experiment-of-nature,” so that more than one individual or future pregnancies in a family can be at risk. Fourth, elucidation of their etiology and pathogenesis offers insight into skeletal development and the dynamics of bone and cartilage metabolism [1, 2]. Although a specific disorder may be encountered just once in a medical career, patients with rare bone diseases must be appreciated and cared for and are always memorable. Dynamics of Bone and Cartilage Metabolism

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II. OSTEOPENIA Among the rare forms of osteopenia are osteogenesis imperfecta, idiopathic juvenile osteoporosis, heritable disorders of RANKL/OPG/RANK/NF-κB signaling, and cleidocranial dysplasia.

A. Osteogenesis Imperfecta Osteogenesis imperfecta [OI] is a heritable disorder of connective tissue where the fundamental pathogenesis in essentially all patients is a quantitative and often qualitative abnormality in the biosynthesis of the most abundant protein in bone, type I collagen [4]. Consequently, the clinical hallmark of OI is generalized osteopenia associated with recurrent fracture and skeletal deformity [5]. However, type I collagen is also found in dentin, ligaments, skin, and sclerae and patients with OI often have additional clinical problems involving the teeth (dentinogenesis imperfecta) and other tissues that normally contain this major structural protein [4, 5]. Nevertheless, the severity of OI is extremely variable and ranges from stillbirth to perhaps lifelong absence of symptoms. The classification scheme proposed by Sillence in 1979 for OI, according to clinical and radiographic features and apparent patterns of inheritance, provided a framework for prognostication and served as a foundation for further biochemical/genetic investigation [6] (Table I). However, this nosology has been largely superseded by insight from recombinant DNA studies [4] showing numerous different mutations in the OI patient population involving either of the two genes that encode the α1 or α2 chains of Table I. OI type I

II

III

IV

the type I collagen heterotrimer, and autosomal dominant inheritance for nearly all affected families [7]. The nosology of Sillence describes four major types of OI [6]. Type I OI features relatively mild osteopenia and infrequent fractures (deformity is uncommon or not severe), sclerae with obvious bluish discoloration that is especially striking during childhood (Color Plate A), and deafness that typically manifests during early adulthood (30% incidence). Stature is usually normal. Type I OI has been subclassified into forms I-A and I-B, depending upon the absence or (more rarely) the presence, respectively, of dentinogenesis imperfecta (Color Plate B). Type II OI is manifest at birth with profound osteopenia and is typically fatal within a few weeks due to respiratory compromise from thoracic insufficiency. Affected newborns are often premature, small for gestational age, and have short and bowed limbs, numerous fractures, markedly soft skulls, and small chests. Type III OI is distinguished by progressive skeletal deformity and short stature acquired during childhood that result from recurrent fractures as well as from fragmentation of growth plates due to moderately severe osteopenia (Fig. 1). Type IV OI was once considered rare, but increasingly is used imprecisely to explain multigenerational osteoporosis not classified as type I or III OI and not understood at the molecular/biochemical level. The sclerae are typically white, but characteristics of OI can include skeletal deformity, dentinogenesis imperfecta, and hearing loss. Within the OI population worldwide, >800 distinctive defects have been identified in either of the genes that encode the two large proteins that intertwine to become

Clinical Heterogeneity and Biochemical Defects in Osteogenesis Imperfecta

Clinical features Normal stature, little or no deformity; blue sclerae, hearing loss in about 50 percent of individuals; dentinogenesis imperfecta rare and may distinguish a subset Lethal in the perinatal period, minimal calvarial mineralization, beaded ribs, compressed femurs, marked long bone deformity, platyspondyly Progressively deforming bones, usually with moderate deformity at birth; scleras variable in hue, often lighten with age; dentinogenesis imperfecta common, hearing loss common; stature very short Normal sclerae, mild to moderate bone deformity; variable short stature; dentinogenesis common; hearing loss occurs in some

Reproduced with permission from Byers [4]. *AD, autosomal dominant; AR, autosomal recessive.

Inheritance*

Biochemical defects

AD

Decreased production of type I procollagen; substitution for residue other than glycine in triple helix of α1(I)

AD (new mutation) AR (rare)

Rearrangements in the COL1A1 and COL1A2 genes; substitutions for glycyl residues in the triple-helical domain of the α1(I) α2(I) chain; small deletion in α2(I) on the background of a null allele Point mutation in the α1(I) or α2(I) chain Frameshift mutation that prevents incorporation of proα2(I) into molecules (non-collagenous defects)

AD AR

AD

Point mutations in the α2(I) chain; rarely, point mutations in the α1(I) chain; small deletions in the α2(I) chain

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Chapter 49 Rare Bone Diseases

Color Plate A

This 14-year-old girl has remarkably blue sclera characteristic of type I osteogenesis imperfecta (photographed digitally with

a built-in flash).

the type I collagen heterotrimer [7]. The genetic basis for nearly all genuine cases of OI is a mutation involving one allele of either the COL1A1 or the COL1A2 genes [4, 7]. These genes direct the biosynthesis of the proα1(I) and the proα2(I) proteins that undergo post-translational modification and combine in a 2:1 ratio, respectively, to form ultimately the important triple-helical (structural) domain of the type I collagen fibril [4] (see Chapters 1 and 2). Diminished synthesis of type I collagen, often also associated with a distorted heterotrimer, is the principal

Color Plate B osteogenesis imperfecta.

biochemical abnormality in OI [4]. When a flaw in one of the four alleles leads to functional loss of approximately 50% of either the proα1(I) or the proα2(I) chains, there is mild OI [4, 5]. Severe OI is typically explained by either a deletion of a single exon or by a point mutation, leading to a substitution of one of the many critical glycyl residues in the triple-helical domain of a proα1(I) or proα2(I) chain [4, 5]. Consequently, delayed physical association of these chains can also cause their posttranslational overmodification, further distorting the

The translucent teeth of dentinogenesis imperfecta encountered in a child with

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Figure 1 Osteogenesis imperfecta. Progressively deforming (type III) OI in this 4-year-old girl manifests with osteopenia and fragmentation of the distal growth plate of her left femur forming characteristic “popcorn calcifications” (arrows).

important triple-helical region. Subsequently, most heterotrimer molecules form defectively, and dominantnegative effects due to a single abnormal allele become a key heritable and pathogenetic factor. OI represents the archetypical disorder in which one mutated allele encoding a complex structural protein causes a dominantly inherited disease. The large size of the COL1A1 and COL1A2 genes partly explains why the great majority of affected families differ due to “private mutations” and how the clinical severity of OI can be so remarkably diverse [4, 7]. In fact, approximately one-third of patients with type I OI reflect new dominant mutations, and the estimated 6% recurrence risk for siblings with type II OI seems to be explained by gonadal mosaicism [4]. Bone histology in OI often reveals an abnormal skeletal matrix, especially in severely affected patients. Irregular architecture of the fibrillar network of collagen is a characteristic feature; polarized-light microscopy shows disorganized “woven” bone where there should be predominantly lamellar osseous tissue. Furthermore, the diameter of single collagen fibers can be either reduced [8] or increased [9]. Early histological studies of the OI skeleton were interpreted as showing low bone turnover [10, 11]. However, in 1983, Baron and colleagues [12] reported dynamic histomorphometry of iliac crest specimens from nine children with mild (“tarda”) OI, suggesting increased skeletal remodeling, although bone formation by individual osteoblasts was decreased. Here, the defect

MICHAEL P. WHYTE

in matrix synthesis by single osteoblasts seemed to be partly compensated for by increased numbers of these cells. As a remnant of this pathologic process, many osteocytes became embedded in cortical bone [12]. In 1984, Ste-Marie and colleagues [11] also described many osteoblasts working with decreased efficiency. In 1991, Vetter and associates showed that apatite crystals were relatively small in OI bone [13], and that non-collagenous proteins in the matrix were also altered [14]. This was confirmed by additional studies [8, 13–17]. In 1992, Fedarko et al. [8] reported reduced amounts of several glycoproteins including osteonectin, biglycan, and decorin. In 1995, these investigators described elevations in thrombospondin, fibronectin, and hyaluronan [18]. OI bone cells in vitro have also revealed important findings. In 1992, Mörike et al. [19] reported that osteoblast-like cells from all clinical forms of OI produce low extracellular levels of type I collagen irrespective of the absence or (more frequently) the presence of structural abnormalities within the heterotrimer itself. However, significant differences occurred among the Sillence types of OI. Diminished production of type I procollagen (versus aberrant processing, secretion, and pericellular accumulation) distinguished type I OI from types III and IV, respectively [19]. Types III and IV featured intracellular retention of the nascent protein. In 1993, Mörike and colleagues [19] also showed that osteoblast-like cells in OI nevertheless correctly express markers of osteoblast activity (alkaline phosphatase, 1,25-dihydroxyvitamin D3-stimulated osteocalcin, and PTH-induced cAMP). Hence, osteoblast differentiation is not altered in OI [20]. In 1995, overhydroxylation of lysyl and prolyl residues in type I collagen was demonstrated using OI patient fibroblasts in culture, and the degree of this post-translational overmodification correlated with the severity of the clinical phenotype [21]. Despite these quantitative and qualitative defects of osseous tissue in OI, routine biochemical parameters of mineral homeostasis are typically unremarkable in patients except for hypercalciuria that was documented in 36% of children with OI followed at Shriners Hospital for Children, St. Louis, Missouri [22]. The magnitude of the hypercalciuria in this pediatric OI population correlated with the clinical severity of their skeletal disease [22], but was not associated with renal dysfunction [23] and seemed to stem from poor utilization of dietary calcium due to impaired skeletal growth. In 1979, Neilsen et al. [24] reported normal serum calcitonin levels in OI patients. In 1993, Marini and coworkers [25] found no evidence of growth hormone deficiency in children with OI, although some patients seemed to have blunted serum IGF-1 values after growth hormone challenge. Since 1969,

Chapter 49 Rare Bone Diseases

increased urinary levels of inorganic pyrophosphate (PPi) have been documented in OI patients, but they do not reflect the disorder’s severity or change with magnesium oxide supplementation [26]. Among the rare bone diseases, biochemical markers of skeletal turnover have been investigated most thoroughly in OI. Elevations in some patients of serum alkaline phosphatase (ALP) activity and urinary levels of hydroxyproline (OHP) were documented repeatedly [10, 27]. Occasionally, serum ALP activity is low in type II OI [28]. In 1986, Castells et al. [29] reported that plasma osteocalcin levels were increased in OI. However, the following year Rico [30] described low osteocalcin levels. Furthermore, in 1991, Rico and colleagues [31] observed that serum tartrate-resistant acid phosphatase (TRAP) activity was reduced in types I and III OI, suggesting quiescent bone turnover in these clinical subtypes consistent with an osteoblast defect. In 1993, Brenner et al. [32] found that serum levels of type I procollagen C-terminal propeptide were also generally low in OI (especially type I). They concluded that this marker would be especially useful for the clinical management and assessment of future therapeutic interventions for OI [32]. They also noted increased serum osteocalcin levels, but only during childhood [32], in keeping with findings using OI bone itself [14]. In 1994, Minisola and coworkers [33] reported a family with type I-A OI whose serum levels of C-terminal propeptide were low, yet other biochemical parameters of skeletal remodeling were unremarkable. In 1994, Brenner et al. [34] discovered generous urinary levels of type I collagen crosslinked N-telopeptide in children and adolescents with OI which correlated positively with urinary calcium excretion. They concluded that the synthesis and turnover of mature, crosslinked type I collagen is disturbed in OI, thereby providing evidence for accelerated bone remodeling [34]. Rauch and colleagues provide more recent evidence for this disturbance [35, 36]. In summary, biochemical markers of bone remodeling in OI generally support histomorphometric studies showing enhanced rates of cancellous bone turnover, despite impaired efficiency of individual osteoblasts. Measurement and correct interpretation of markers of bone remodeling is now becoming especially important for OI because of increasing use of bisphosphonate therapy [36], with potential toxicity causing oversuppression of both skeletal modeling and remodeling [37].

B. Idiopathic Juvenile Osteoporosis Osteoporosis before adult life is usually unmasked when radiographs taken for fracture also show generalized

815 hypomineralization of the skeleton (i.e. osteopenia), but no evidence of rickets or excessive parathyroid hormone (PTH)-mediated bone resorption (e.g. osteitis fibrosa). Corticosteroid therapy is probably the most common cause of acquired osteoporosis in childhood and adolescence, but a considerable number of conditions are associated with this disorder [38, 39]. Idiopathic juvenile osteoporosis (IJO) refers to a rare problem that is usually observed just before puberty, but can also trouble young children, especially those who are growing rapidly [39]. The term was coined by Dent and Friedman in 1965 [40]. Since first described by Schippers in 1938 [41], fewer than 100 cases have been reported in the medical literature [38], but the disorder is more common than this might suggest, and there is concern for many future cases in the United States (see below) [42]. The patient with IJO is typically prepubertal and otherwise healthy [39]. Family history is generally negative in the medical literature, but we find using dual-energy X-ray absorptiometry (DXA) that cases often seem to reflect a familial, and probably heritable, predisposition (unpublished observations). Both sexes are affected with equal prevalence. Usually, symptoms begin with discomfort in the lower back and difficulty walking. Knee and ankle pain and fractures of the lower extremities are typical. Physical examination can be unremarkable or reveal loss of height, limping, thoracolumbar kyphosis or kyphoscoliosis, “pigeon chest” deformity, reduced uppersegment to lower-segment ratio, and long bone deformity. Radiographs show osteopenia and fractures, especially in the spine and metaphyses of the lower limbs in areas of “neo-osseous osteoporosis” (Fig. 2) [43]. Reportedly, IJO often resolves spontaneously with onset of puberty and generally physical deformities are reversible [44, 45]. However, in 1995, in a longitudinal follow-up study of 21 affected children, Smith [44] cautioned that although most patients “substantially or completely recovered,” several developed crippling skeletal deformities leaving them wheelchair-bound with cardiopulmonary compromise as adults. Because spontaneous recovery from IJO does occur, Smith [44] cautioned that it was “impossible to assess the many forms of treatment given.” The clinical onset of IJO, usually just before puberty, may reflect the fact that approximately 25% of peak bone mass is acquired during the 2-year period across peak height velocity [46]. Early on, investigation of a limited number of patients suggested that IJO develops from negative or inappropriately neutral mineral balance at this critical time. Recovery is heralded by improvement in calcium balance [40]. Now, decreasing dietary calcium intake during the adolescent years, especially for girls consuming less than 500 mg daily which can not be

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skin or synthesized by dermal fibroblasts in culture. That same year, however, Pocock and coworkers [52] indicated that quantitative and qualitative abnormalities of type I collagen may characterize a minority of IJO patients who represent a subpopulation of this disorder. There are no characteristic biochemical disturbances of mineral homeostasis in IJO [44]. Case reports mention deficiencies or excesses, including low circulating levels of 1,25(OH)2D or calcitonin, and beneficial responses to treatment with these hormones [53, 54]. However, in a relatively large study, circulating 25(OH)D and 1,25(OH)2D levels were unremarkable [44]. Increased urinary excretion of OHP as well as hypercalcemia and suppressed serum PTH levels have been observed in some patients. Such parathyroid gland suppression could reduce 1,25(OH)2D synthesis and decrease intestinal calcium absorption, thereby contributing to the negative calcium balance.

C. Heritable Disorders of RANKL/OPG/RANK/NF-κB Signaling

Figure 2 Idiopathic juvenile osteoporosis. Osteopenia is present, but there is no evidence of rickets or hyperparathyroidism in this 10-year-old girl. Characteristic “neo-osseous osteoporosis” is apparent in the metaphysis of the proximal left tibia (arrow).

compensated for by enhanced gastrointestinal absorption [47], predicts an increase in IJO. In 2004, the U.S. Surgeon General’s report concerning osteoporosis and bone health addressed this issue [42]. Studies of bone tissue have been reported in IJO. Jowsey and Johnson [45] and Cloutier et al. [48] published histologic and microradiographic evidence of increased skeletal resorption, speculating that excessive dietary phosphorus stimulated PTH-mediated bone resorption. Smith [44] described decreased bone formation from static histomorphometry of iliac crest specimens. Using tetracycline labeling techniques, Evans and colleagues [49] suggested that severe IJO in one boy likely resulted from a reduction in bone formation. In 2002, Rauch et al. described both decreased cancellous bone volume and formation rate [50]. In 1992, Bertelloni et al. [51] reported normal osteoblast function in six IJO patients using a stimulation test that measured serum osteocalcin levels after 1,25(OH)2D3 challenge. In 1995, Smith [44] found that most patients with IJO manifest no abnormality of collagen in their

Identification of the RANKL/OPG/RANK/NF-κB signaling pathway as the foundation for regulating osteoclast formation and action, and thereby skeletal remodeling, followed discovery of a new ligand (RANKL) and two receptors (OPG, RANK) in the tumor necrosis factor (TNF) superfamily [55]. Subsequent studies of genetically altered mice revealed key physiologic roles for these molecules in bone biology. Here, RANKL binds to receptor activator of nuclear factor-κB (RANK) on osteoclast precursor cells to enhance osteoclastogenesis and to osteoclasts to increase their activity and delay apoptosis. Osteoprotegerin (OPG) acts as a “decoy receptor” for RANKL to diminish its effects [55]. Nevertheless, full appreciation of the significance for this pathway in humans derived from clinical characterization of several extremely rare, heritable diseases followed by elucidation of their genetic bases [56]. These remarkable disorders, that primarily involve the skeleton, reflect gene defects leading to constitutive activation of RANK or to deficiency of OPG. In 2000, Hughes and colleagues investigated familial expansile osteolysis (FEO) and identified an activating 18-bp tandem duplication in the gene encoding RANK (TNFRSF11A) in three affected kindreds, and a similar 27-bp duplication causing a familial form of early-onset Paget’s disease of bone (PDB2) in Japan. In 2002, Whyte and coworkers reported that a seemingly unique disorder designated expansile skeletal hyperphosphatasia (ESH) was allelic to FEO and explained by a 15-bp tandem

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duplication in RANK. That same year, Whyte and colleagues discovered homozygous deletion of the gene encoding OPG (TNFRSF11B) as a molecular explanation for idiopathic hyperphosphatasia, also called juvenile Paget’s disease (JPD), in two Navajo patients. Homozygous deactivating mutations in TNFRSF11B from a founder, causing OPG deficiency, account for most, but not all, cases of JPD [56]. Hence, the clinical features of FEO, ESH, and JPD illustrate the crucial role RANKL/OPG/ RANK/NF-κB signaling has for skeletal homeostasis in humans. Each disorder is associated with rapid bone remodeling throughout the skeleton, deafness during early childhood from defects in ossicles, premature loss of adult teeth from “external osteolysis,” and focal osteolytic lesions in appendicular bones that mimic the active phase of PDB. Biochemical markers of bone remodeling are increased, especially in JPD, and indicate accelerated bone turnover due to the generalized as well as the focal skeletal changes. As expected, antiresorptive therapy with bisphosphonates can be effective [56].

In 1992, Moore et al. [57] reported in a preliminary communication that serum ALP activity was low in several patients with CCD consistent with decreased osteoblast function. Subsequently, in 1997, mutations were identified in CBFA1 [58] – the gene that encodes the transcriptional activator of osteoblast differentiation, OSF2/CBFA1 [59]. In 2002, low serum ALP activity (hypophosphatasemia) was verified in a few CCD cases in whom additional biochemical features of hypophosphatasia (deactivation of TNSALP causing deficiency of the tissue-non-specific isoenzyme of ALP) were reportedly also present [60, 61]. We found normal TNSALP substrate levels as well as serum osteocalcin concentrations in two children with CCD due to CBFA1 defects [28]. Other biochemical markers of skeletal remodeling in patients with CCD await investigation.

D. Cleidocranial Dysplasia

Osteosclerosis and hyperostosis refer to trabecular and cortical bone thickening, respectively [62]. Focal or generalized osteosclerosis and/or hyperostosis are caused by many rare (primarily hereditary) dysplastic diseases, as well as by a variety of dietary, metabolic, endocrine, hematologic, infectious, and neoplastic conditions (Table II). Several of the dysplastic disorders are discussed here because they can be especially informative concerning

Cleidocranial dysplasia (CCD) is an autosomal dominant disorder characterized by large cranial fontanelles, aplasia or hypoplasia of the clavicles, delayed skeletal development, and a variety of dental anomalies (Fig. 3A, B) [1]. This clinical phenotype suggests a fundamental defect in ossification.

A Figure 3

III. OSTEOSCLEROSIS AND HYPEROSTOSIS

B

Cleidocranial dysplasia. (A) Open cranial sutures are readily apparent in this 4-year-old boy. (B) At 5 years of age, a chest radiograph reveals characteristic clavicular hypoplasia.

818 Table II.

MICHAEL P. WHYTE Disorders That Cause High Bone Mass*

Dysplasias and dysostoses Autosomal dominate osteosclerosis Central osteosclerosis with ectodermal dysplasia Craniodiaphyseal dysplasia Craniometaphyseal dysplasia Dysosteosclerosis Endosteal hyperostosis (van Buchem disease/sclerosteosis) Frontometaphyseal dysplasia Infantile cortical hyperostosis (Caffey disease) Lenz–Majewski syndrome Melorheostosis Metaphyseal dysplasia (Pyle disease) Mixed-sclerosing bone dystrophy Oculodento-osseous dysplasia Osteodysplasia of Melnick and Needles Osteoectasia with hyperphosphatasia (hyperostosis corticalis) Osteopathia striata Osteopetrosis Osteopoikilosis Pachydermoperiostosis Progressive diaphyseal dysplasia (Engelmann disease) Pycnodysostosis Tubular stenosis (Kenny–Caffey syndrome) Metabolic Carbonic anhydrase II deficiency Fluorosis Heavy metal poisoning Hepatitis C-associated osteosclerosis Hypervitaminosis A, D

Hyper-, hypo-, and pseudohypoparathyroidism Hypophosphatemic osteomalacia Milk-alkali syndrome Renal osteodystrophy X-linked hypophosphatemia Other Axial osteomalacia Diffuse idiopathic skeletal hyperostosis (DISH) Erdheim–Chester disease Fibrogenesis imperfecta ossium Hypertrophic osteoarthropathy Ionizing radiation Leukemia Lymphomas Mastocytosis Multiple myeloma Myelofibrosis Osteomyelitis Osteonecrosis Paget’s bone disease Polycythemia vera Sarcoidosis Sickle cell disease Skeletal metastases Tuberous sclerosis

Table III. Types of Osteopetrosis in Humans Type Benign (adult) Type I? Type II Malignant Carbonic anhydrase II deficiency Intermediate Lethal Malignant with neuronal storage disease Transient infantile Post-infectious

Inheritance* AD

AR AR AR AR AR ? –

*From Whyte [62]. AD, autosomal dominant; AR, autosomal recessive.

1993 there seemed to be at least eight clinical/genetic types (Table III) [62]. Malignant OP caused anemia and infection due to myelophthisis and cranial nerve compression. A rarer, autosomal recessive, “intermediate” form of OP presented during childhood with some of the signs and symptoms of malignant OP, but its impact on life expectancy was not well characterized (Fig. 4).

*Updated from Whyte, MP [2].

skeletal development and the dynamics of bone and cartilage metabolism. For these conditions, one wonders if increased skeletal mass per se influences blood or urinary levels of markers of bone turnover and how this should be considered in their interpretation.

A. Osteopetrosis Osteopetrosis (OP), sometimes called “marble bone disease,” was first described just over one century ago by Albers-Schönberg [63]. More than 300 cases have been reported in the medical literature [62]. Early on, two major clinical forms of OP were well delineated: autosomal dominant, adult (“benign”) type OP associated with relatively few or no symptoms, and autosomal recessive, infantile (“malignant”) type OP which, if untreated, typically kills during infancy or early childhood. However, by

Figure 4 Osteopetrosis. Skeletal mass of this 35-year-old man with intermediate severity osteopetrosis is remarkably increased, yet his bone is of poor quality and fractured in the subtrochanteric region. There is diffuse thickening of cortical and trabecular bone. He has severe anemia because medullary space has not formed.

Chapter 49 Rare Bone Diseases

A fourth clinical type, inherited as an autosomal recessive trait, had been called briefly the syndrome of “osteopetrosis with renal tubular acidosis (RTA) and cerebral calcification,” but became the first form of marble bone disease to be understood biochemically and then genetically representing an inborn error of metabolism, carbonic anhydrase II (CA II) deficiency, due to deactivating mutations in the CA II gene [64]. Neuronal storage disease with malignant OP in several infants seemed to be a distinct phenotype. There also appeared to be especially rare “lethal,” “transient infantile,” and “post-infectious” forms of OP of unknown etiology (Table III) [62]. All genuine forms of OP reflect failure of osteoclastmediated resorption of the skeleton [62]. This pathogenesis is suggested by the radiographic findings demonstrating impaired formation or expansion of medullary spaces, together with typical modeling errors such as the “Erlenmeyer flask” deformity or club-shaping of metaphyses [37]. Histological studies verify this pathogenesis showing that primary spongiosa (calcified cartilage deposited during endochondral bone formation) is not subsequently removed by bone remodeling [62]. In malignant OP, serum calcium levels generally reflect dietary mineral intake [62]. Hypocalcemia can occur and be severe enough to cause what have been interpreted to be rachitic changes on skeletal radiographs [65]. Actually, secondary hyperparathyroidism may be the explanation for these X-ray abnormalities [65]. In benign OP, standard biochemical indices of mineral homeostasis are usually unremarkable. Description of nearly 50 cases of CA II deficiency has revealed considerable clinical variability among families [64]. Typically, diagnostic investigation begins in infancy or early childhood because of failure to thrive, fracture, developmental delay, or short stature. Hyperchloremic metabolic acidosis, however, occurs as early as the neonatal period. Both proximal and distal RTA (types II and I, respectively) have been reported, although distal RTA seems to be characterized best [64]. The RTA is occasionally associated with hypokalemia which perhaps explains the apathy, hypotonia, and muscle weakness that troubles some patients. The apparent diversity of clinical/hereditary forms of OP in humans (Table III) suggested that a considerable number of genes might be compromised leading to osteoclast failure. Furthermore, the relatively large number of animal models for OP [66] and “bench” revelation of many essential factors for osteoclastogenesis and osteoclast action [55], suggested that the molecular basis for osteoclast failure in OP patients would be complex [62]. Abnormalities in the osteoclast stem cell microenvironment, progenitor cells, or mature polykaryons, or bone matrix itself could be at fault [55, 66]. Nevertheless, when

819 the first edition of this book was published in 1999, although most forms of human OP seemed to be transmitted as autosomal traits (Table III), the molecular defects were unknown except for CA II deficiency featuring deactivating mutations within the candidate CA II gene [64]. Now, there has been substantial progress in our understanding of the genetic bases of OP [67]. In 1997, benign OP had been localized to chromosome 1p21 [68] and serum M-CSF levels were unremarkable in malignant OP [69]. The other human forms of OP had not been genetically mapped. A lysosomal defect was suspected for the especially rare cases of OP with neuronal storage disease (characterized by accumulation of ceroid lipofuscin). In 1982, Marcucci et al. [70] had reported elevated levels of acid mucopolysaccharides (heparan sulfate, especially) in the urine of two patients with severe OP, providing evidence for a deficiency of lysosomal hydrolytic enzyme activity. In 2000, we found increased urinary heparan and chondroitin sulfate excretion [71]. Viral-like inclusions had been observed in osteoclasts in a few sporadic cases of benign OP, but their significance was uncertain [62]. Defective production of interleukin-2 or superoxide (factors necessary for bone resorption) or synthesis of an abnormal PTH were also potential pathogenetic defects [72]. Furthermore, in 1996, Lajeunesse and coworkers [73] implicated osteoblast dysfunction in two cases of malignant OP, suggesting disrupted interaction between abnormal osteoblasts and osteoclasts. One year earlier, the role of cytokines in bone resorption had been reviewed by Key et al. [66, 72] who provided a rationale for therapy of OP using M-CSF and interferon gamma. Findings from a number of important animal models for OP were also summarized [66]. Beginning in 2000, the cause of OP was appreciated to be deactivating mutations in additional genes that enable osteoclasts to secrete acid into the extracellular space they create overlying bone [62]. For autosomal recessive malignant OP, defects in the subunit of the osteoclast proton pump encoded by TCIRG1 and in chloride channel 7 encoded by CLCN7 were published in 2000 and 2001, respectively [74, 75]. In 2001, autosomal dominant “benign” OP (Albers–Schönberg disease) and, soon after, intermediate forms of OP, were also discovered to reflect CLCN7 defects [75]. In 2003, a defect in GL, homologous for the genetic abnormality causing the “grey-lethal” mouse model of OP, was discovered in one severely affected infant [76]. Also in 2003, a patient who was a phenocopy for CA II deficiency was discovered to combine a homozygous defect in TCIRG1, causing OP with a homozygous mutation in ATP6V1B1, causing RTA [77]. In fact, the majority of patients with OP are now understood at the gene level, with the common pathogenetic

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theme being inability of osteoclasts to acidify their pericellular milieu to resorb calcified cartilage and bone [62]. Leukocyte function studies in malignant OP revealed abnormalities in circulating monocytes and granulocytes, helping to explain predisposition to infection [72]. A variety of disturbances in osseous tissue have been characterized in patients with OP. In 1994, Danielsen and colleagues [78] used thermal stability studies of cortical bone collagen in affected adults to show that an increased proportion of this protein was aged. Skeletal fragility in OP occurs because impaired bone resorption diminishes remodeling of woven bone to compact bone and few collagen fibers properly connect osteons. Markers of bone turnover have been reported in surprisingly few individuals with OP. In the infantile form, TRAP activity is often increased in serum despite the osteoclast failure. Urinary OHP levels can be low [65, 79]. Decreased bone resorption occurs although there may be secondary hyperparathyroidism and the number of osteoclasts is increased [65]. In adult OP, serum TRAP activity and PTH levels are often increased, and urinary OHP is decreased [80]. For all genuine forms of OP in humans, the brain isoenzyme of creatine kinase (BB-CK) seems to be aberrantly present in serum [81]. Both TRAP and BB-CK likely originate from the inactive osteoclasts [81], and bisphosphonate-induced OP can cause these same enzymatic disturbances [37].

B. Pycnodysostosis Pycnodysostosis is the skeletal dysplasia that perhaps affected the French impressionist painter Henri de Toulouse-Lautrec (1864–1901) [62]. More than 100 cases from 50 kindreds have been described since the condition was delineated in 1962. Pycnodysostosis is transmitted as an autosomal recessive trait, and parental consanguinity has been reported for approximately 30% of patients. Most case descriptions have come from Europe or the United States, but the disorder has also been diagnosed in Israel, Indonesia, India, and Africa and seems to be especially prevalent in Japan [62]. Pycnodysostosis is generally discovered during infancy or early childhood because of dysmorphic physical features that include disproportionate short stature, fronto-occipital prominence, relative macrocephaly, obtuse mandibular angle, small facies and chin, high-arched palate, dental malocclusion with retained deciduous teeth, proptosis, bluish sclerae, and a beaked nose [1, 2]. Additionally, the anterior fontanelle and other cranial sutures are usually open, fingers are short and clubbed

from acro-osteolysis or aplasia of terminal phalanges, hands are small and square, fingernails are hypoplastic, and the thorax is narrow. There can also be pectus excavatum, kyphoscoliosis, and increased lumbar lordosis [2]. Mental retardation affects approximately 10% of cases [2, 62]. Adult height ranges from 4 ft 3 in to 4 ft 11 in. Recurrent fractures typically involve the lower limbs and cause knock knee deformity. Patients are, however, usually able to walk independently. Visceral manifestations and rickets have also been described. Recurrent respiratory tract infections and right heart failure (from chronic upper airway obstruction due to micrognathia) can occur. Anemia is generally not a feature. Histopathologic study shows cortical bone structure that appears to be normal despite possibly decreased osteoclastic and osteoblastic activity and slowed skeletal turnover [82]. Both the rate of bone accretion and the size of the exchangeable calcium pool can be reduced. Diminished skeletal resorption has been suggested to explain the osteosclerosis. Electron microscopy of bone from two patients indicated that degradation of collagen could be defective, perhaps from an abnormality in the skeletal matrix or in the osteoclasts themselves [83]. In fact, abnormal inclusions in osteoclasts, consisting of collagen-containing vacuoles, have been described. Virus-like inclusions were also found in the osteoclasts of two affected brothers. Serum calcium and inorganic phosphate levels and ALP activity are usually unremarkable in pycnodysostosis. Absorption of dietary calcium has been reported to be markedly increased. In 1993, the killing activity and IL-1 secretion of circulating monocytes was found to be low [84]. In 1996, defective growth hormone secretion and low serum IGF-1 levels were reported in five of six affected children [85]. That same year, the genetic basis for pycnodysostosis was discovered to be deactivating mutation within the gene encoding cathepsin K [86]. Cathepsin K, a lysosomal cysteine protease, is highly expressed in osteoclasts. Impaired bone resorption from diminished collagen degradation seems to be the fundamental pathogenetic disturbance.

C. Progressive Diaphyseal Dysplasia Progressive diaphyseal dysplasia (PDD) was characterized by Cockayne in 1920. Camurati discovered it was a heritable disease, and Engelmann described the typical severe form in 1929 [1, 2]. PDD is transmitted as an autosomal dominant trait. Descriptions of more than 100 cases show that the clinical and radiologic penetrance

Chapter 49 Rare Bone Diseases

is quite variable [1, 87]. Some especially mild cases of PDD were assumed to reflect autosomal recessive inheritance (i.e. “Ribbing disease”) [1]. However, sporadic cases do occur, and mild disease can be transmitted as an autosomal dominant trait with variable penetrance [87]. PDD typically presents during childhood with limping or a broad-based and waddling gait, leg pain, muscle wasting, and decreased subcutaneous fat in the extremities; it may be mistaken for muscular dystrophy [88]. Severely affected individuals have a characteristic body habitus that includes an enlarged head with prominent forehead, proptosis, and strikingly thin limbs with palpably thickened and tender bones and little muscle mass. Some patients have hepatosplenomegaly, Raynaud’s phenomenon, or other findings suggestive of vasculitis [89]. Cranial nerve palsies and raised intracranial pressure can occur from skull involvement. Puberty is sometimes delayed. The defining feature of PDD is hyperostosis that occurs gradually on both the periosteal and endosteal surfaces of long bone diaphyses (Fig. 5). In severe cases, osteosclerosis is widespread and the skull and axial skeleton are also involved. Some carriers manifest no radiographic changes,

Figure 5 Progressive diaphyseal dysplasia. Cortical bone thickening partly envelops the proximal diaphyseal shaft of the tibia of this 17-year-old girl, but characteristically does not extend to the metaphysis or epiphysis.

821 but bone scintigraphy is abnormal [2, 90]. Although radiographic studies typically show progressive disease, the clinical course is variable, and remission of symptoms sometimes occurs during the adult years. Bone scanning generally reveals focally increased radionuclide accumulation in affected areas [90]. Clinical, radiographic, and scintigraphic findings in PDD are generally concordant, yet bone scans can be unremarkable despite considerable X-ray changes accompanying advanced but quiescent disease. Markedly increased radioisotope accumulation with minimal radiographic findings can reflect early and active disease [2]. Cranial involvement has been assessed with magnetic resonance imaging and computed tomography. Histopathology shows that peripheral to the bony cortex, newly formed woven bone undergoes centripetal maturation and then incorporation into the cortex. Electron microscopy of muscle has shown myopathic changes and vascular abnormalities [2]. Routine biochemical parameters of bone and mineral metabolism are typically normal in PDD, although serum ALP activity and urinary OHP levels can be increased [91]. Modest hypocalcemia and significant hypocalciuria occur in some individuals with severe disease, apparently due to their markedly positive calcium balance [92]. Some of the clinical and laboratory features of PDD, together with its well-documented responsiveness to glucocorticoid treatment, led several investigators to consider PDD a systemic condition (i.e. an inflammatory connective tissue disease) [2]. In fact, mild anemia and leukopenia and an elevated erythrocyte sedimentation rate can occur [88, 89]. Aberrant differentiation of monocytes/ macrophages to fibroblasts and to osteoblasts has been discussed as a fundamental pathogenetic feature. In 1997, investigation of 18 affected individuals in a three-generation family indicated that males who inherited PDD from their fathers perhaps reflected a dynamic mutation with repeat expansion enhanced by father-to-son transmission [87]. Now, PDD is understood at the molecular level. Beginning in 2000, deactivating mutations were reported within the domain of the gene encoding transforming growth factor B1 (TGFB1) latency-associated peptide [93, 94]. In 1997, investigation of several biochemical markers of skeletal remodeling in four cases in one affected family showed increased levels for indices of both bone formation and bone resorption. The magnitude of the changes generally reflected the PDD activity determined by bone scanning. The findings were interpreted as showing rapid skeletal remodeling and utility for certain biochemical parameters in evaluating therapy [91].

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D. Endosteal Hyperostosis

E. Hepatitis C-Associated Osteosclerosis

Endosteal hyperostosis refers to several disorders featuring increased skeletal mass due to cortical bone thickening at especially endosteal surfaces [2]. Van Buchem disease (hyperostosis corticalis generalisata) is an autosomal recessive endosteal hyperostosis characterized also by enlargement of the jaw and thickening of the skull sometimes causing cranial nerve compression. Sclerosteosis is an autosomal recessive condition that resembles van Buchem disease but includes gigantism and syndactyly. These two entities were postulated more than a decade ago to be allelic disorders [1, 2]. Limited histological and biochemical investigation of skeletal remodeling suggested that osteoblast activity is excessive, with a secondary increase in bone turnover [95]. In 1998, van Buchem disease was mapped to chromosome 17q12–q21 [96]. In 2001, sclerosteosis was found to involve deactivating mutations in a gene encoding sclerostin (SOST) [97]. In 2002, van Buchem disease was attributed to a 52 kb deletion downstream to SOST [98]. The SOST protein appears to be a key factor for osteoblast action – functioning as a modulator of bone morphogenetic protein signaling [97]. In 2002, another rare type of endosteal hyperostosis gained considerable attention. During that year, gain-offunction mutation (Gly171Val) of the gene encoding lowdensity lipoprotein receptor-related protein 5 (LRP5) was identified in two American kindreds [99]. Both families manifested asymptomatic high bone mass and one also featured asymptomatic torus palatinus and a wide jaw. In 2003, however, six novel LRP5 missense mutations affecting the same LRP5 protein first β-propeller domain were reported in patients from the Americas and Europe, some with clinically significant dense bone disease. In fact, another patient with the LRP5(Gly171Val) mutation and high bone mass had significant skeletal disease including diffuse bone pain, cranial nerve palsies, extensive maxillary and mandibular exostoses surrounding teeth, and underwent craniectomy for a Chiari I malformation [99]. Because the LRP5 missense mutations associated with high bone mass alter the receptor’s antagonism by dickkopf (Dkk) or interaction with a chaperone, this endosteal hyperostosis likely reflects increased Wnt signaling through LRP5. However, as yet uncertain nutritional, environmental, or more likely genetic factors condition the clinical phenotype. Biochemical markers of skeletal remodeling are increasingly documented to remain in normal ranges, offering no clues to the pathogenesis. Conversely, deactivating mutations in LRP5 cause the osteoporosis pseudoglioma syndrome characterized by low bone mass with fractures and early-onset blindness [1].

Hepatitis C-associated osteosclerosis (HCAO) refers to a metabolic bone disease discovered in 1992 [100]. The principal radiographic feature is severe, acquired, generalized osteosclerosis and hyperostosis. Fewer than 15 cases have been reported worldwide. Evidence of infection from hepatitis C virus is documented in all patients [101]. In HCAO, forearm and leg bones are painful, seemingly from subperiosteal new bone formation. Periosteal, endosteal, and trabecular bone enhancement occurs throughout the skeleton although radiographs do not suggest thickening of the skull. Densitometry of the hip and spine reveals bone mass reaching 200–300% above control mean values. Skeletal formation is in fact accelerated on dynamic histomorphometry as implied by elevated serum total ALP, bone ALP, and osteocalcin levels, as well as enhanced radionuclide uptake on bone scanning [100]. Increased urinary OHP and deoxypyridinoline levels suggest compensatory accelerated skeletal resorption. However, biochemical markers are coming from markedly increased skeletal mass [100]. Sometimes, symptoms abate with calcitonin or bisphosphonate therapy consistent with osteoclast–osteoblast “cross talk” [100]. In fact, I have encountered gradual and seemingly spontaneous remission in pain and correction in bone density (unpublished observation). A disturbance in the IGF system, featuring elevations in circulating IGF-IIE peptide and IGF binding protein 2, could explain the high bone mass in these interesting patients [102].

F. Fibrogenesis Imperfecta Ossium Fibrogenesis imperfecta ossium (FIO) was first described in 1950. Approximately ten cases have been reported in the medical literature [2]. Although radiographs show generalized osteopenia, coarse and dense appearance of areas of trabecular bone explain why FIO is included among the osteosclerotic disorders (Table II). FIO typically presents during or after middle-age. Either sex can be affected. Intractable skeletal pain that rapidly progresses is characteristic. The debilitating course limits mobility. Spontaneous fracture is a prominent feature. Patients generally become bedridden. Physical examination typically shows marked bony tenderness. Acute agranulocytosis and macroglobulinemia have been reported [2]. The histopathological lesion of FIO resembles a form of osteomalacia, although the amount of diseased

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Chapter 49 Rare Bone Diseases

osseous tissue varies considerably from area to area within the skeleton. Cortical bone in the femora and tibiae demonstrates the least abnormality. Osteoid seams are thick and osteoblasts and osteoclasts can be abundant. Aberrant collagen is found in regions with abnormal mineralization patterns, yet collagen is unremarkable in other tissues. Polarized-light microscopy shows that collagen fibrils lack birefringence in areas of abnormal bone. Electron microscopy reveals thin and randomly organized collagen fibrils in a tangled pattern. In some areas, one sees peculiar, circular, matrix structures of 300–500 nm diameter. Unless bone is viewed with polarized-light or electron microscopy, FIO can be mistaken for osteoporosis or some other form of osteomalacia [2]. The etiology of FIO is unknown. Genetic factors have not been implicated because cases have been sporadic. FIO seems to be an acquired disorder of collagen synthesis in lamellar bone. Serum calcium and inorganic phosphate levels are normal, yet ALP activity is increased. OHP levels in urine may be normal or elevated. Typically, there is no aminoaciduria or other evidence of renal tubular dysfunction.

Table IV.

Disorders Associated with Extraskeletal Calcification or Ossification

Metastatic calcification Hypercalcemia Milk-alkali syndrome Hypervitaminosis D Sarcoidosis Hyperparathyroidism Renal failure Hyperphosphatemia Tumoral calcinosis Hyperparathyroidism Pseudohypoparathyroidism Cell lysis following chemotherapy for leukemia Renal failure Dystrophic calcification Calcinosis (universalis or circumscripta) Childhood dermatomyositis Scleroderma Systemic lupus erythematosus Post-traumatic Ectopic ossification Myositis ossificans (post-traumatic) Burns Surgery Neurologic injury Fibrodysplasia (myositis) ossificans progressiva Reproduced with permission from Wenkert and Whyte [105].

G. Pachydermoperiostosis Pachydermoperiostosis (primary hypertrophic osteoarthropathy) is a rare, autosomal dominant disorder characterized by periosteal new bone formation, thick skin, and clubbing of the fingers and toes [1, 2]. There are also autosomal recessive forms [1, 103]. Patients suffer from arthralgias. Alteration of blood flow to involved skeletal regions has been reported as a possible pathogenetic disturbance. In one affected individual, skin fibroblasts in culture synthesized decreased amounts of collagen and significant amounts of the proteoglycan, decorin [104]. Despite these remarkable clinical manifestations, markers of skeletal homeostasis have not been adequately studied in pachydermoperiostosis.

IV. ECTOPIC CALCIFICATION A significant number and variety of disorders cause extraskeletal deposition of calcium and phosphate (Table IV) [105]. In some, mineral is precipitated as amorphous calcium phosphate or as hydroxyapatite crystals; in others, bone is formed. The pathogenesis of the ectopic mineralization is generally attributed to one of three mechanisms: metastatic calcification, dystrophic calcification, or ectopic ossification.

A. Fibrodysplasia (Myositis) Ossificans Progressiva Fibrodysplasia ossificans progressiva (FOP) is a heritable disorder of connective tissue characterized by congenital malformations of the great toes and recurrent, painful, soft-tissue swellings that often become heterotopic bone. More than 600 cases have been reported involving all ethnic groups. Autosomal dominant transmission is rare, but established [1]. FOP can be suspected at birth, before soft-tissue lesions appear, if the typical congenital digital malformations are recognized. The hallmark is shortening and deviation of the big toes (hallux valgus) due to defective formation of the cartilaginous anlagen of the first metatarsal and proximal phalange. Nevertheless, these digital anomalies are not pathognomonic of FOP. Usually, diagnosis occurs when characteristic masses with radiographic evidence of heterotopic ossification are discovered [1]. Episodes of soft-tissue swelling from FOP typically commence during the first decade of life, although I have recently encountered a woman who first manifested such lesions at age 47 years [106]. Painful, tender, and rubbery masses appear spontaneously, or may follow even minor trauma (e.g. intramuscular injection). Typically, swellings occur in paraspinal muscles or in the limb girdle and can last for several weeks. Most lesions then mature to

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contain heterotopic bone. Gradually, ectopic bone can immobilize joints and cause contractures and deformity, particularly in the neck and shoulders. Another remarkable radiographic feature of FOP is fusion of cervical vertebrae early in life; scoliosis is also common [2]. Ankylosis of the spine and rib cage eventually imperil cardiopulmonary function. Bone scans show increased radionuclide uptake before ossification appears on radiographs. Computerized tomography and magnetic resonance imaging of early lesions has been described. Radiographs as well as histopathologic examination of abnormal bone indicate that modeling and remodeling occurs within the heterotopic skeleton. The soft-tissue masses of FOP initially comprise one or more edematous skeletal muscles. Edema of fascial planes is one of the earliest findings. Subsequently, endochondral ossification leads to heterotopic bone. Immunostaining for bone morphogenetic protein (BMP) 2/4 is intense. Mature osseous lesions have Haversian systems and cancellous bone contains hematopoietic tissue [107]. In FOP, routine biochemical studies of mineral metabolism are usually normal, although ALP activity in serum may be increased, especially during disease flare-ups.

Figure 6 Ectopic calcification in juvenile dermatomyositis. Numerous radiopaque areas of calcification are present above the ankle of this 16-year-old boy with a 6-year history of juvenile dermatomyositis.

B. Juvenile Dermatomyositis Dermatomyositis is a multisystem connective tissue disorder caused by small vessel vasculitis and features a non-suppurative inflammatory process of especially skin and striated muscle [108]. Dystrophic calcification often follows episodes of inflammation. There are more female than male patients and two peak ages of incidence: childhood (5–15 years) and adulthood (50–60 years). When dermatomyositis manifests before age 16 years, it is called the juvenile form. The adult form is associated with malignancy [2]. Juvenile dermatomyositis (JD) is now responding better to treatment using vigorous pharmacologic and immunologic approaches; consequently, the prevalence of calcinosis is likely decreasing. The patient’s sex and age of disease onset do not, however, seem to influence the severity of calcinosis, which is typically noted 1–3 years after diagnosis and usually following the acute phase (Fig. 6). Mineral deposits present over 1–3 years. This dystrophic calcification then typically remains stable, although there can be some regression [108]. Radiographic studies delineate four types of dystrophic calcification in JD: (i) superficial masses; (ii) deep, discrete, subcutaneous, nodular masses; (iii) deep, linear, sheetlike deposits within intramuscular fascial planes (calcinosis universalis); and (iv) lacy, reticular, subcutaneous deposits

that encase the torso to form a generalized “exoskeleton” [108]. Children with severe JD refractory to therapy seem especially prone to developing an exoskeleton featuring severe calcinosis and poor physical function. With calcinosis universalis, mineral deposition occurs throughout the subcutaneous tissues, but primarily involves periarticular regions or areas subject to trauma. Calcinosis circumscripta deposits are more localized and typically surround joints. The ectopic calcification can cause pain, ulcerate the skin, limit mobility, and predispose to abscesses. JD seems to be a form of complement-mediated microangiopathy. Nevertheless, the precise mechanism for the dystrophic calcification is unknown. Predating aggressive medical therapies, mineral deposition seemed to occur in the majority of long-term survivors reflecting a “scarring” process. This hypothesis was bolstered by the observation that calcification occurs primarily in the muscles that are most severely affected during the acute phase. Among the postulated mechanisms for the dystrophic calcification in JD has been release of ALP or discharge of free fatty acids from diseased muscle directly precipitating calcium or first binding acid mucopolysaccharides.

Chapter 49 Rare Bone Diseases

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Electron microscopy shows hydroxyapatite crystals [105]. Increased urinary levels of gamma-carboxylated peptides suggested that calcium-binding proteins may engender the mineral deposition. In 1994, Perez et al. [109] reported dual-tracer, stable isotope studies of 12 affected children and concluded that patients also risk bone mineral loss due, in part, to decreased dietary calcium absorption, especially acutely and during glucocorticoid therapy. Hypercalcemia with hypercalciuria and hyperphosphaturia may occur.

V. OTHER DISORDERS Fibrous dysplasia, including the McCune–Albright syndrome, and systemic mastocytosis are among the rare bone diseases that have been assessed using biochemical markers of skeletal remodeling.

A. Fibrous Dysplasia and McCune–Albright Syndrome Fibrous dysplasia (FD) is a sporadic, developmental disorder characterized by a unifocal or multifocal expanding fibrous lesion of bone-forming mesenchyme that often results in deformity and sometimes in fracture. McCune–Albright syndrome (MAS) refers to patients with polyostotic FD and hyperpigmented macules, called café-au-lait spots, who also manifest primary hyperfunction of one or more endocrine glands. Both sexes are affected [1, 2]. Monostotic FD characteristically develops during the second or third decade of life; polyostotic disease typically manifests before 10 years of age. Monostotic disease is more common. Generally, an expansile FD lesion causes deformity or fracture and may occasionally entrap nerves. The skull and long bones are most often affected (Fig. 7). Pregnancy may “reactivate” quiescent lesions. Sarcomatous degeneration of FD sometimes occurs (incidence <1%), especially when facial bones or femora are involved. MAS features café-au-lait spots with rough borders, compared to smooth borders in neurofibromatosis (i.e. “Coast-of-Maine” versus “Coast-of-California,” respectively). Most often, the endocrinopathy is pseudoprecocious puberty in girls due to ovarian hyperactivity. Less commonly, there is thyrotoxicosis, Cushing’s disease, acromegaly, hyperprolactinemia, hyperparathyroidism, or pseudoprecocious puberty in boys. Increasingly, patients with MAS are recognized to have acquired renal

Figure 7 Fibrous dysplasia in McCune–Albright syndrome. Heterogeneous areas of osteopenia and osteosclerosis with cortical thinning (arrows) are present in the proximal right femur of this 3-year-old girl.

phosphate wasting causing hypophosphatemic rickets or osteomalacia – especially with widespread FD lesions. Both monostotic and polyostotic FD have a similar histological appearance. Lesions appear well defined, but are not encapsulated. Spindle-shaped fibroblasts form characteristic “swirls” within the marrow space. Haphazardly arranged trabeculae contain woven bone. Cartilage is a component, especially with polyostotic disease. Cystic regions may occur lined by multinucleated giant cells. The histopathology resembles hyperparathyroid bone disease (osteitis fibrosa cystica), but few osteoblasts reflect the basic biological disturbance. Skeletal lesions result from the formation of woven bone and fibrotic matrix as mesenchymal cells proliferate but do not differentiate fully to osteoblasts. Endocrine hyperfunction in MAS is typically end-organ overactivity. FD and MAS reflect post-zygotic mosaicism of an activating mutation in the gene that encodes the Gsα subunit of the receptor/adenylate cyclase-coupling G protein [110–112]. Such mutations enhance this pathway for transmembrane signaling, leading to overactivity of

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adenyl cyclase, and can be impacted by genetic imprinting [111]. Fibrotic areas in bone contain excesses of preosteogenic cells, and the lesional bone is produced by mature, but peculiarly shaped, osteoblasts. Bone matrix is rich and poor in certain antiadhesion and proadhesion molecules, respectively [113]. Synthesis of osteocalcin is diminished [112]. In 1996, increased IL-6 secretion was reported [112]. Although serum ALP activity may be elevated in FD or MAS, calcium levels are usually normal unless there is hyperparathyroidism. Studies of biochemical markers of skeletal turnover, often to assess bisphosphonate therapy, indicate that remodeling in fibrodysplastic bone is accelerated [114]. Hypophosphatemic rickets in MAS is associated with increased circulating levels of fibroblast growth factor 23.

plasma cells, lymphocytes, and characteristic oval- or spindle-shaped mast cells. These spindle-shaped mast cells superficially resemble histiocytes or fibroblasts, but contain granules that stain metachromatically [116]. Additionally, marrow includes increased numbers of such mast cells individually or in smaller aggregates [116]. Histomorphometric studies following tetracycline labeling show that skeletal remodeling is usually normal or rapid in SM. Evaluation of biochemical markers of bone remodeling reveals elevations in serum ALP and TRAP activity and in osteocalcin levels in some patients [115]. Activating c-kit mutations are considered the hallmark of the neoplastic forms of SM [117].

B. Mastocytosis

Supported in part by Shriners Hospitals for Children and The Clark and Mildred Cox Inherited Metabolic Bone Disease Research Fund. Becky Whitener, CPS, provided expert secretarial help.

Systemic mastocytosis (SM) features increased numbers of mast cells in the viscera. The skin also can contain many hyperpigmented macules reflecting dermal mast cell accumulation called urticaria pigmentosa. Bone marrow is typically involved, causing the skeletal pathology. Symptoms of SM result primarily from release of mediator substances by the deranged mast cells and include generalized pruritus, urticaria, flushing, episodic hypotension, syncope, diarrhea, weight loss, and peptic ulcer. Skeletal manifestations are relatively infrequent but include bone pain or tenderness from deformity resulting from fracture [2]. Radiographic abnormalities of the skeleton are common in SM (about 70% of patients). Diffuse, poorly demarcated, sclerotic, and lucent areas are typically seen where hematopoietic marrow is normally present (i.e. the axial skeleton in adults). However, circumscribed lesions can occur, especially in the extremities as well as in the skull. Lytic areas are often small and have a surrounding rim of osteosclerosis. Generalized osteopenia alone is also a common radiographic presentation of SM. Urinary levels of methylimidazoleacetic acid, a metabolite of histamine and an indicator of mast cell burden, correlate with bone mineral density [115]. The histopathological findings of SM in the skeleton are now well characterized [116]. In fact, examination of non-decalcified sections of iliac crest can be a particularly effective way to establish the diagnosis. Iliac crest biopsy may be superior to bone marrow aspiration or biopsy. Multiple nodules, 150–450 µm in diameter, that resemble granulomas are found in the marrow space. Within these cellular aggregations are eosinophils,

Acknowledgments

References 1. McKusick, V. A. (1998). Mendelian inheritance in man: A catalog of human genes and genetic disorders, 12th ed. Johns Hopkins University Press, Baltimore, MD and 2000 Online Mendelian Inheritance in Man, OMIM™, Baltimore, MD: McKusick-Nathans Institute for Genetic Medicine, Johns Hopkins University and Bethesda, MD: National Center for Biotechnology Information, National Library of Medicine. 2. Whyte, M. P. (2002). Hereditary metabolic and dysplastic skeletal disorders. In “Disorders of Bone and Mineral Metabolism,” 2nd Ed. (F. L. Coe, M. J. Favus, eds.), pp. 937–984. Lippincott, Williams & Wilkins, Philadelphia, PA. 3. Kanis, J. A. (1998). Pathophysiology and Treatment Of Paget’s Disease Of Bone, 2nd Ed. Martin Dunitz, London, UK. 4. Byers, P. H. (2001). Disorders of collagen biosynthesis and structure. In “The Metabolic and Molecular Bases of Inherited Disease,” 8th Ed. (C. R. Scriver, A. L. Beaudet, W. S. Sly, D. Valle, eds.), pp. 5241–5285. McGraw-Hill, New York. 5. Rowe, D. W., and Shapiro, J. R. (1995). Osteogenesis imperfecta. In “Metabolic Bone Disease” (L.V. Avioli, S.M. Krane, eds.), pp. 651–695. Academic Press, San Diego, CA. 6. Sillence, D. O., Senn, A., and Danks, D. M. (1979). Genetic heterogeneity in osteogenesis imperfecta. J. Med. Genet. 16, 101–116. 7. Dalgleish, R. (1997). The human type I collagen mutation database. Nuc. Acids Res. 25, 181–187. Dalgleish, R. (1998). The human collagen mutation database. Nuc. Acids Res. 26, 253–255. http://www.le.ac.uk/genetics/collagen 8. Fedarko, N. S., Mörike, M., Brenner, R., Robey, P. G., and Vetter, U. (1992). Extracellular matrix formation by osteoblasts from patients with osteogenesis imperfecta. J. Bone Miner. Res. 7, 921–930. 9. Cassella, J. P., and Ali, S. Y. (1992). Abnormal collagen and mineral formation in osteogenesis imperfecta. Bone Min. 17, 123–128. 10. Doty, S. B., and Mathews, R. S. (1971). Electron microscopic and histochemical investigation of osteogenesis imperfecta tarda. Clin. Orthop. 80, 191–201.

Chapter 49 Rare Bone Diseases 11. Ste-Marie, L. G., Charhon, S. A., Edouard, C., Chapuy, M. C., and Meunier, P. J. (1984). Iliac bone histomorphometry in adults and children with osteogenesis imperfecta. J. Clin. Pathol. 37, 1081–1089. 12. Baron, R., Gertner, J. M., Lang, R., and Vignery, A. (1983). Increased bone turnover with decreased bone formation by osteoblasts in children with osteogenesis imperfecta tarda. Pediatr. Res. 17, 204–207. 13. Vetter, U., Eanes, E. D., Kopp, J. B., Termine, J. D., and Robey, P. G. (1991). Changes in apatite crystal size in bones of patients with osteogenesis imperfecta. Calcif. Tissue Int. 49, 248–250. 14. Vetter, U., Fisher, L. W., Mintz, K. P., Kopp, J. B., Tuross, N., Termine, J. D., and Robey, P. G. (1991). Osteogenesis imperfecta: Changes in noncollagenous proteins in bone. J. Bone Miner. Res. 6, 501–505. 15. Dickson, I. R., Millar, E. A., and Veis, A. (1975). Evidence for abnormality of bone-matrix proteins in osteogenesis imperfecta. Lancet 2, 586–587. 16. Riley, F. C., Jowsey, J., and Brown, D. M. (1973). Osteogenesis imperfecta: Morphologic and biochemical studies of connective tissue. Pediatr. Res. 7, 757–768. 17. Shoenfeld, Y. (1975). Osteogenesis imperfecta. Review of the literature with presentation of 29 cases. Am. J. Dis. Child. 129, 679–687. 18. Fedarko, N. S., Robey, P. G., and Vetter, U. K. (1995). Extracellular matrix stoichiometry in osteoblasts from patients with osteogenesis imperfecta. J. Bone Miner. Res. 10, 1122–1129. 19. Mörike, M., Brenner, R. E., Bushart, G. B., Teller, W. M., and Vetter, U. (1992). Collagen metabolism in cultured osteoblasts from osteogenesis imperfecta patients. Biochem. J. 286, 73–77. 20. Mörike, M., Schulz, M., Brenner, R. E., Bushart, G. B., Teller, W. M., and Vetter, U. (1993). In vitro expression of osteoblastic markers in cells isolated from normal fetal and postnatal human bone and from bone of patients with osteogenesis imperfecta. J. Cell. Physiol. 157, 439–444. 21. Lehmann, H. W., Rimek, D., Bodo, M., Brenner, R. E., Vetter, U., Wörsörfer, O., Karbowski, A., and Müller, P. K. (1995). Hydroxylation of collagen type I: evidence that both lysyl and prolyl residues are overhydroxylated in osteogenesis imperfecta. Eur. J. Clin. Invest. 25, 306–310. 22. Chines, A., Petersen, D. J., Schranck, F. W., and Whyte, M. P. (1991). Hypercalciuria in children severely affected with osteogenesis imperfecta. J. Pediatr. 119, 51–57. 23. Chines, A., Boniface, A., McAlister, W., and Whyte, M. (1995). Hypercalciuria in osteogenesis imperfecta: A follow-up study to assess renal effects. Bone 16, 333–339. 24. Nielsen, H. E., Pedersen, U., Hansen, H. H., and Elbrond, O. (1979). Serum calcitonin and bone mineral content in patients with osteogenesis imperfecta. Acta Orthop. Scand. 50, 639–643. 25. Marini, J. C., Bordenick, S., Heavner, G., Rose, S., Hintz, R., Rosenfeld, R., and Chrousos, G. P. (1993). The growth hormone and somatomedin axis in short children with osteogenesis imperfecta. J. Clin. Endocrinol. Metab. 76, 251–256. 26. Granda, J. L., Falvo, K. A., and Bullough, P. G. (1977). Pyrophosphate levels and magnesium oxide therapy in osteogenesis imperfecta. Clin. Orthop. 126, 228–231. 27. Cropp, G. J., and Myers, D. N. (1972). Physiological evidence of hypermetabolism in osteogenesis imperfecta. Pediatrics 49, 375–391. 28. Wyckoff, M. H., El-Turk, C., Laptook, A., Timmons, C., Gannon, F. H., Zhang, X., Mumm, S., and Whyte, M. P. (2005). Neonatal lethal osteochondrodysplasia with low serum levels of alkaline phosphatase and osteocalcin J. Clin. Endocrin. Metab. (in press) 29. Castells, S., Yasumura, S., Fusi, M. A., Colbert, C., Bachtell, R. S., and Smith, S. (1986). Plasma osteocalcin levels in patients with osteogenesis imperfecta. J. Pediatr. 109, 88–91. 30. Rico, H. (1987). Osteocalcin levels in osteogenesis imperfecta. J. Pediatr. 111, 634. 31. Rico, H., Revilla, M., Iritia, M., Arribas, I., and Villa, L. F. (1991). Total body bone mineral and tartrate-resistant acid phosphatase levels

827 in type I and III osteogenesis imperfecta. Miner. Electrolyte Metab. 17, 396–398. 32. Brenner, R. E., Schiller, B., Vetter, U., Ittner, J., and Teller, W. M. (1993). Serum concentrations of procollagen I C-terminal propeptide, osteocalcin and insulin-like growth factor-I in patients with non-lethal osteogenesis imperfecta. Acta Paediatr. 82, 764–767. 33. Minisola, S., Piccioni, A. L., Rosso, R., Romagnoli, E., Pacitti, M. T., Scarnecchia, L., and Mazzuoli, G. (1994). Reduced serum levels of carboxy-terminal propeptide of human type I procollagen in a family with type I-A osteogenesis imperfecta. Meta. Clin. Exp. 43, 1261–1266. 34. Brenner, R. E., Vetter, U., Bollen, A. M., Mörike, M., and Eyre, D. R. (1994). Bone resorption assessed by immunoassay of urinary crosslinked collagen peptides in patients with osteogenesis imperfecta. J. Bone Miner. Res. 9, 993–997. 35. Rauch, F., Travers, R., Plotkin, H., and Glorieux, F. H. (2002). The effects of intravenous pamidronate on the bone tissue of children and adolescents with osteogenesis imperfecta. J. Clin. Invest. 110, 1293–1299. 36. Rauch, F., Plotkin, H., Travers, R., Zeitlin, L., and Glorieux, F. H. (2003). Osteogenesis imperfecta types I, III, and IV: effect of pamidronate therapy on bone and mineral metabolism. J. Clin. Endocrinol. Metab. 88, 986–992. 37. Whyte, M. P., Wenkert, D., Clements, K. L., McAlister, W. H., and Mumm, S. (2003). Bisphosphonate-induced osteopetrosis. New Engl. J. Med. 349, 455–461. 38. Norman, M. E. (1996). Idiopathic juvenile osteoporosis. In “Primer on the Metabolic Bone Diseases and Disorders of Mineral Metabolism” (M. Favus, ed.), pp. 275–278. Lippincott-Raven, New York. 39. Smith, R. (1980). Idiopathic osteoporosis in the young. J. Bone Joint Surg. 62-B, 417–427. 40. Dent, C. E., and Friedman, M. (1965). Idiopathic juvenile osteoporosis. Q. J. Med. 34, 177–210. 41. Schippers, J. C. (1938). “Spontaneous” generalized osteoporosis in girl 10 years old. Maandschr Kindergeneeskd 8, 108–116. 42. US Surgeon General’s Report, “Bone Health and Osteoporosis” 2004. 43. Smith, R. (1980). Idiopathic osteoporosis in the young. J. Bone Joint. Surg. Br. 62, 417–427. 44. Smith, R. (1995). Idiopathic juvenile osteoporosis: Experience of twenty-one patients. Br. J. Rheumatol. 34, 68–77. 45. Jowsey, J., and Johnson, K. A. (1972). Juvenile osteoporosis: Bone findings in seven patients. J. Pediatr. 81, 511–517. 46. Bailey, D. A., McKay, H. A., Mirwald, R. L., Crocker, P. R., and Faulkner, R. A. (1999). A six-year longitudinal study of the relationship of physical activity to bone mineral accrual in growing children: The University of Saskatchewan bone mineral accrual study. J. Bone Miner. Res. 14, 1672–1679. 47. Abrams, S. A., Griffin, I. J., Hicks, P. D., and Gunn, S. K. (2004). Pubertal girls only partially adapt to low dietary calcium intakes. J. Bone Miner. Res. 19, 759–763. 48. Cloutier, M. D., Hayles, A. B., Riggs, B. L., Jowsey, J., and Bickel, W. H. (1967). Juvenile osteoporosis: Report of a case including a description of some metabolic and microradiographic studies. Pediatrics 40, 649–655. 49. Evans, R. A., Dunstan, C. R., and Hills, E. (1983). Bone metabolism in idiopathic juvenile osteoporosis: A case report. Calcif. Tissue Int. 35, 5–8. 50. Rauch, F., Travers, R., Norman, M. E., Taylor, A., Parfitt, A. M., and Glorieux, F. H. (2002). The bone formation defect in idiopathic juvenile osteoporosis is surface–specific. Bone 31, 85–89. 51. Bertelloni, S., Baroncelli, G. I., Di Nero, G., and Saggese, G. (1992). Idiopathic juvenile osteoporosis: Evidence of normal osteoblast function by 1,25-dihydroxyvitamin D3 stimulation test. Calcif. Tissue Int. 51, 20–23. 52. Pocock, A. E., Francis, M. J., and Smith, R. (1995). Type I collagen biosynthesis by skin fibroblasts from patients with idiopathic juvenile osteoporosis. Clin. Sci. 89, 69–73.

828 53. Jackson, E. C., Strife, F., Tsang, R. C., and Marder, H. K. (1988). Effect of calcitonin replacement therapy in idiopathic juvenile osteoporosis. Am. J. Dis. Child. 142, 1237–1239. 54. Saggese, G., Bertelloni, S., Baroncelli, G. I., Perri, G., and Calderazzi, A. (1991). Mineral metabolism and calcitriol therapy in idiopathic juvenile osteoporosis. Am. J. Dis. Child. 145, 457–462. 55. Martin, T. J. (2004). Paracrine regulation of osteoclast formation and activity: Milestones in discovery. J. Musculoskel. Neuro. Interact. 4, 243–253. 56. Whyte, M. P., and Mumm, S. (2004). Heritable disorders of the RANKL/OPG/RANK signaling pathway. J. Musculoskel. Neuro. Interact 4, 254–267. 57. Moore, C. A., Ayangade, G., Okolo, P., Bull, M. J., Whyte, M. P., and Bixler, D. (1992). Cleidocranial dysplasia: A natural history study including evaluation of a five generation family (Abstract). Proc. Greenwood Genet. Center 10, 91. 58. Mundlos, S., Otto, F., Mundlos, C., Mulliken, J. B., Aylsworth, A. S., Albright, S., Lindhout, D., Cole, W. G., Henn, W., Knoll, J. H. M., Owen, M. J., Mertelsmann, R., Zabel, B. U., and Olsen, B. R. (1997). Mutations involving the transcription factor CBFA1 cause cleidocranial dysplasia. Cell 89, 773–779. 59. Otto, F., Thornell, A. P., Crompton, T., Denzel, A., Gilmour, K. C., Rosewell, I. R., Stamp, G. W., Beddington, R. S., Mundlos, S., Olsen, B. R., Selby, P. B., and Owen, M. J. (1997). Cbfal, a candidate gene for cleidocranial dysplasia syndrome, is essential for osteoblast differentiation and bone development. Cell 89, 765–771. 60. Unger, S., Mornet, E., Mundlos, S., Blaser, S., and Cole, D. E. (2002). Severe cleidocranial dysplasia can mimic hypophosphatasia. Eur. J. Pediatr. 161, 623–626. 61. Morava, E., Karteszi, J., Weisenbach, J., Caliebe, A., Mundlos, S., and Mehes, K. (2002). Cleidocranial dysplasia with decreased bone density and biochemical findings of hypophosphatasia. Eur. J. Pediatr. 161, 619–622. 62. Whyte, M. P. (2002). Osteopetrosis. In “Connective Tissue and Its Heritable Disorders” (P. M. Royce, and B. Steinmann, eds.), 2nd Edition, pp. 789–807. Wiley-Liss, New York. 63. Albers-Schönberg, H. (1904). Röntgenbilder einer seltenen Knochenerkrankung. Münch. Med. Wochenschr. 51, 365. 64. Sly, W. S., and Hu, P. Y. (2001). The carbonic anhydrase II deficiency syndrome: Osteoporosis with renal tubular acidosis and cerebral calcification. In “The Metabolic and Molecular Bases of Inherited Disease” (C. R. Scriver, A. L. Beaudet, S. William, S. Sly, D. Valle, eds.), 8th ed, pp. 5331–5343. McGraw-Hill, Inc., New York. 65. Reeves, J., Arnaud, S., Gordon, S., Subryan, B., Block, M., Huffer, W., Arnaud, C., Mundy, G., and Haussler, M. (1981). The pathogenesis of infantile malignant osteopetrosis: Bone mineral metabolism and complications in five infants. Metab. Bone Dis. Rel. Res. 3, 135–142. 66. Key, L. L., Jr., Rodriguiz, R. M., and Wang, W. C. (1995). Cytokines and bone resorption in osteopetrosis. Int. J. Pediatr. Hematol./Oncol. 2, 143–149. 67. Tolar, J., Teitelbaum, S. L., and Orchard, P. J. (2004). Osteopetrosis. N. Engl. J. Med. 351, 2839–2849. 68. Van Hul, W., Bollerslev, J., Gram, J., Van Hul, E., Wuyts, W., Benichou, O., Vanhoenacker, F., and Willems, P. J. (1997). Localization of a gene for autosomal dominant osteopetrosis (Albers-Schönberg disease) to chromosome 1p21. Am. J. Hum. Genet. 61, 363–369. 69. Naffakh, N., Le Gall, S., Danos, O., Heard, J. M., Cournot, G., Motoyoshi, K., and Vilmer, E. (1993). Macrophage colony-stimulating factor: Serum levels and cDNA structure in malignant osteopetrosis (letter). Blood 81, 2817–2818. 70. Marcucci, F., Rufini, S., and Sensi, L. (1982). Abnormal urinary excretion of glycosaminoglycans in Albers-Schönberg disease. Clin. Chim. Acta 120, 161–170. 71. Steiner, R. D., Whyte, M. P., Chang, E., Hanks, J., Mattes, C., Senephansiri, H., and Gibson, K. M. (2000). Increased urine heparan

MICHAEL P. WHYTE

72.

73.

74.

75.

76.

77.

78.

79.

80.

81.

82.

83.

84.

85.

86.

87.

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and chondroitin sulfate excretion in patients with osteopetrosis. J. Inher. Metab. Dis. 23, 88–90. Key, L. L., Jr., Rodriguiz, R. M., Willi, S. M., Wright, N. M., Hatcher, H. C., Eyre, D. R., Cure, J. K., Griffin, P. P., and Ries, W. L. (1995). Long-term treatment of osteopetrosis with recombinant human interferon gamma. N. Engl. J. Med. 332, 1594–1599. Lajeunesse, D., Busque, L., Mènard, P., Brunette, M. G., and Bonny, Y. (1996). Demonstration of an osteoblast defect in two cases of human malignant osteopetrosis. Correction of the phenotype after bone marrow transplant. J. Clin. Invest. 98, 1835–1842. Frattini, A., Orchard, P. J., Sobacchi, C., Giliani, S., Abinun, M., Mattson, J. P., Keeling, D. J., Andersson, A-K., Wallbrandt, P., Zecca, L., Notarangelo, L. D., Vezzoni, P., and Villa, A. (2000). Defects in TCIRG1 subunit of the vacuolar proton pump are responsible for a subset of human autosomal recessive osteopetrosis. Nat. Genet. 25, 343–346. Cleiren, E., Benichou, O., Van Hul, E., Gram, J., Bollerslev, J., Singer, F. R., Beaverson, K., Aledo, A., Whyte, M. P., Yoneyama, T., deVernejoul, M. C., and Van Hul, W. (2001). Albers-Schönberg disease (autosomal dominant osteopetrosis, type II) results from mutations in the ClCN7 chloride channel gene. Hum. Mol. Genet. 10, 2861–2867. Chalhoub, N., Benachenhou, N., Rajapurohitam, V., Pata, M., Ferron, M., Frattini, A., Villa, A., and Vacher, J. (2003). Grey-lethal mutation induces severe malignant autosomal recessive osteopetrosis in mouse and human. Nat. Med. 9, 399–406. Borthwick, K. J., Kandemir, N., Topaloglu, R., Kornak, U., Bakkaloglu, A., Yordam, N., Ozen, S., Mocan, H., Shah, G. N., Sly, W. S., and Karet, F. E. (2003). A phenocopy of CAII deficiency: a novel genetic explanation for inherited infantile osteopetrosis with distal renal tubular acidosis. J. Med. Genet. 40, 115–121. Danielsen, C. C., Mosekilde, L., Bollerslev, J., and Mosekilde, L. (1994). Thermal stability of cortical bone collagen in relation to age in normal individuals and in individuals with osteopetrosis. Bone 15, 91–96. Key, L. L., Jr., Ries, W. L., Rodriguiz, R. M., and Hatcher, H. C. (1992). Recombinant human interferon gamma therapy for osteopetrosis. J. Pediatr. 121, 119–124. Bollerslev, J. (1989). Autosomal dominant osteopetrosis: Bone metabolism and epidemiological, clinical, and hormonal aspects. Endocr. Rev. 10, 45–67. Whyte, M. P., Chines, A., Silva, D. P., Landt, Y., and Ladenson, J. H. (1996). Creatine kinase brain isoenzyme (BB-CK) presence in serum distinguishes osteopetroses among the sclerosing bone disorders. J. Bone Miner. Res. 11, 1438–1443. Soto, R. J., Mautalen, C. A., Hojman, D., Codevilla, A., Piquè J., and Pángaro, J. A. (1969). Pycnodysostosis: Metabolic and histologic studies. Birth Defects Orig. Artic. Ser. 5, 109–116. Everts, V., Aronson, D. C., and Beertsen, W. (1985). Phagocytosis of bone collagen by osteoclasts in two cases of pycnodysostosis. Calcif. Tissue Int. 37, 25–31. Karkabi, S., Reis, N. D., Linn, S., Edelson, G., Tzehoval, E., Zakut, V., Dolev, E., Bar-Meir, E., and Ish-Shalom, S. (1993). Pycnodysostosis: Imaging and laboratory observations. Calcif. Tissue Int. 53, 170–173. Soliman, A. T., Rajab, A., AlSalmi, I., Darwish, A., and Asfour, M. (1996). Defective growth hormone secretion in children with pycnodysostosis and improved linear growth after growth hormone treatment. Arch. Dis. Child. 75, 242–244. Gelb, B. D., Shi, G. P., Chapman, H. A., and Desnick, R. J. (1996). Pycnodysostosis, a lysosomal disease caused by cathepsin K deficiency. Science 273, 1236–1238. Saraiva, J. M. (1997). Progressive diaphyseal dysplasia: a threegeneration family with markedly variable expressivity. Am. J. Med. Genet. 71, 348–352. Low, L. C., Stephenson, J. B., and Stuart-Smith, D. A. (1985). Progressive diaphyseal dysplasia mimicking childhood myopathy: Clinical and biochemical response to prednisolone. Aust. Paediatr. J. 21, 193–196.

Chapter 49 Rare Bone Diseases 89. Naveh, Y., Alon, U., Kaftori, J. K., and Berant, M. (1985). Progressive diaphyseal dysplasia: Evaluation of corticosteroid therapy. Pediatrics 75, 321–323. 90. Sty, J. R., Babbitt, D. P., and Starshak, R. J. (1978). Bone scintigraphy demonstrating Engelmann’s disease. Clin. Nucl. Med. 3, 69–70. 91. Hernandez, M. V., Peris, P., Guanabens, N., Alvarez, L., Monegal, A., Pons, F., Ponce, A., and Munoz-Gomez, J. (1997). Biochemical markers of bone turnover in Camurati-Engelmann disease: a report on four cases in one family. Calcif. Tis. Internat. 61, 48–51. 92. Smith, R., Walton, R. J., Corner, B. D., and Gordon, I. R. (1977). Clinical and biochemical studies in Engelmann’s disease (progressive diaphyseal dysplasia). Q. J. Med. 46, 273–294. 93. Kinoshita, A., Saito, T., Tomita, H., Makita, Y., Yoshida, K., Ghadami, M., Yamada, K., Kondo, S., Ikegawa, S., Nishimura, G., Fukushima, Y., Nakagomi, T., Saita, H., Sugimoto, T., Kamegaya, M., Hisa, K., Murray, J. C., Taniguchi, N., Niikawa, N., and Yoshiura, K. (2000). Domain-specific mutations in TGFB1 result in CamuratiEngelmann disease. Nat. Genet. 26, 19–20. 94. Janssens, K., Gershoni-Baruch, R., Guanabens, N., Migone, N., Ralston, S., Bonduelle, M., Lissens, W., Van Maldergem, L., Vanhoenacker, F., Verbruggen, L., and Van Hul, W. (2000). Mutations in the gene encoding the latency-associated peptide of TGF-beta 1 cause Camurati-Engelmann disease. Nat. Genet. 26, 273–275. 95. Stein, S. S., Witkop, C., Hill, S., Fallon, M. D., Viernstein, L., Gucer, G., McKeever, P., Long, D., Altman, J., Miller, N. R., Teitelbaum, S. L., and Schlesinger, S. (1983). Sclerosteosis: Neurogenetic and pathophysiologic analysis of an American kinship. Neurology 33, 267–277. 96. Van Hul, W., Balemans, W., Van Hul, E., Dikkers, F. G., Obee, H., Stokroos, R. J., Hildering, P., Vanhoenacker, F., Van Camp, G., and Willems, P. J. (1998). Van Buchem disease (hyperostosis corticalis generalisata) maps to chromosome 17q12–q21. Am. J. Hum Genet. 62, 391–399. 97. Winkler, D. G., Sutherland, M. S. K., Ojala, E., Turcott, E., Geoghegan, J. C., Shpektor, D., Skonier, J. E., Yu, C., and Latham, J. A. (2005). Sclerostin inhibition of wnt-3a-induced C3H10T1/2 cell differentiation is indirect and mediated by bone morphogenetic proteins J. Biol. Chem. 280, 2498–2502. 98. Balemans, W., Patel, N., Ebeling, M., Van Hul, E., Wuyts, W., Lacza, C., Dioszegi, M., Dikkers, F. G., Hildering, P., Willems, P. J., Verheij, J. B., Lindpaintner, K., Vickery, B., Foernzler, D., and Van Hul, W. (2002). Identification of a 52 kb deletion downstream of the SOST gene in patients with van Buchem disease. J. Med. Genet. 39, 91–97. 99. Rickels, M. R., Zhang, X., Mumm, S., and Whyte, M. P. (2005). Skeletal disease accompanying high bone mass and novel LRP5 mutation. J. Bone Miner. Res. (in press). 100. Villareal, D. T., Murphy, W. A., Teitelbaum, S. L., Arens, M. Q., and Whyte, M. P. (1992). Painful diffuse osteosclerosis after intravenous drug abuse. Am. J. Med. 93, 371–381. 101. Shaker, J. L., Reinus, W. R., and Whyte, M. P. (1998). Hepatitis C-associated osteosclerosis: Late onset after blood transfusion in an elderly woman. J. Clin. Endocrinol. Metab. 84, 93–98. 102. Khosla, S., Hassoun, A. A. K., Baker, B. K., Liu, F., Zien, N. N., Whyte, M. P., Reasner, C. A., Nippoldt, T. B., Tiegs, R. D., Hintz, R. L. and Conover, C. A. (1998). Insulin-like growth factor system abnormalities in hepatitis C-associated osteosclerosis: A means to increase bone mass in adults? J. Clin. Invest. 101, 2165–2173.

829 103. Sinha, G. P., Curtis, P., Haigh, D., Lealman, G. T., Dodds, W., and Bennett, C. P. (1997). Pachydermoperiostosis in childhood. B. J. Rheumatol. 36, 1224–1227. 104. Wegrowski, Y., Gillery, P., Serpier, H., Georges, N., Combemale, P., Kalis, B., and Maquart, F. X. (1996). Alteration of matrix macromolecule synthesis by fibroblasts from a patient with pachydermoperiostosis. J. Invest. Dermatol. 106, 70–74. 105. Wenkert, D., and Whyte, M. P. (2005). Calcinosis and heterotopic ossification. In “Bone Disease In Rheumatology” (M. Maricic, O. S. Gluck, eds.), pp. 197–202. Lippincott Williams & Wilkins, Philadelphia, PA. 106. Di Martino, S. J., DiCarlo, E. F., Boskey, A. L., and Whyte, M. P., Fibrodysplasia ossificans progressiva: middle-age onset with poor response to aggressive immunosuppressive therapy (in press). 107. Shafritz, A. B., and Kaplan, F. S. (1998). Differential expression of bone and cartilage related genes in fibrodysplasia ossificans progressiva, myositis ossificans progressiva, myositis ossificans traumatica, and osteogenic sarcoma. Clin. Orthop. Rel. Res. 346, 46–52. 108. Eddy, M. E., Leelawattana, R., McAlister, W. H., and Whyte, M. P. (1997). Calcinosis universalis complicating juvenile dermatomyositis: Resolution during probenecid therapy. J. Clin. Endocrinol. Metab. 82, 3536–3542. 109. Perez, M. D., Abrams, S. A., Koenning, G., Stoff, J. E., O’Brien, K. O., and Ellis, K. J. (1994). Mineral metabolism in children with dermatomyositis. J. Rheumatol. 21, 2364–2369. 110. Marie, P. J., de Pollak, C., Chanson, P., and Lomri, A. (1997). Increased proliferation of osteoblastic cells expressing the activating Gs alpha mutation in monostotic and polyostotic fibrous dysplasia. Am. J. Pathol. 150, 1059–1069. 111. Weinstein, L. S., Shenker, A., Gejman, P. V., Merino, M. J., Friedman, E., and Spiegel, A. M. (1991). Activating mutations of the stimulatory G protein in the McCune-Albright syndrome. N. Engl. J. Med. 325, 1688–1695. 112. Yamamoto, T., Ozono, K., Kasayama, S., Yoh, K., Hiroshima, K., Takagi, M., Matsumoto, S., Michigami, T., Yamaoka, K., Kishimoto, T., and Okada, S. (1996). Increased IL-6-production by cells isolated from the fibrous bone dysplasia tissues in patients with McCuneAlbright symdrome. J. Clin. Invest. 98, 30–35. 113. Riminucci, M., Fisher, L. W., Shenker, A., Spiegel, A. M., Bianco, P., and Gehron-Robey, P. (1997). Fibrous dysplasia of bone in the McCune-Albright syndrome: abnormalities in bone formation. Am. J. Pathol. 151, 1587–1600. 114. Chapurlat, R. D., Delmas, P. D., Liens, D., and Meunier, P. J. (1997). Long-term effects of intravenous pamidronate in fibrous dysplasia of bone. J. Bone Miner. Res. 12, 1746–1752. 115. Johansson, C., Roupe, G., Lindstedt, G., and Mellström, D. (1996). Bone density, bone markers and bone radiological features in mastocytosis. Age Ageing 25, 1–7. 116. Fallon, M. D., Whyte, M. P., and Teitelbaum, S. L. (1981). Systemic mastocytosis associated with generalized osteopenia: histological characterization of the skeletal lesion using undecalcified bone from two patients. Hum. Pathol. 12, 813–820. 117. Pullarkat, V.A., Bueso-Ramos, C., Lai, R., Kroft, S., Wilson, C. S., Pullarkat, S. T., Bu, X., Thein, M., Lee, M., and Brynes, R. K. (2003). Systemic mastocytosis with associated clonal hematological nonmast-cell lineage disease: analysis of clinicopathologic features and activating c-kit mutations. A. J. Hematol. 73, 12–17.

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Chapter 50

Osteogenesis Imperfecta FRANCIS H. GLORIEUX AND FRANK RAUCH

VI. Bisphosphonate Therapy in OI VII. Medical Therapies Other than Bisphosphonates VIII. Potential Future Therapies IX. Conclusions References

I. Introduction II. Classification III. Diagnosis IV. Differential Diagnosis V. Pathogenesis

I. INTRODUCTION

there are many patients with type I collagen gene mutations who do not exhibit these secondary characteristics. In addition, the OI patient may present with Wormian bones in the sutures of the skull, and may be of decreased height and have skeletal deformity. OI has been recognized as a disease entity since the 17th century, when it was termed congenital osteomalacia. The term osteogenesis imperfecta was adopted in the late 19th century, and by the start of the 20th century the disorder had been subclassified into congenita and tarda forms to reflect the severe or mild disease that may be present in the young juvenile. As more patients with OI were investigated, the broad clinical spectrum of the disease was more fully recognized and more comprehensive classification systems were adopted. The system in common use today was presented by Sillence et al. and subdivides patients into four types based on disease severity and progression (Table I) [2]. However, it is appreciated that in reality the disorder represents a continuum of severity and that patients do not always fall conveniently into one clinical category. The application of bone histology to OI has also revealed that individuals with

Osteogenesis imperfecta (OI) is a heritable disorder of bone formation that may affect more than 1:10 000 individuals. It is characterized by bone fragility due to low bone mass giving an increased fracture incidence [1]. The bone fragility has led to the adoption of the lay name of “brittle bone disease”. The heritable nature of the disorder distinguishes it from idiopathic juvenile osteoporosis, although clinical osteoporosis is also a consequence of OI. Patients with OI do not have perturbations in serum calcium and vitamin D metabolite levels as a consequence of their disease. Patients with OI may present with other clinical features, but these are by no means universal. Such features include the presence of blue sclera, dentinogenesis imperfecta, skin hyperlaxity, and joint hypermobility. These are all features that one might expect to be associated with a disorder involving type I collagen, though such a relationship is not absolute. While individuals with OI having blue sclera and dentinogenesis imperfecta commonly have mutations in one of their type I collagen genes, Dynamics of Bone and Cartilage Metabolism

Genetics Unit, Shriners Hospital for Children, 1529 Cedar Avenue, Montréal, Québec, Canada H3G 1A6

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Type

Clinical severity

I II

Mild non-deforming OI Perinatal lethal

III

Severely deforming

IV

Moderately deforming

V

Moderately deforming

VI

Moderately to severely deforming

VII

Moderately deforming

Expanded Sillence Classification of OI Typical features

Typically associated mutations

Normal height or mild short stature; blue sclera; no DI Multiple rib and long-bone fractures at birth; marked deformities; broad long bones; low density of skull bones on X-rays; dark sclera Very short; triangular face; severe scoliosis; grayish sclera; DI Moderately short; mild to moderate scoliosis; grayish or white sclera; DI Mild to moderate short stature; dislocation of radial head; mineralized interosseous membrane; hyperplastic callus; white sclera; no DI Moderately short; scoliosis; accumulation of osteoid in bone tissue, fish scale pattern of bone lamellation; white sclera; no DI Mild short stature; short humeri and femora; coxa vara; white sclera; no DI

Premature stop codon in COL1A1 Glycine substitutions in COL1A1 or COL1A2 Glycine substitutions in COL1A1 or COL1A2 Glycine substitutions in COL1A1 or COL1A2 Unknown

Unknown

Unknown

Note: The “typically associated mutations” may or may not be detectable in a given patient. DI: dentinogenesis imperfecta.

similar clinical presentations may exhibit vastly different changes in their bone architecture. This has led to further subdivisions of the disorder, with seven subtypes now having been defined. One feature that is apparent from the preceding paragraph is that OI has never been defined using gene mutation as a criterion. Yet there has been a tendency in the past decade to try to redefine OI as a type I collagenopathy [3, 4]. This would imply that most if not all OI must arise from a mutation in either of the two genes for type I collagen, COL1A1 or COL1A2. This is not the case, and while the majority of OI may arise from mutations in these genes, there are a considerable number of individuals in whom such mutations are absent and mutation in a different gene is the causative event [5–7]. A priori there is no reason why mutations in different genes cannot give rise to the same clinical phenotype, and therefore the definition of OI should remain a clinical one. However, it is certainly possible that in the future genetic diversity may result in a further subdivision of the disease classification.

II. CLASSIFICATION Even though the spectrum of clinical severity in OI is a continuum, categorizing patients into separate types can be useful to assess prognosis and to help evaluate the effects of therapeutic interventions. The classification system of Sillence et al. divides OI into four severity-based types, with the type IV category representing the clinically most diverse group [2]. It is from this heterogeneous

group that OI types V, VI, and VII have been identified as separate entities on the basis of distinct clinical and bone histological features [8–10]. The clinically most relevant characteristic of all types of OI is bone fragility, the severity of which increases in the order type I < types IV, V, VI, VII < type III < type II. Heredity follows an autosomal dominant pattern in OI types I through V and is autosomal recessive in types VI and VII. Mutations affecting collagen type I are usually present in OI types I through IV, but are absent in types V, VI, and VII.

A. Type I OI Type I OI is the mildest form of the disease, being non-deforming and resulting in patients attaining a normal height. However, vertebral fractures are common and can lead to mild scoliosis. Patients typically have blue sclera, but dentinogenesis imperfecta is rarely evident clinically. Fractures are not commonly observed at birth, but begin with ambulation and subsequent falls during juvenile development, then commonly decrease following puberty.

B. Type II OI Type II OI is the most severe form of the disease, resulting in death in the perinatal period, with patients rarely surviving for more than a few days. These individuals exhibit multiple intrauterine rib and long bone fractures,

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and show severe skeletal deformity, which eventually results in respiratory failure. Bone histology reveals a marked decrease in both cortical bone thickness and the amount of trabecular bone.

tomography can be useful to distinguish hyperplastic callus from osteosarcoma in unclear cases [13, 14]. Patients with OI type V appear to constitute between 4% and 5% of the OI population seen in hospitals [1].

C. Type III OI

F. Type VI OI

Type III OI is the most severe form of the disease compatible with survival past the perinatal period. It is characterized by severe progressive skeletal deformity that may have begun by birth. Individuals may possess multiple fractures at birth and suffer frequent fractures thereafter because of the highly fragile nature of their bones. The incidence of fractures remains high even in adult life. Individuals have severely short stature and because of their deformity and bone fragility are frequently confined to a wheelchair for life. Scoliosis can lead to respiratory problems, which have been identified as a leading cause of death in this patient group [11, 12]. Dentinogenesis imperfecta is commonly present.

Type VI OI patients also present with moderate to severe skeletal deformity and do not have blue sclera or dentinogenesis imperfecta [8]. The distinctive features of this OI type are the fish scale-like appearance of the bone lamellae and the presence of excessive osteoid upon histological examination. Although the osteoid accumulation suggests a mineralization defect reminiscent of osteomalacia, there is no abnormality in calcium, phosphate, parathyroid hormone, or vitamin D metabolism, and growth plate mineralization proceeds normally. Inheritance probably is autosomal recessive. OI type VI may be present in about 4% of moderately to severely affected OI patients [1].

D. Type IV OI

G. Type VII OI

Type IV OI is clinically the most diverse group. The phenotype can vary from severe to mild, with the more severely affected patients presenting with fractures at birth, suffering moderate skeletal deformity, and attaining a relatively short stature.

Type VII OI patients also have moderate to severe skeletal deformity and bone fragility, and lack blue sclera and dentinogenesis imperfecta. The distinctive clinical feature of the disease is a rhizomelic shortening of the humerus and femur [10]. Coxa vara may be present even in infancy. So far this disorder has been observed only in a community of Native Americans in northern Quebec. Type VII OI exhibits autosomal recessive inheritance. The disease has been localized to chromosome 3p22–24.1, which is outside the loci for collagen type I genes [7].

E. Type V OI Type V OI is moderately deforming, and patients suffer from moderate to severe bone fragility [9]. Blue sclera and dentinogenesis imperfecta are not present. The patients are characterized by three distinctive features: the presence of hypertrophic callus formation at fracture sites, calcification of the interosseous membranes between the bones of the forearm, and the presence of a radio-opaque metaphyseal band immediately adjacent to the growth plates on X-ray. Upon histological examination, the lamellar organization of the bone has an irregular mesh-like appearance, which is distinct from the normal lamellar organization. The calcified interosseous membrane often severely limits the pronation/supination movement of the hand and may lead to secondary dislocation of the radial head. Importantly, the hyperplastic callus that may develop after fractures or surgical interventions may mimic osteosarcoma. Magnetic resonance imaging and computed

III. DIAGNOSIS The clinical diagnosis of OI is based primarily on the signs and symptoms enumerated above. Traditionally, much emphasis has been laid on the presence or absence of blue sclera and dentinogenesis imperfecta as diagnostic signs of OI. This still holds true, but some limitations should be recognized. Dark or bluish sclera are very common in healthy infants and therefore this finding is not of much diagnostic use in this age group. Dentinogenesis imperfecta is more often clinically evident in primary than in permanent teeth of OI patients [15]. Even in patients whose teeth look normal on inspection, radiological or histological examinations frequently show abnormalities [16–18].

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Clinically evident hearing loss is rare in the first two decades of life, even though subtle audiometric abnormalities can be found in a large percentage of children and adolescents with OI [19–21]. About half of the patients above age 50 report hearing loss, and an even higher percentage of adults have clearly pathologic audiometric findings [21, 22]. Diagnosing OI is straightforward in cases with a positive family history or where several typical features are present, but can be difficult in the absence of affected family members and when bone fragility is not associated with obvious extraskeletal abnormalities. The uncertainty in such cases is compounded by the fact that there are no agreed minimal criteria that establish a clinical diagnosis of OI. In that situation, analysis of the collagen type I genes may provide helpful information. This can be done by examining the amount and structure of type I procollagen molecules that are derived from the patient’s cultured skin fibroblasts [23]. Alternatively, genomic DNA can be extracted from white blood cells and the coding region of the COL1A1 and COL1A2 genes can then be screened for mutations [24]. Both of these approaches are believed to detect almost 90% of all collagen type I mutations [25]. A positive collagen type I study thus confirms the diagnosis of OI. However, a negative result leaves open the possibilities that either a collagen type I mutation is present but was not detected, or that the patient has a form of OI that is not associated with collagen type I mutations (see below). Therefore, a negative collagen type I study does not rule out OI. (For biochemical parameters of mineral homeostasis and bone turnover, see Chapter 53.)

IV. DIFFERENTIAL DIAGNOSIS A number of primary skeletal disorders can be confused with OI. The clinical resemblance is highlighted by the fact that Bruck syndrome and osteoporosispseudoglioma syndrome have previously been called “OI with congenital joint contractures” and “ocular form of OI”, respectively [26, 27]. Panostotic fibrous dysplasia is the extreme form of polyostotic fibrous dysplasia, where all bones are affected [28]. Idiopathic autosomal recessive hyperphosphatasia, also known as juvenile Paget’s disease, is characterized by markedly increased bone turnover [29, 30]. It is usually easily distinguishable from OI on the basis of extremely high serum alkaline phosphatase activity. Hypophosphatasia is extremely variable in clinical expression, ranging from stillbirth without mineralized bone to pathologic fractures which develop only late in adulthood [30]. Cole–Carpenter syndrome has been

described only in a few patients and therefore the mode of inheritance is not established. Idiopathic juvenile osteoporosis is a transient, non-hereditary form of childhood osteoporosis without extraskeletal involvement that typically develops in a prepubertal, previously healthy child of either sex [31]. Spontaneous recovery is the rule after 3–5 years of evolution, although spine deformities and severe functional impairment may persist [31]. Child abuse is a frequent cause of fractures, with the highest incidence in the first year of life [32]. The clinical differentiation of mild OI from child abuse can be difficult at times, especially if the family history is negative for OI. Bone mineral density examinations using dual energy X-ray absorptiometry or computed tomography have been proposed to help in the differential diagnosis [33, 34]. However, there is little information on the range of bone mineral density that is to be expected in infants with mild OI. Collagen type I analysis can be very useful when the test is unequivocally positive, thus proving OI [25]. However, a negative collagen type I analysis evidently does not prove child abuse. Thus, in many cases the distinction between mild OI and child abuse still relies entirely on careful clinical evaluation.

V. PATHOGENESIS This section deals with those forms of OI that are positive for collagen type I mutations, as little is known on the pathogenesis of the other types of the disease. A collagen type I molecule comprises three polypeptide chains (two alpha 1 and one alpha 2 chain) which form a triplehelical structure [35] (see also Chapter 1). For the three chains to intertwine correctly, they must have a glycine residue at every third position. The most frequently encountered types of sequence abnormality associated with OI are point mutations that affect a glycine residue in either COL1A1 or COL1A2. Cells harboring such a mutation produce a mixture of normal and abnormal collagen [36]. The resulting phenotype can vary from very mild to lethal, depending on which of the two alpha chains is affected, at what position in the triple helix the substitution occurs and by what amino acid glycine is substituted. Until now, the genotype–phenotype correlations are too loose to predict the phenotypic effect of a particular glycine mutation with certainty. Mutations that create a premature stop codon within COL1A1 have a more predictable outcome, as in most cases they result in an OI type I phenotype [37]. The transcription products of genes harboring such a mutation are usually unstable and are destroyed by a process called

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nonsense-mediated decay [38]. Consequently, only normal collagen type I chains are produced by fibroblasts of affected individuals, but the rate of collagen production is reduced [36]. Most of these molecular studies have used skin fibroblasts to examine collagen production. Much less is known about the effect of mutations on osteoblasts, which may differ from fibroblasts with regard to post-translational modifications of mutated collagen [39–43] as well as in the propensity to incorporate abnormal collagen molecules into extracellular matrix [44]. For the vast majority of mutations it is unknown how osteoblasts process mutated gene products, how much mutated protein is secreted and whether it is incorporated into organic bone matrix. Osteoblasts harboring a mutated collagen type I gene may have an abnormal expression pattern of other matrix proteins, such as proteoyglycans, hyaluronan, decorin, fibronectin, and thrombospondin [45–47]. These abnormalities in organic composites also affect the mineral phase. Compared to age-matched controls, bone from OI patients shows a higher average mineralization density [48]. The oim mouse, a model of moderate to severe OI, has smaller and less-well-aligned mineral crystals than normal mice [49, 50]. The disturbances in the organic and mineral bone composites are associated with altered biomechanical behavior. Collagen from oim mice has reduced tensile strength [51]. Mineralized OI bone may be harder at the material level [50], but it breaks more easily when deformed, and fatigue damage accumulates much faster on repetitive loading [52]. The sum of these abnormalities may explain the brittleness of OI bone. In addition, OI is characterized by an insufficient amount of bone. Both cortical thickness and the amount of trabecular bone are low [53].

VI. BISPHOSPHONATE THERAPY IN OI Physiotherapy, rehabilitation, and orthopedic surgery are the mainstay of treatment in OI [54, 55]. Therapeutic efforts aim at maximizing mobility and other functional capabilities [54]. Physical activity programs are encouraged (as far as is compatible with the increased risk of fracture) to prevent contractures and immobility-induced bone loss [55]. Orthoses are used to protect the lower limbs during the earlier phases of mobilization [56]. Standing and walking can often only be achieved after femora and tibiae have been straightened using intramedullary rods [55, 57]. This traditional treatment approach can be successful, but does not alter the often

extreme bone fragility in these patients. For this reason, there has been a longstanding search for medical approaches to strengthen the bones. Bisphosphonates are potent antiresorptive agents that inhibit osteoclast function. The hypothesis initially underlying the use of an antiosteoclast medication in an osteoblast disorder such as OI was that a decrease in the activity of the bone-resorbing system might compensate for the weakness of the bone-forming cells. After a case report on the use of oral pamidronate in a child with OI appeared in 1987 [58], various investigators started to treat small groups of pediatric OI patients with bisphosphonates. The use of these drugs in OI and other pediatric disorders became more widespread after the 1998 publication of a larger series of children and adolescents with OI who had been treated with cyclical intravenous pamidronate [59]. Since then a number of groups have reported on their experience with intravenous pamidronate and, more recently, oral forms of bisphosphonate treatment [60].

A. Intravenous Pamidronate The majority of OI patients who were described in published reports received cyclical intravenous pamidronate. None of these studies compared various dosing regimens against each other and therefore the optimal dose of pamidronate and the best treatment interval are unknown. Nevertheless, investigators agreed that intravenous pamidronate infusions, given every 1–4 months, led to a marked and rapid decrease of chronic bone pain, an increased sense of well-being and a rapid rise in vertebral bone mineral mass. Collapsed vertebral bodies were also noted to regain a more normal size and shape [59, 61–64]. The two largest studies reported improved mobility in more than half of the patients [59, 61]. It is unknown at present whether pamidronate treatment prevents long-bone deformities or delays the progression of scoliosis. Histomorphometric studies of iliac bone samples showed that the main effect of pamidronate treatment was to increase cortical thickness [65]. The amount of trabecular bone also increased, which was due to a higher number of trabeculae. In contrast, pamidronate therapy had no detectable effect on trabecular thickness. With regard to long-term safety, there was initially a great deal of concern regarding the effect on growth, given the well-known growth-suppressive effect of highdose bisphosphonates in animals. Fortunately, no negative effect of pamidronate on growth has been detected in children with moderate to severe OI [61, 66].

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B. Other Intravenously Administered Bisphosphonates Neridronate, a bisphosphonate which is similar to pamidronate, was used in an open-label controlled study on adults with OI [67]. This resulted in a significant increase in areal bone mineral density. Zoledronate is a newer intravenously applied bisphosphonate that has been used to treat postmenopausal osteoporosis. Its utility for the treatment of children and adolescents with OI is currently under investigation in an international multicenter trial.

large glass of water with the pill, staying in an upright position for at least 30 minutes thereafter). However, oral therapy also has a number of drawbacks, such as uncertain compliance, low and variable bioavailability, as well as the possibility of gastrointestinal side effects. Oral treatment exposes the skeleton to frequent small doses of medication, whereas intravenous treatment acts with lower frequency but at higher doses. In growing children this difference leads to specific radiographic features: oral treatment causes a continuous dense band in metaphyses, whereas intravenous treatment leads to discrete metaphyseal lines (Fig. 1) [71]. It is currently unknown whether other skeletal effects differ between oral and intravenous bisphosphonate therapy in OI patients.

C. Oral Bisphosphonate Therapy In an observational trial, 15 children and adolescents received alendronate pills at a dose of 1 mg/kg per week in three to seven doses per week, combined with calcitriol 0.25 µg once or twice per week [68]. This treatment was reported to increase lumbar spine areal bone mineral density and decrease the number of fractures, compared to the pretreatment period. A double-blind placebo-controlled trial in 139 children and adolescents found that 2 years of oral alendronate significantly decreased bone turnover, increased spine bone mineral density, and was generally well tolerated [69]. However, no significant effect on the incidence of fractures, bone pain, or functional status was evident. Sakkers et al. tested oral olpadronate at a daily dose of 10 mg per m2 body surface area in a randomized placebocontrolled study that comprised 34 children and adolescents with OI. After a treatment period of 2 years, the group receiving active therapy had a higher lumbar spine areal bone mineral density and a lower incidence of longbone fractures. No difference in functional outcome, such as mobility and muscle force, was detected [70].

E. Who Should Receive Bisphosphonates? The observational trials on pamidronate discussed earlier have evolved from the compassionate use of this drug in desperate cases. As experience with this treatment approach increased, it was also used in less severe (“moderate”) forms of OI. Nevertheless, most reports on

D. Intravenous or Oral Bisphosphonate Therapy? There is currently little published evidence to help in the selection among different bisphosphonate regimens for an OI patient. No study has directly compared the efficacy and safety of different therapeutic approaches. Nevertheless, it is the impression of many clinicians that intravenous pamidronate has a more marked effect on bone pain than oral bisphosphonate therapy. Oral medication has obvious practical advantages over intravenous infusions, at least in patients who are able to swallow pills and who can take the precautions that are required with oral bisphosphonates (such as drinking a

Figure 1

Radiograph of the wrist of a 14-year-old girl with OI type IV. Cyclical intravenous pamidronate had been given every 4 months from age 6 to age 12 years. Metaphyseal lines corresponding to the last few treatment cycles are clearly visible in the radius. Bone that was added after the last pamidronate cycle does not contain any transverse lines and appears to be of normal density.

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intravenous pamidronate therapy state that this treatment was offered only to patients who had long-bone deformities, vertebral compression fractures, and frequent fractures. The treatment decisions of most investigators thus appear to be dictated by clinical severity rather than collagen mutation status, bone mineral density, or OI type. However, given that OI types III and IV (as well as the newly described types V and VI) represent the more severe part of the spectrum of the disorder, most patients who are classified in these categories will probably fulfill the stated criteria for intravenous pamidronate treatment. The results from studies in moderate to severe OI cannot be simply extrapolated to mild forms of the disease (with two or fewer fractures per year, no vertebral compression fractures, and no long-bone deformities). Children with mild OI have less to gain from therapy than severely affected patients, simply because their functional status is better even without treatment. Therefore, the fact that the long-term side effects of bisphosphonate therapy in growing subjects are unknown weighs more heavily in the risk–benefit balance of mildly affected patients. In our view, bisphosphonate therapy is not justified in children with mild forms of OI unless ongoing placebo-controlled trials can establish the efficacy and safety of this approach in this particular patient group. It is unclear what, if any, role bone mineral density measurements should have in the decision to treat an OI patient with bisphosphonates. The many limitations of bone mineral density measurements by dual-energy X-ray absorptiometry, in particular the dependency of results on bone size, have been highlighted in a number of recent reviews [72, 73]. In moderate to severely affected OI patients, lumbar spine areal bone mineral density is usually very low, but such patients would receive treatment based on their clinical picture alone. In milder cases bone mineral density may also be low, but the predictive value of such results in OI patients is not established. Despite the lack of evidence that bone density results actually provide relevant information in OI patients, a clinician may feel that stagnant or decreasing bone density values in a growing child should be taken as an additional argument for starting bisphosphonate therapy. Nevertheless, it is unclear whether there is any benefit in treating young and asymptomatic OI patients whose only “problem” is low bone density.

F. At What Age Should Bisphosphonates be Given? Most of the patients described in the published studies were above 2 years of age when pamidronate treatment

was started. In a congenital disease such as OI, it appears logical to start treatment as early as possible. Indeed, promising results were reported in a small group of patients who received pamidronate in the first 2 years of life [64]. As the clinical effect of a pamidronate infusion, especially on bone pain, was more short-lived in these young children than in older patients, pamidronate cycles were repeated more frequently [1]. The effect of bisphosphonates on the skeleton is clearly growth-dependent [65]. Therefore, postpubertal adolescents and adults cannot be expected to benefit as much from treatment as do younger patients. Nevertheless, two studies suggest that adults with OI also may have some benefit from intravenous pamidronate or neridronate, a bisphosphonate which is similar to pamidronate [67, 74]. In the larger of these, Adami et al. found that intravenous neridronate induced a significant increase in areal bone mineral density at the spine and at the hip. The incidence of fractures was significantly lower during than before treatment.

G. How Long Should Bisphosphonate Treatment be Given? Should the treatment be stopped at all? If so, when? There is little published evidence that would allow answering these questions in one way or another. However, one might try to tackle the problem by addressing it from three different sides: (1) What is the benefit of prolonged therapy? (2) What are the drawbacks of continuing therapy? (3) What is the effect of discontinuing therapy? Regarding the first question, no study has specifically looked at the benefit of long-term therapy, as these reports invariably focused on the initial treatment effect. However, a densitometric study on 56 pediatric OI patients indicated that the rate of change in bone density is slowing down with the duration of pamidronate therapy [62]. For example, the age-specific Z score for lumbar spine areal bone mineral density increased by 2.0 during the first 2 years of treatment, but only by 0.6 between 2 years and 4 years of treatment. Similarly, histomorphometric studies have shown that the cortical width of iliac bone almost doubles during the first 2.4 years of pamidronate treatment, but changes little when therapy is continued for another 3 years thereafter (see reference [65] and unpublished observations) (Fig. 2). As to question 2, it is an inevitable drawback of antiresorptive drugs such as bisphosphonates to decrease the activity of bone remodeling and modeling (shaping) [62, 75]. This was highlighted in a recent case report of a teenage boy who for unclear reasons received massive

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asymptomatic hypocalcemia on the first day of life and the other had bilateral talipes equinovarus. However, it is unclear whether these adverse events in the babies were related to the previous pamidronate treatment of the mothers. It will be important to collect such cases in far greater number to gain some certainty on the effect of bisphosphonate treatment on future pregnancies. Concerning the effect of discontinuing therapy, there is no published information on this question. However, we are following a group of young OI patients who have stopped pamidronate treatment between 1 and 3 years ago. In most of these children and adolescents lumbar spine areal bone mineral density remained stable or continued to increase after pamidronate was stopped. However, a few patients requested that pamidronate treatment be restarted, as they complained about a lack of stamina and recurrence of bone pain after they had been off pamidronate for several months.

H. Bisphosphonates in OI: Conclusions

Figure 2 Series of iliac bone samples in a girl with OI type I. (A) Before treatment (cortical width 505 µm, cancellous bone volume 4.5%). (B) After 2.4 years of pamidronate treatment (cortical width 1686 µm, cancellous bone volume 19.0%). (C) After 5.3 years of pamidronate treatment (cortical width 2025 µm, cancellous bone volume 21.9%).

doses of pamidronate over a period of 3 years and who developed abnormally shaped long-bone metaphyses [76]. A sustained decrease in remodeling activity during growth may also be harmful, as it can lead to the accumulation of growth plate residues within trabecular bone tissue [65, 76]. Calcified cartilage has a high mineral density and therefore contributes to increased densitometric results, but is less resistant to fractures than is normal bone. Low remodeling activity might also delay bone healing after injury. In fact, pamidronate treatment delayed the healing of osteotomy sites after intramedullary rodding procedures [77]. This can lead to pain and fracture at the affected site and may necessitate further surgical procedures. Bisphosphonates are known to persist in bone tissue for many years. Therefore, bisphosphonate treatment of girls and premenopausal women might have an effect on future pregnancies. In two young women with OI who received intravenous pamidronate before conception no maternal ill effects were detected [77]. Both of their babies inherited OI from their mother. One baby had transient

Bisphosphonate therapy does not constitute a cure of OI, but rather is an adjunct to physiotherapy, rehabilitation, and orthopedic care. These drugs have brought clear improvements to the lives of patients suffering from moderate to severe OI. In contrast, children and adolescents with OI who have few fractures and no functional limitations have less to gain from treatment. It therefore appears advisable not to treat such patients unless clinical benefit can be demonstrated in placebocontrolled studies.

VII. MEDICAL THERAPIES OTHER THAN BISPHOSPHONATES Growth hormone has long been proposed as a possible treatment for OI [78]. Small studies suggest that growth hormone treatment may accelerate short-term height velocity in some patients [79–81]. Calcium kinetic studies after 1 year of growth hormone therapy revealed that bone turnover had increased, but that calcium retention was unchanged compared to the pretreatment situation [80]. Increased bone turnover during growth hormone therapy was also found in histomorphometric studies of iliac bone samples [81]. As bone turnover is already abnormally high in untreated children with OI [53], further stimulation does not appear to be a desirable goal. Possibly, growth hormone would be more useful in combination with bisphosphonate therapy, but this remains to be tested.

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Parathyroid hormone is a potent bone anabolic agent and has been shown to reduce the fracture incidence in postmenopausal osteoporosis [82]. These results made parathyroid hormone look like an attractive candidate for treating children with OI. However, a substantial proportion of young rats receiving parathyroid hormone subsequently developed osteosarcoma [83]. It cannot be excluded that a similar effect could happen in humans [84]. Thus, parathyroid hormone should probably not be used in children until these issues have been resolved.

VIII. POTENTIAL FUTURE THERAPIES Bone marrow stromal cells can differentiate into a variety of cell lineages, including osteoblasts [85]. This observation led to the straightforward hypothesis that transplanting bone marrow stromal cells from healthy subjects might improve the clinical course of OI. For this approach to be successful, intravenously infused bone marrow stromal cells must find their way into the skeleton and differentiate into osteoblasts that start producing normal bone. Also, it must be hoped that the bone produced by transplanted cells is not subjected to the same rapid removal as the bone formed by the patient’s original osteoblasts. These hopes formed the rationale for performing bone marrow transplantation in a small group of children with severe OI [86, 87]. The reported results elicited mixed reactions. Bone marrow transplantation experts were excited, because a small percentage of the patients’ osteoblasts seemed to be of donor origin [88]. Bone disease specialists remained skeptical, because no convincing evidence was presented to show that the patients had actually benefited from the procedure [36, 85, 89–91]. Whatever the clinical effects may have been, the authors conceded that they were short-lived [87]. Therefore, they went on to retreat the same patients with a modified approach. This time isolated marrow stromal cells were infused [92]. Similar to the first study, a small fraction of transplanted cells was detectable in a number of patient tissues, including bone. Clinical benefit was claimed, based mainly on increased growth velocity in the 6 months following the procedure. However, the difficulty of measuring short-term growth velocity in children with bone deformities is obvious. In our view, enthusiasm about an innovative technique should not be a substitute for scientific stringency and objective evaluation of results. Techniques based on marrow stem cells may offer therapeutic potential for OI patients in the future. It is therefore important not to discredit this approach by premature clinical application.

The basic science and technical issues of this approach should be worked out in animal experiments before further studies are performed in humans. Currently available medical treatment options at best achieve symptomatic improvement of OI. The only hope for actually curing the disease is by eliminating the mutated gene or the gene product. Unfortunately, there are major obstacles to gene-based therapy of OI. The majority of severe OI cases result from the presence of abnormal collagen molecules. Thus, it is not sufficient to replace a missing protein, as is the case in many recessive enzyme disorders. Rather, it is necessary to first inactivate the mutant allele and then substitute for its product [93]. Current research is still grappling with the first of these two tasks. Various investigators are studying the use of so-called hammerhead ribozymes [94–96], small RNA molecules that can cut mRNA in the absence of protein cofactors [97]. Various viruses have been tested that allow for transfection of mesenchymal progenitor cells [96, 98]. When ribozymes were transfected into such cells COL1A1 mRNA levels could be suppressed by approximately 50% [96]. Such results are encouraging, but they nevertheless represent only a first step on a long and difficult path towards gene-based therapy of OI.

IX. CONCLUSIONS OI has long been a topic of interest for basic scientists and a source of frustration to clinicians. Much progress has been made concerning the molecular, cellular, and material basis of the disorder, but up to now these insights have contributed little to alleviate the disease burden on affected individuals. Only the recent advent of pamidronate therapy has brought clear improvements to the lives of children and adolescents suffering from moderate to severe OI. Nevertheless, this form of treatment does not constitute a cure, but rather is an adjunct to physiotherapy, rehabilitation, and orthopedic care. Gene-based therapy offers the hope for a curative treatment of OI, but remains at present in the early stages of preclinical research. New therapeutic avenues in severe OI will have to be evaluated against the results obtained with bisphosphonates.

References 1. Rauch, F., and Glorieux, F. H. (2004). Osteogenesis imperfecta. Lancet 363, 1377–1385. 2. Sillence, D. O., Senn, A., and Danks, D. M. (1979). Genetic heterogeneity in osteogenesis imperfecta. J. Med. Genet. 16, 101–116.

840 3. Byers, P. H., and Steiner, R. D. (1992). Osteogenesis imperfecta. Annu. Rev. Med. 43, 269–282. 4. Prockop, D. J., Colige, A., Helminen, H., Khillan, J. S., Pereira, R., and Vandenberg, P. (1993). Mutations in type 1 procollagen that cause osteogenesis imperfecta: effects of the mutations on the assembly of collagen into fibrils, the basis of phenotypic variations, and potential antisense therapies. J. Bone Miner. Res. 8 (Suppl 2), S489–492. 5. Aitchison, K., Ogilvie, D., Honeyman, M., Thompson, E., and Sykes, B. (1988). Homozygous osteogenesis imperfecta unlinked to collagen I genes. Hum. Genet. 78, 233–236. 6. Wallis, G. A., Sykes, B., Byers, P. H., Mathew, C. G., Viljoen, D., and Beighton, P. (1993). Osteogenesis imperfecta type III: mutations in the type I collagen structural genes, COL1A1 and COL1A2, are not necessarily responsible. J. Med. Genet. 30, 492–496. 7. Labuda, M., Morissette, J., Ward, L. M., Rauch, F., Lalic, L., Roughley, P. J., and Glorieux, F. H. (2002). Osteogenesis imperfecta type VII maps to the short arm of chromosome 3. Bone 31, 19–25. 8. Glorieux, F. H., Ward, L. M., Rauch, F., Lalic, L., Roughley, P. J., and Travers, R. (2002). Osteogenesis imperfecta type VI: a form of brittle bone disease with a mineralization defect. J. Bone Miner. Res. 17, 30–38. 9. Glorieux, F. H., Rauch, F., Plotkin, H., Ward, L., Travers, R., Roughley, P., Lalic, L., Glorieux, D. F., Fassier, F., and Bishop, N. J. (2000). Type V osteogenesis imperfecta: a new form of brittle bone disease. J. Bone Miner. Res. 15, 1650–1658. 10. Ward, L. M., Rauch, F., Travers, R., Chabot, G., Azouz, E. M., Lalic, L., Roughley, P. J., and Glorieux, F. H. (2002). Osteogenesis imperfecta type VII: an autosomal recessive form of brittle bone disease. Bone 31, 12–18. 11. McAllion, S. J., and Paterson, C. R. (1996). Causes of death in osteogenesis imperfecta. J. Clin. Pathol. 49, 627–630. 12. Paterson, C. R., Ogston, S. A., and Henry, R. M. (1996). Life expectancy in osteogenesis imperfecta. BMJ 312, 351. 13. Rieker, O., Kreitner, K. F., and Karbowski, A. (1998). Hyperplastic callus formation in osteogenesis imperfecta: CT and MRI findings. Eur. Radiol. 8, 1137–1139. 14. Dobrocky, I., Seidl, G., and Grill, F. (1999). MRI and CT features of hyperplastic callus in osteogenesis imperfecta tarda. Eur. Radiol. 9, 665–668. 15. Petersen, K., and Wetzel, W. E. (1998). Recent findings in classification of osteogenesis imperfecta by means of existing dental symptoms. ASDC J. Dent. Child. 65, 305–309. 16. Lygidakis, N. A., Smith, R., and Oulis, C. J. (1996). Scanning electron microscopy of teeth in osteogenesis imperfecta type I. Oral Surg. Oral Med. Oral Pathol. Oral Radiol. Endod. 81, 567–572. 17. Lund, A. M., Jensen, B. L., Nielsen, L. A., and Skovby, F. (1998). Dental manifestations of osteogenesis imperfecta and abnormalities of collagen I metabolism. J. Craniofac. Genet. Dev. Biol. 18, 30–37. 18. Malmgren, B., and Norgren, S. (2002). Dental aberrations in children and adolescents with osteogenesis imperfecta. Acta. Odontol. Scand. 60, 65–71. 19. Pedersen, U. (1984). Hearing loss in patients with osteogenesis imperfecta. A clinical and audiological study of 201 patients. Scand. Audiol. 13, 67–74. 20. Kuurila, K., Grenman, R., Johansson, R., and Kaitila, I. (2000). Hearing loss in children with osteogenesis imperfecta. Eur. J. Pediatr. 159, 515–519. 21. Paterson, C. R., Monk, E. A., and McAllion, S. J. (2001). How common is hearing impairment in osteogenesis imperfecta? J. Laryngol. Otol. 115, 280–282. 22. Kuurila, K., Kaitila, I., Johansson, R., and Grenman, R. (2002). Hearing loss in Finnish adults with osteogenesis imperfecta: a nationwide survey. Ann. Otol. Rhinol. Laryngol. 111, 939–946. 23. Wenstrup, R. J., Willing, M. C., Starman, B. J., and Byers, P. H. (1990). Distinct biochemical phenotypes predict clinical severity in nonlethal variants of osteogenesis imperfecta. Am. J. Hum. Genet. 46, 975–982.

Francis H. Glorieux and Frank Rauch 24. Korkko, J., Ala-Kokko, L., De Paepe, A., Nuytinck, L., Earley, J., and Prockop, D. J. (1998). Analysis of the COL1A1 and COL1A2 genes by PCR amplification and scanning by conformation-sensitive gel electrophoresis identifies only COL1A1 mutations in 15 patients with osteogenesis imperfecta type I: identification of common sequences of null-allele mutations. Am. J. Hum. Genet. 62, 98–110. 25. Marlowe, A., Pepin, M. G., and Byers, P. H. (2002). Testing for osteogenesis imperfecta in cases of suspected non-accidental injury. J. Med. Genet. 39, 382–386. 26. McPherson, E., and Clemens, M. (1997). Bruck syndrome (osteogenesis imperfecta with congenital joint contractures): review and report on the first North American case. Am. J. Med. Genet. 70, 28–31. 27. Beighton, P., Winship, I., and Behari, D. (1985). The ocular form of osteogenesis imperfecta: a new autosomal recessive syndrome. Clin. Genet. 28, 69–75. 28. Cole, D. E., Fraser, F. C., Glorieux, F. H., Jequier, S., Marie, P. J., Reade, T. M., and Scriver, C. R. (1983). Panostotic fibrous dysplasia: a congenital disorder of bone with unusual facial appearance, bone fragility, hyperphosphatasemia, and hypophosphatemia. Am. J. Med. Genet. 14, 725–735. 29. Cundy, T., Hegde, M., Naot, D., Chong, B., King, A., Wallace, R., Mulley, J., Love, D. R., Seidel, J., Fawkner, M., Banovic, T., Callon, K. E., Grey, A. B., Reid, I. R., Middleton-Hardie, C. A., and Cornish, J. (2002). A mutation in the gene TNFRSF11B encoding osteoprotegerin causes an idiopathic hyperphosphatasia phenotype. Hum. Mol. Genet. 11, 2119–2127. 30. Whyte, M. P., Obrecht, S. E., Finnegan, P. M., Jones, J. L., Podgornik, M. N., McAlister, W. H., and Mumm, S. (2002). Osteoprotegerin deficiency and juvenile Paget’s disease. N. Engl. J. Med. 347, 175–184. 31. Smith, R. (1995). Idiopathic juvenile osteoporosis: experience of twenty-one patients. Br. J. Rheumatol. 34, 68–77. 32. Nimkin, K., and Kleinman, P. K. (2001). Imaging of child abuse. Radiol. Clin. North. Am. 39, 843–864. 33. Moore, M. S., Minch, C. M., Kruse, R. W., Harcke, H. T., Jacobson, L., and Taylor, A. (1998). The role of dual energy x-ray absorptiometry in aiding the diagnosis of pediatric osteogenesis imperfecta. Am. J. Orthop. 27, 797–801. 34. Miller, M. E., and Hangartner, T. N. (1999). Bone density measurements by computed tomography in osteogenesis imperfecta type I. Osteoporos. Int. 9, 427–432. 35. Kadler, K. E., Holmes, D. F., Trotter, J. A., and Chapman, J. A. (1996). Collagen fibril formation. Biochem. J. 316 (Pt 1), 1–11. 36. Byers, P. H. (2000). Osteogenesis imperfecta: perspectives and opportunities. Curr. Opin. Pediatr. 12, 603–609. 37. Willing, M. C., Deschenes, S. P., Slayton, R. L., and Roberts, E. J. (1996). Premature chain termination is a unifying mechanism for COL1A1 null alleles in osteogenesis imperfecta type I cell strains. Am. J. Hum. Genet. 59, 799–809. 38. Byers, P. H. (2002). Killing the messenger: new insights into nonsensemediated mRNA decay. J. Clin. Invest. 109, 3–6. 39. Sarafova, A. P., Choi, H., Forlino, A., Gajko, A., Cabral, W. A., Tosi, L., Reing, C. M., and Marini, J. C. (1998). Three novel type I collagen mutations in osteogenesis imperfecta type IV probands are associated with discrepancies between electrophoretic migration of osteoblast and fibroblast collagen. Hum. Mutat. 11, 395–403. 40. Bank, R. A., Robins, S. P., Wijmenga, C., Breslau-Siderius, L. J., Bardoel, A. F., van der Sluijs, H. A., Pruijs, H. E., and TeKoppele, J. M. (1999). Defective collagen crosslinking in bone, but not in ligament or in Bruck syndrome: indications for a bone-specific telopeptide lysyl hydroxylase on chromosome 17. Proc. Natl. Acad. Sci. USA 96, 1054–1058. 41. Cabral, W. A., Chernoff, E. J., and Marini, J. C. (2001). G76E substitution in type I collagen is the first nonlethal glutamic acid substitution in the alpha1(I) chain and alters folding of the N-terminal end of the helix. Mol. Genet. Metab. 72, 326–335.

Chapter 50 Osteogenesis Imperfecta 42. Cabral, W. A., Fertala, A., Green, L. K., Korkko, J., Forlino, A., and Marini, J. C. (2002). Procollagen with skipping of alpha 1(I) exon 41 has lower binding affinity for alpha 1(I) C-telopeptide, impaired in vitro fibrillogenesis, and altered fibril morphology. J. Biol. Chem. 277, 4215–4222. 43. Galicka, A., Wolczynski, S., and Gindzienski, A. (2002). Comparative studies of osteoblast and fibroblast type I collagen in a patient with osteogenesis imperfecta type IV. J. Pathol. 196, 235–237. 44. Mundlos, S., Chan, D., Weng, Y. M., Sillence, D. O., Cole, W. G., and Bateman, J. F. (1996). Multiexon deletions in the type I collagen COL1A2 gene in osteogenesis imperfecta type IB. Molecules containing the shortened alpha2(I) chains show differential incorporation into the bone and skin extracellular matrix. J. Biol. Chem. 271, 21068–21074. 45. Fedarko, N. S., Robey, P. G., and Vetter, U. K. (1995). Extracellular matrix stoichiometry in osteoblasts from patients with osteogenesis imperfecta. J. Bone Miner. Res. 10, 1122–1129. 46. Fedarko, N. S., Sponseller, P. D., and Shapiro, J. R. (1996). Long-term extracellular matrix metabolism by cultured human osteogenesis imperfecta osteoblasts. J. Bone Miner. Res. 11, 800–805. 47. Grzesik, W. J., Frazier, C. R., Shapiro, J. R., Sponsellor, P. D., Robey, P. G., and Fedarko, N. S. (2002). Age-related changes in human bone proteoglycan structure: impact of osteogenesis imperfecta. J. Biol. Chem. 6, 6. 48. Boyde, A., Travers, R., Glorieux, F. H., and Jones, S. J. (1999). The mineralization density of iliac crest bone from children with osteogenesis imperfecta. Calcif. Tissue Int. 64, 185–190. 49. Fratzl, P., Paris, O., Klaushofer, K., and Landis, W. J. (1996). Bone mineralization in an osteogenesis imperfecta mouse model studied by small-angle x-ray scattering. J. Clin. Invest. 97, 396–402. 50. Grabner, B., Landis, W. J., Roschger, P., Rinnerthaler, S., Peterlik, H., Klaushofer, K., and Fratzl, P. (2001). Age- and genotype-dependence of bone material properties in the osteogenesis imperfecta murine model (oim). Bone 29, 453–457. 51. Misof, K., Landis, W. J., Klaushofer, K., and Fratzl, P. (1997). Collagen from the osteogenesis imperfecta mouse model (oim) shows reduced resistance against tensile stress. J. Clin. Invest. 100, 40–45. 52. Jepsen, K. J., Schaffler, M. B., Kuhn, J. L., Goulet, R. W., Bonadio, J., and Goldstein, S. A. (1997). Type I collagen mutation alters the strength and fatigue behavior of Mov13 cortical tissue. J. Biomech. 30, 1141–1147. 53. Rauch, F., Travers, R., Parfitt, A. M., and Glorieux, F. H. (2000). Static and dynamic bone histomorphometry in children with osteogenesis imperfecta. Bone 26, 581–589. 54. Engelbert, R. H., Pruijs, H. E., Beemer, F. A., and Helders, P. J. (1998). Osteogenesis imperfecta in childhood: treatment strategies. Arch. Phys. Med. Rehabil. 79, 1590–1594. 55. Zeitlin, L., Fassier, F., and Glorieux, F. H. (2003). Modern approach to children with osteogenesis imperfecta. J. Pediatr. Orthop. B. 12, 77–87. 56. Gerber, L. H., Binder, H., Berry, R., Siegel, K. L., Kim, H., Weintrob, J., Lee, Y. J., Mizell, S., and Marini, J. (1998). Effects of withdrawal of bracing in matched pairs of children with osteogenesis imperfecta. Arch. Phys. Med. Rehabil. 79, 46–51. 57. Wilkinson, J. M., Scott, B. W., Clarke, A. M., and Bell, M. J. (1998). Surgical stabilisation of the lower limb in osteogenesis imperfecta using the Sheffield Telescopic Intramedullary Rod System. J. Bone Joint Surg. Br. 80, 999–1004. 58. Devogelaer, J. P., Malghem, J., Maldague, B., and Nagant, de Deuxchaisnes, C. (1987). Radiological manifestations of bisphosphonate treatment with APD in a child suffering from osteogenesis imperfecta. Skeletal Radiol. 16, 360–363. 59. Glorieux, F. H., Bishop, N. J., Plotkin, H., Chabot, G., Lanoue, G., and Travers, R. (1998). Cyclic administration of pamidronate in children with severe osteogenesis imperfecta. N. Engl. J. Med. 339, 947–952.

841 60. Rauch, F., and Glorieux, F. H. (2005). Bisphosphonate treatment in osteogenesis imperfecta: which drug, for whom, for how long? Ann. Med. 37, 295–302. 61. Astrom, E., and Soderhall, S. (2002). Beneficial effect of long term intravenous bisphosphonate treatment of osteogenesis imperfecta. Arch. Dis. Child. 86, 356–364. 62. Rauch, F., Plotkin, H., Zeitlin, L., and Glorieux, F. H. (2003). Bone mass, size and density in children and adolescents with osteogenesis imperfecta: Effect of intravenous pamidronate therapy. J. Bone Miner. Res. 18, 610–614. 63. Arikoski, P., Silverwood, B., Tillmann, V., and Bishop, N. J. (2004). Intravenous pamidronate treatment in children with moderate to severe osteogenesis imperfecta: assessment of indices of dual-energy X-ray absorptiometry and bone metabolic markers during the first year of therapy. Bone 34, 539–546. 64. Munns, C. F., Rauch, F., Travers, R., and Glorieux, F. H. (2005). Effects of intravenous pamidronate treatment in infants with osteogenesis imperfecta: clinical and histomorphometric outcome. J. Bone Miner. Res. 20, 1235–1243. 65. Rauch, F., Travers, R., Plotkin, H., and Glorieux, F. H. (2002). The effects of intravenous pamidronate on the bone tissue of children and adolescents with osteogenesis imperfecta. J. Clin. Invest. 110, 1293–1299. 66. Zeitlin, L., Rauch, F., Plotkin, H., and Glorieux, F. H. (2003). Height and weight development during long-term therapy with cyclical intravenous pamidronate in children and adolescents with osteogenesis imperfecta types I, III and IV. Pediatrics 111, 1030–1036. 67. Adami, S., Gatti, D., Colapietro, F., Fracassi, E., Braga, V., Rossini, M., and Adami, S. (2003). Intravenous neridronate in adults with osteogenesis imperfecta. J. Bone Miner. Res. 18, 126–130. 68. Maasalu, K., Haviko, T., and Martson, A. (2003). Treatment of children with osteogenesis imperfecta in Estonia. Acta. Paediatr. 92, 452–455. 69. Glorieux, F. H., Rauch, F., Ward, L. M., Smith, P., Verbruggen, N., Heyden, N., and Lombardi, A. (2004). Alendronate in the treatment of pediatric osteogenesis imperfecta. J. Bone Miner. Res. 19, S12. 70. Sakkers, R., Kok, D., Engelbert, R., van Dongen, A., Jansen, M., Pruijs, H., Verbout, A., Schweitzer, D., and Uiterwaal, C. S. P. M. (2004). Skeletal effects and functional outcome with olpadronate in children with osteogenesis imperfecta: a 2-year randomised placebocontrolled study. Lancet 363, 1427–1431. 71. Rauch, F., Travers, R., Munns, C., and Glorieux, F. H. (2004). Sclerotic metaphyseal lines in a child treated with pamidronate: histomorphometric analysis. J. Bone Miner. Res. 19, 1191–1193. 72. Leonard, M. B. (2003). Assessment of bone health in children and adolescents with cancer: promises and pitfalls of current techniques. Med. Pediatr. Oncol. 41, 198–207. 73. Fewtrell, M. S. (2003). Bone densitometry in children assessed by dual X-ray absorptiometry: uses and pitfalls. Arch. Dis. Child 88, 795–798. 74. Shapiro, J. R., McCarthy, E. F., Rossiter, K., Ernest, K., Gelman, R., Fedarko, N., Santiago, H. T., and Bober, M. (2003). The effect of intravenous pamidronate on bone mineral density, bone histomorphometry, and parameters of bone turnover in adults with type IA osteogenesis imperfecta. Calcif. Tissue Int. 72(2), 103–112. 75. Rauch, F., Plotkin, H., Travers, R., Zeitlin, L., and Glorieux, F. H. (2003). Osteogenesis imperfecta types I, III and IV: effect of pamidronate therapy on bone and mineral metabolism. J. Clin. Endocrinol. Metab. 88, 986–992. 76. Whyte, M. P., Wenkert, D., Clements, K. L., McAlister, W. H., and Mumm, S. (2003). Bisphosphonate-induced osteopetrosis. N. Engl. J. Med. 349, 457–463. 77. Munns, C. F., Rauch, F., Zeitlin, L., Fassier, F., and Glorieux, F. H. (2004). Delayed osteotomy but not fracture healing in pediatric osteogenesis imperfecta patients receiving pamidronate. J. Bone Miner. Res. 19, 1779–1786.

842 78. Kruse, H. P., and Kuhlencordt, F. (1975). On an attempt to treat primary and secondary osteoporosis with human growth hormone. Horm. Metab. Res. 7, 488–491. 79. Antoniazzi, F., Bertoldo, F., Mottes, M., Valli, M., Sirpresi, S., Zamboni, G., Valentini, R., and Tato, L. (1996). Growth hormone treatment in osteogenesis imperfecta with quantitative defect of type I collagen synthesis. J. Pediatr. 129, 432–439. 80. Vieira, N. E., Marini, J. C., Hopkins, E., Abrams, S. A., and Yergey, A. L. (1999). Effect of growth hormone treatment on calcium kinetics in patients with osteogenesis imperfecta type III and IV. Bone 25, 501–505. 81. Marini, J. C., Hopkins, E., Glorieux, F. H., Chrousos, G. P., Reynolds, J. C., Gundberg, C. M., and Reing, C. M. (2003). Positive linear growth and bone responses to growth hormone treatment in children with types III and IV osteogenesis imperfecta: high predictive value of the carboxyterminal propeptide of type I procollagen. J. Bone Miner. Res. 18, 237–243. 82. Neer, R. M., Arnaud, C. D., Zanchetta, J. R., Prince, R., Gaich, G. A., Reginster, J. Y., Hodsman, A. B., Eriksen, E. F., Ish-Shalom, S., Genant, H. K., Wang, O., and Mitlak, B. H. (2001). Effect of parathyroid hormone (1–34) on fractures and bone mineral density in postmenopausal women with osteoporosis. N. Engl. J. Med. 344, 1434–1441. 83. Vahle, J. L., Sato, M., Long, G. G., Young, J. K., Francis, P. C., Engelhardt, J. A., Westmore, M. S., Linda, Y., and Nold, J. B. (2002). Skeletal changes in rats given daily subcutaneous injections of recombinant human parathyroid hormone (1–34) for 2 years and relevance to human safety. Toxicol. Pathol. 30, 312–321. 84. Kuijpers, G., Schneider, B., Stadel, B., and Colman, E. (2002). Recombinant human parathyroid hormone. Preclinical data on rat osteosarcoma were not dismissed. BMJ 324, 1218; discussion. 85. Bianco, P., and Gehron Robey, P. (2000). Marrow stromal stem cells. J. Clin. Invest. 105, 1663–1668. 86. Horwitz, E. M., Prockop, D. J., Fitzpatrick, L. A., Koo, W. W., Gordon, P. L., Neel, M., Sussman, M., Orchard, P., Marx, J. C., Pyeritz, R. E., and Brenner, M. K. (1999). Transplantability and therapeutic effects of bone marrow-derived mesenchymal cells in children with osteogenesis imperfecta. Nat. Med. 5, 309–313.

Francis H. Glorieux and Frank Rauch 87. Horwitz, E. M., Prockop, D. J., Gordon, P. L., Koo, W. W., Fitzpatrick, L. A., Neel, M. D., McCarville, M. E., Orchard, P. J., Pyeritz, R. E., and Brenner, M. K. (2001). Clinical responses to bone marrow transplantation in children with severe osteogenesis imperfecta. Blood 97, 1227–1231. 88. Gerson, S. L. (1999). Mesenchymal stem cells: no longer second class marrow citizens. Nat. Med. 5, 262–264. 89. Marini, J. C. (1999). Osteogenesis imperfecta calls for caution. Nat. Med. 5, 466–467. 90. Bishop, N. J. (1999). Osteogenesis imperfecta calls for caution. Nat. Med. 5, 466–467. 91. Smith, R. (2001). Severe osteogenesis imperfecta: new therapeutic options? BMJ 322, 63–64. 92. Horwitz, E. M., Gordon, P. L., Koo, W. K., Marx, J. C., Neel, M. D., McNall, R. Y., Muul, L., and Hofmann, T. (2002). Isolated allogeneic bone marrow-derived mesenchymal cells engraft and stimulate growth in children with osteogenesis imperfecta: Implications for cell therapy of bone. Proc. Natl. Acad. Sci. USA 99, 8932–8937. 93. Forlino, A., and Marini, J. C. (2000). Osteogenesis imperfecta: prospects for molecular therapeutics. Mol. Genet. Metab. 71, 225–232. 94. Dawson, P. A., and Marini, J. C. (2000). Hammerhead ribozymes selectively suppress mutant type I collagen mRNA in osteogenesis imperfecta fibroblasts. Nucleic Acids Res. 28, 4013–4020. 95. Toudjarska, I., Kilpatrick, M. W., Niu, J., Wenstrup, R. J., and Tsipouras, P. (2001). Delivery of a hammerhead ribozyme specifically downregulates mutant type I collagen mRNA in a murine model of osteogenesis imperfecta. Antisense Nucleic Acid Drug Dev. 11, 341–346. 96. Millington-Ward, S., Allers, C., Tuohy, G., Conget, P., Allen, D., McMahon, H. P., Kenna, P. F., Humphries, P., and Farrar, G. J. (2002). Validation in mesenchymal progenitor cells of a mutation-independent ex vivo approach to gene therapy for osteogenesis imperfecta. Hum. Mol. Genet. 11, 2201–2206. 97. Doudna, J. A., and Cech, T. R. (2002). The chemical repertoire of natural ribozymes. Nature 418, 222–228. 98. Liu, P., Kalajzic, I., Stover, M. L., Rowe, D. W., and Lichtler, A. C. (2001). Human bone marrow stromal cells are efficiently transduced by vesicular stomatitis virus-pseudotyped retrovectors without affecting subsequent osteoblastic differentiation. Bone 29, 331–335.

Chapter 51

Rheumatoid Arthritis and other Inflammatory Joint Pathologies STEVEN R. GOLDRING, M.D. MARY B. GOLDRING, PH.D.

AND

Department of Medicine, Rheumatology Division, Beth Israel Deaconess Medical Center, Harvard Medical School, Boston, MA; New England Baptist Bone and Joint Institute, Boston, MA 02215

I. Abstract II. Introduction III. Effects of Joint Inflammation on Skeletal Remodeling

IV. Effects of Joint Inflammation on Cartilage Remodeling V. Conclusion References

I. ABSTRACT

techniques for assessing bone mass are among the approaches that have been used to assess the effects of the inflammatory joint diseases on local joint structures and generalized skeletal tissue remodeling. This chapter will review the results of these various analytical tools and assays that have been developed to assess the effects of inflammatory conditions on cartilage and bone remodeling and to correlate these biomarkers with standard indices of systemic inflammatory activity and immune system activation.

The inflammatory joint diseases comprise a heterogeneous group of disorders that share in common their propensity to produce destruction of the extracellular matrices of joint cartilage and bone. In addition, the underlying disturbance in immune regulation that is responsible for the localized joint pathology may target extra-articular organs and tissues leading to manifestations in the lung, kidney, central nervous system, and cardiovascular and gastrointestinal systems. Under these circumstances, the generalized systemic inflammatory process may produce generalized effects on bone and connective tissue remodeling that may affect the entire skeleton. Among the inflammatory joint diseases, rheumatoid arthritis, the seronegative spondyloarthropathies, and juvenile rheumatoid arthritis represent excellent models for gaining insights into the effects of local as well as systemic consequences of inflammatory processes on skeletal tissue and cartilage remodeling. Histomorphometric analyses of bone remodeling indices, assessment of the breakdown products of the matrix components of joint cartilage and bone, and quantitative imaging Dynamics of Bone and Cartilage Metabolism

II. INTRODUCTION Rheumatoid arthritis (RA) and related inflammatory joint diseases are a clinically diverse group of disorders that share in common their propensity to affect the anatomic components of articular and juxta-articular tissues. Of the structurally distinct types of joints, the amphiarthrodial and diarthrodial joints are the most frequently affected sites of involvement. The diarthrodial joints are the most common of the articular design patterns [1]. These joints join two opposing bone surfaces that are covered by 843

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Steven R. Goldring and Mary B. Goldring

a specialized hyaline cartilage that provides a low-friction, articulating interface. A highly specialized membrane termed the synovium lines the joint cavity. This membrane is the site of production of synovial fluid that provides the nutritional requirements for the articular cartilage and also “lubricates” the cartilage surfaces. In most forms of inflammatory arthritis, this lining membrane is the site of the initial inflammatory process [2–5]. In contrast to the diarthrodial joints, amphiarthroses lack a true joint cavity and synovial lining and are characterized by a fibrocartilaginous union that joins the two opposing bone surfaces. These joints are best represented by the intervertebral disks. Many of the inflammatory joint diseases are accompanied by generalized systemic symptoms such as fatigue, weight loss, and fever that may dominate the clinical picture. In addition, the underlying disturbance in immune regulation that is responsible for the localized joint pathology may target extra-articular organs and tissues leading to manifestations in the lung, kidney, and cardiovascular, gastrointestinal, and central nervous systems. Under these circumstances, the generalized systemic inflammatory process may produce generalized effects on the remodeling of bone and connective tissues influence the entire skeleton.

III. EFFECTS OF JOINT INFLAMMATION ON SKELETAL REMODELING Among the inflammatory joint diseases, rheumatoid arthritis (RA), the seronegative spondyloarthropathies, and juvenile rheumatoid arthritis represent excellent models for gaining insights into the effects of local as well as systemic consequences of inflammatory processes on skeletal tissue and cartilage remodeling. The effects of RA on skeletal tissues and indices of bone remodeling will be discussed first.

expansion of the synovial lining, there is eventual extension of the inflammatory tissue mass to the adjacent articular cartilage with progressive overgrowth of the articular surface and formation of the so-called pannus, which is derived from the Greek word, meaning “covering” or “mantle”. Directly, at the interface between the RA synovium and articular cartilage, tongues of proliferating cells can be seen penetrating the extracellular matrix of the cartilage. The progression of RA to severe and irreversible joint damage is highly variable from patient to patient. The potential predictors of radiographic progression identified in retrospective and prospective studies have generally included clinical signs of disease activity, systemic indices of inflammation, and radiographic damage [11–15]. Although there is an association between inflammation and the development of joint damage, damage may progress in spite of attenuated inflammatory activity, or erosions may develop in the absence of clinical signs of inflammation [16–19]. The recent development of assays for specific biological markers that reflect quantitative and dynamic changes in the remodeling of joint tissues offers the possibility of identifying patients at risk for rapid joint damage and also for early monitoring of the efficacy of disease-modifying antirheumatic therapies [20–22]. These biomarkers include the synthetic and degradation products of components of the extracellular matrices of the bone and cartilage [23–29], as listed in Table I (see also Chapters 24–27 and 34–37). The specific cellular and biochemical mechanisms that account for the disruption of the integrity of the cartilage matrix and the release of soluble components that can be used to monitor these events are reviewed in the subsequent sections. Similarly, at the interface between the inflamed synovial tissues and adjacent subchondral bone, there is evidence of local activation of bone resorption with destruction of the mineralized bone matrix. The effects of the RA synovial lesion on bone remodeling will be discussed first.

B. Bone Disease Associated with RA A. Rheumatoid Arthritis Although RA may manifest as a systemic disorder, the hallmark of this condition is the presence of a symmetrical inflammatory polyarthritis that targets the proximal small joints of the hands, wrists, elbows, shoulders, knees, ankles, and feet. The synovial lining of diarthrodial joints represents the initial target of the inflammatory joint pathology. This lesion is characterized by proliferation of the synovial lining cells and infiltration of the tissue by inflammatory cells, including lymphocytes, plasma cells, endothelial cells, and activated macrophages [2, 6–10]. With the growth and

Three principal forms of bone disease have been described in patients with RA. The first is characterized by focal bone loss affecting the immediate subchondral bone and bone at the joint margins (Fig. 1). This gives rise to the “bone erosions” that are characteristic of RA and related forms of inflammatory arthritis. The second form of bone disease is manifested by the development of periarticular osteopenia adjacent to inflamed joints. The third form of bone disease in patients with RA is the presence of generalized osteoporosis involving the axial and appendicular skeleton [30–32].

TABLE I.

Molecular Markers of Turnover of Bone, Cartilage, and Synovium in Synovial Fluid and Blood

Molecular Bone Type I collagen

Non-collagenous proteins Cartilage Type II collagen

Synthesis N-propeptide (PINP; S, SF) C-propeptide (PICP; S, SF)

Osteocalcin (S, SF) Alkaline phosphatase (S) N-propeptides (PIIANP and PIIBNP; S, SF, U) C-propeptide (PIICP, or CPII; S, SF, U)

Aggrecan

Chondroitin sulfate epitopes (846, 3-B-3, 7-D-4; S, SF)

Other non-collagen proteins

Glycoprotein 39 (YKL-40; S, SF) CD-RAP (SF)

Synovium Type III collagen Non-collagen proteins

Type III N propeptide (PIIINP) Hyaluronan (S) Glycoprotein 39 (YKL-40; S, SF) COMP (S, SF)

Degradation Pyridinoline (PYD; S, SF, U) Deoxypyridinoline (DPD) C-telopeptide (CTX-I; S, U) N-telopeptide (NTX-I; S, U) Bone sialoprotein (S) TRAP, isoenzyme 5b (plasma) PYD (S, SF, U) C-telopeptide (CTX-II; S, SF, U) Collagenase cleavage epitopes COL2-3/4long mono (C2C) COL2-3/4Cshort Core protein cleavage epitopes VDIPEN (MMP epitope) NITEGE (ADAMTS epitope) Keratan sulfate epitopes (5-D-4, ANP9) Cartilage oligomeric matrix protein (COMP)

Glc-Gal-PYD (U)

All of the markers listed can be found in serum (S) or synovial fluid (SF) of RA patients and some (PYD) may also be detected in the urine (U). The absolute levels of a given marker may not be indicative of disease progression or tissue damage without correlation with another molecular marker of tissue destruction or synthesis, as outlined in this chapter, or of inflammation or catabolism (see Table II). Synthetic markers may represent a repair response or degradation of newly synthesized molecules.

Figure 1

Radiographic and histopathologic features of bone erosions in RA. A plain radiograph of the hand from a patient with RA demonstrates extensive marginal and subchondral bone erosions as well as joint space narrowing. The circle highlights the region of focal marginal joint erosion. A low-power photomicrograph demonstrates the progressive destruction of the cartilage associated with the invading synovial pannus and the interface between the pannus and adjacent mineralized bone from a patient with RA. The histological section shows the presence of multinucleated osteoclast-like cells, characterized by TRAP-positive staining at the bone–pannus interface eroding the bone surface at the joint margin in a patient with RA.

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Steven R. Goldring and Mary B. Goldring 1. FOCAL SUBCHONDRAL AND MARGINAL BONE EROSIONS

The earliest skeletal lesions are localized to the sites where the inflamed synovial tissue comes into direct contact with the marginal bone surfaces. Focal resorption of the bone matrix leads to excavation of the bone surface, which produces the characteristic cystic bone erosions that can be detected radiographically. In addition, the synovial tissue may penetrate the subchondral bone plate, permitting invasion of the marrow spaces. The inflammatory tissue may then undermine the bone plate and produce additional focal areas of subchondral bone loss (see Fig. 1). Important insights into the pathogenesis of the progressive focal bone erosions and subchondral osteolysis that characterizes the synovial lesion of RA have been provided by the studies of Bromley and Woolley [33, 34]. They noted the presence of increased numbers of multinucleated cells with phenotypic features of osteoclasts localized to osteolytic lacunae at the junction of the subchondral bone and adjacent marrow space. Multinucleated cells, which expressed abundant tartrate-resistant acid phosphatase activity, were also observed adjacent to the calcified cartilage matrix and they identified these cells as “chondroclasts” [33]. These chondroclasts were observed in 30% of the specimens from large joints such as the knee where they appeared to make a major contribution to the subchondral bone erosions. Interestingly, they rarely observed these cells in specimens from small joints, and they speculated that the mechanisms associated with small joint pathology in RA might differ from that in the large joints. Allard et al. [35] have made similar speculations regarding the differential mechanism of focal bone loss in large and small joints. Characterization of cells isolated from RA synovial tissues has provided further support to the concept that cells with phenotypic features and functional activities of authentic osteoclasts are responsible for at least a component of the focal bone resorption that characterizes the RA synovial lesion [36, 37]. This does not preclude the possibility that other cell types present in the synovial lesions also have the capacity to resorb bone. Studies by several authors have demonstrated that macrophages and macrophage polykaryons, as well as synovial fibroblasts, have the ability to resorb bone, although their resorptive capacity is much less than that of osteoclasts [37–40]. Studies in vitro using explant cultures of synovial tissues from patients with RA also support the concept that mononuclear cells derived from synovial tissues from patients with RA can differentiate into cells expressing an osteoclastic phenotype, including the enhanced capacity to resorb bone. In initial studies by Fujikawa et al. [41], synovial fragments from patients with RA were digested

and the dispersed cells added to bone slices in the presence or absence of the rat osteoblast-like cell line UMR 106. Addition of 1,25(OH)2 vitamin D3 and human macrophage colony stimulating factor (M-CSF) resulted in differentiation of CD11b- and CD14-positive mononuclear into multinucleated cells expressing tartrate-resistant acid phosphatase activity, calcitonin receptor and vitronectin receptor that were capable of extensive bone resorption. Subsequently, several investigators have provided further support indicating that RA synovial tissues contain abundant osteoclast precursors [37, 42–45]. These results indicate that the RA synovium represents a site to which osteoclast precursor cells of monocyte-macrophage lineage are recruited in response to the local inflammatory mediators. In the presence of appropriate stimuli, these cells have the capacity to differentiate into authentic osteoclasts that in turn mediate the local bone-resorptive process. Characterization of the phenotype of the cells at the bone–pannus junction has been hampered by the lack of a suitable marker to definitively identify osteoclasts and to distinguish these cells from other cells of macrophage lineage or other bone cell types. Our laboratory has cloned the human calcitonin receptor and utilized reagents developed from this cDNA to prepare riboprobes to determine whether mono- and multinucleated cells in resorptive lacunae at the bone–pannus interface in samples from patients with RA express calcitonin receptor mRNA [46–48]. Expression of this gene coincides with the terminal differentiation of the osteoclast to a fully activated resorbing cell and distinguishes the osteoclast from its monocyte/macrophage precursor [49]. We also examined the cells at the bone–pannus interface for expression of other osteoclast-associated genes, including cathepsin K and tartrate-resistant acid phosphatase (TRAP), which are additional phenotypic markers of osteoclast-like cells. These studies confirm that the multinucleated (and some mononuclear cells) in resorption lacunae express the full repertoire of osteoclast-associated markers, including the expression of abundant calcitonin receptor mRNA, adding further support to the concept that they are functionally indistinguishable from authentic osteoclasts. These cells were also examined for the expression of mRNA encoding the parathyroid hormone receptor, and of interest, no message was detected. These findings are consistent with the observations of others who speculate that parathyroid hormone does not act directly on osteoclasts but rather mediates its effects on early osteoclast progenitors or indirectly via effects on osteoblast lineage and/or marrow stromal cells [50–52]. To more definitively establish an essential role for osteoclasts in the pathogenesis of focal erosions, our laboratory

Chapter 51 Rheumatoid Arthritis and other Inflammatory Joint Pathologies

and others have utilized experimental models of inflammatory arthritis employing transgenic and knockout mice lacking the capacity to produce osteoclast. In our own studies, we generated inflammatory arthritis in mice in which the gene for receptor activator of nuclear factor-B ligand (RANKL) was deleted [53]. This cytokine is a factor that is essential for osteoclast differentiation and activation, and mice lacking this gene are devoid of osteoclasts and exhibit severe osteopetrosis [54]. Inflammatory arthritis was induced in these mice and control littermates using a serum transfer model of inflammatory arthritis developed by Mathis, Benoist, and coworkers [53, 55–57]. This model of inflammatory arthritis exhibits many of the features of RA, including the formation of pannus and extensive bone and cartilage destruction. After serum transfer, wild-type and RANKL-deficient mice developed comparable levels of synovitis and joint inflammation. However, the RANKL knockout mice, which lacked the capacity to form osteoclasts, failed to develop significant bone erosions. Further evidence indicating an essential role for osteoclasts in the pathogenesis of focal bone erosions in inflammatory arthritis is proved by the studies of Redlich et al. [58]. They backcrossed mice lacking the c-fos gene with TNF-α transgenic mice. This transcription factor is essential for osteoclastogenesis and the c-fos null mice are, therefore, unable to generate osteoclasts [59, 60]. The TNF-α transgenic mice spontaneously develop a form of inflammatory arthritis resembling RA and the backcrossed mice provided a useful model for determining the requirement for osteoclasts in the development of bone erosions. Although the c-fos null mice backcrossed with the TNF-α transgenic mice developed inflammatory arthritis that was comparable to their wild-type littermates, the knockout animals failed to develop bone erosions. The observations from these studies and the results from the RANKL knock-out mice with serum transfer arthritis provide strong evidence indicating that osteoclasts are required for the development of focal bone erosions in inflammatory arthritis. Analysis of the cytokine profiles of RA synovial tissues has provided further insight into the mechanisms involved in the pathogenesis of the osteoclast-mediated bone erosions. RA synovium produces a broad spectrum of factors possessing the capacity to induce osteoclast differentiation and activation, as shown in Figure 2. These include IL-1, interleukin-6 (IL-6), interleukin-7 (IL-7), interleukin-11 (IL-11), interleukin-15 (IL-15), interleukin-17 (IL-17), monocyte/macrophage-colony stimulating factor (M-CSF), TNF-α, and parathyroid hormone-related peptide (PTHrP), the factor associated with humoral hypercalcemia of malignancy [61–67].

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Figure 2

Osteoclast-inducing and -activating factors produced by the RA synovium. IL-1, TNF-α, IL-15, IL-17, and PTHrP enhance osteoclast formation by stimulating bone-lining cells to produce osteoclast-inducing factors such as RANKL, M-CSF, and IL-6. OPG is a soluble decoy receptor that binds RANKL and inhibits its osteoclast-inducing activity. TNF-α also acts directly on osteoclast precursors to induce their differentiation into osteoclasts. IL-1 acts directly on osteoclasts to increase their resorbing activity.

Kong and coworkers [54] were the first to identify RANKL in RA synovial tissue. Subsequently, several laboratories have demonstrated RANKL mRNA and protein in RA synovial T cells and fibroblasts and in synovial tissues from a variety of animal models of inflammatory arthritis [43, 61, 68]. Of interest, this cytokine was originally identified as a T cell product that regulated dendritic cell function [69–72] and was designated as tumor necrosis factor-related activation-induced cytokine (TRANCE). Subsequently, its action as a critical regulator of osteoclastogenesis was established [73, 74]. RANKL transduces its effects via receptor activator of NF-κB (RANK) that is expressed on osteoclast precursors, osteoclasts, dendritic cells, and a variety of non-immune cells, including chondrocytes and osteoblasts. The activity of RANKL is regulated by an inhibitory molecule, osteoprotegerin (OPG), which is a soluble protein that acts as a decoy receptor for RANKL [75–77]. Kong et al. [54] provided the first evidence indicating a critical role for RANKL in the pathogenesis of focal bone erosions in inflammatory arthritis. They showed that treatment of rats with adjuvant arthritis with OPG (the decoy receptor for RANKL) markedly reduced the number and extent of focal articular bone erosions. Subsequently, several laboratories have demonstrated inhibition of bone erosions with OPG in different models of inflammatory arthritis, including the collagen-induced model of inflammatory arthritis [78] and in TNF-α transgenic animals

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with spontaneous arthritis [79]. Synovial fibroblasts and T-cells are the principal sources of RANKL in RA synovium, and these cell types have been shown to possess the capacity to support osteoclast formation [43, 54, 68, 80–82]. Weitzmann et al. [81] demonstrated that treatment of co-cultures of activated T cells and osteoclast precursors with OPG failed to completely block osteoclast formation, indicating that products in addition to RANKL are involved in the osteoclastogenic activity of these immune cells. As described above, T lymphocytes (as well as synovial fibroblasts) produce a variety of osteoclast-inducing factors in addition to RANKL. These factors likely work in concert with RANKL to contribute to the enhanced osteoclastmediated resorption at the site of bone erosions. 2. PERIARTICULAR BONE LOSS

Loss of bone adjacent to inflamed joints is the radiographic hallmark of inflammatory joint disorders such as RA. These radiographic changes are usually associated with sites of active synovitis. The bone loss typically occurs early in the course of RA and usually precedes the appearance of focal bone erosions. For example, Stewart et al. [83] utilized dual-energy X-ray absorptiometry to investigate the predictive value of juxta-articular osteopenia on subsequent development of marginal hand joint erosions detected by standard radiographic techniques. After 1 year, they found that reduced periarticular bone density was specific (100%) and highly sensitive (63%) in predicting individuals who would develop new marginal joint erosions or experience worsening of existing erosions. Shimizo et al. [84] performed histomorphometric analysis of bone samples obtained from patients undergoing joint arthroplasty to gain insights into the cellular mechanisms responsible for the periarticular bone loss in patients with RA. Their analyses revealed evidence of increased bone remodeling with an increase in both bone formation and resorption indices. Local production of proinflammatory mediators that affect bone remodeling, hypervascularity of the synovium, and joint immobilization have been implicated as possible mechanisms responsible for periarticular abnormalities in RA joints [8, 85–90]. 3. GENERALIZED BONE LOSS

Numerous studies have established that in addition to the local effects of the inflammatory process on juxtaarticular bone, patients with RA have a lower bone mineral density (BMD) and an increased risk of fracture compared to age-matched controls [91–99]. The decrease in bone mass affects the axial spine and appendicular bone distant from synovial-lined joints, suggesting that the local joint pathology is associated with a generalized systemic effect [90, 100–107]. Further evidence that RA

adversely affects skeletal remodeling is provided by the study of bone density in monozygotic twins (one with RA and one disease-free). These analyses reveal a significant reduction in BMD in the lumbar spine (4.6%), femoral neck (9.7%), and total body (5.7%) in the twin with RA [108]. More recently, Lodder et al. [99] examined the relationship between BMD, disease activity, and joint damage in 373 patients with low to moderately active RA. They demonstrated an association between the incidence of radiographic joint damage and the presence of low BMD at the hip and suggested that the increased disease activity was a significant contributory factor to the systemic bone loss. In these studies, the use of corticosteroids was not associated independently with BMD. Multiple confounding factors have hampered the investigation of the mechanisms of pathogenesis responsible for the generalized bone loss observed in patients with RA. These include the influence of patient sex, age, mobility, disease activity and duration, and the concomitant use of immunosuppressive therapies and/or glucocorticoids, all of which are known to have profound and independent effects on bone metabolism. Among these factors, disease activity and duration appear to be of particular importance with respect to the risk for reduced bone mass [109]. In a longitudinal prospective study, Gough and coworkers [92] concluded that significant amounts of generalized skeletal bone were lost early in RA and that this loss was associated with disease activity. These findings support the previous observations of Als et al. [110], who also noted a significant decrease in bone mass during the early phases of RA. Further evidence of the impact of disease activity on bone mass is provided by the studies of Sambrook et al. [101], who showed that two measures of disease activity, joint count and C-reactive protein (CRP), correlated with trabecular bone loss in the lumbosacral spine. In addition, epidemiologic studies have consistently shown the relationship between physical activity and functional level and bone density in patients with RA. For example, using the Framingham activity index (as a measure of functional status), Sambrook et al. observed that the level of physical activity correlated with femoral neck bone mineral density in a series of patients with RA [103]. These findings were corroborated by Gough et al. [92], who showed an association between disability level (based on the Health Assessment Questionnaire) and bone loss in the femoral neck and axial spine. Several different approaches have been utilized to gain insights into the pathogenesis of the generalized bone loss associated with RA (Table I). The most specific markers of bone formation are serum osteocalcin, skeletal alkaline phosphatase, and the aminoterminal propeptide of type I

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Chapter 51 Rheumatoid Arthritis and other Inflammatory Joint Pathologies

collagen (PINP) (see also Chapter 34). The most sensitive markers of bone resorption are collagen crosslinks including deoxypyridinoline (DPD) and the amino- and carboxyterminal crosslinked telopeptide of type I collagen (CTX-I, NTX-I) (see Chapter 35). Histomorphometric analyses of bone biopsies from patients with RA indicate that the reduced bone mass observed is related to a decrease in bone formation rather than to an increase in bone resorption [107, 111, 112]. However, contradictory findings have been obtained in studies using biochemical markers of bone turnover [113–116]. Gough and coworkers [117] evaluated 232 patients with RA of less than 2 years duration. They showed that the urinary markers of bone resorption, pyridinoline (PYD) and DPD crosslinks, correlated with the change in BMD at all sites during the 2-year study and that these markers of resorption were increased in subjects with RA compared to age-matched control subjects. Similar observations have been made by several other investigators [91, 114–116, 118–121]. Excretion of both markers was significantly increased in the patients with active disease that lost bone quickly. The levels of these urinary markers were highly correlated with CRP but not with serum levels of interleukin-6 (IL-6) or interleukin-1 (IL-1) (Table II). In the final regression analysis, serial CRP was more closely correlated with BMD change than any of the other markers. Bone formation indices, as assessed by measurement of serum alkaline phosphatase and procollagen I carboxyterminal propeptide (PICP)

levels were not significantly suppressed, suggesting that suppression of bone formation was marginal and not a dominant factor contributing to the accelerated bone loss in RA subjects. Of interest, on an individual basis, the trend was for patients presenting with active RA to have low bone formation indices and then for the levels to rise rapidly if the disease activity was suppressed. However, a cross-sectional study involving 318 patients with established RA, showing decreased bone formation assessed by serum osteocalcin and increased bone resorption by measurement of serum CTX-1 compared to healthy controls, suggested an imbalance in bone remodeling [122]. The serum CTX-I levels were 35% higher in patients with severe radiographic erosions compared to patients without joint damage, whereas the serum osteocalcin levels were similar [122]. Furthermore, a more recent prospective study of 110 patients participating in the COBRA clinical trial showed that high urinary CTX-I levels predict increased risk of radiological progression over 4 years, independent of rheumatoid factor (RF) or erythrocyte sedimentation rate (ESR) [25]. Indices of bone remodeling have also been used to assess the effects of treatments on generalized bone loss in patients with RA (Table I). For example, Hall et al. [113] examined the effects of hormone replacement therapy on markers of bone metabolism in 106 postmenopausal women with RA treated with or without glucocorticoids. They found that the serum osteocalcin levels tended to be

TABLE II. Markers of Inflammation and Joint Destruction Category Inflammation

Proteinases and inhibitors

Cytokines

Chemokines

Proinflammatory or catabolic C-Reactive protein (CRP) Eosinophil sedimentation rate (ESR) Serum amyloid A (SAA) Stromelysin-1 (MMP-3) Collagenase-1, 2, 3 (MMP-1, 8, 13) Gelatinases (MMP-2 and 9) Membrane-type MMP (MMP-14) Cathepsin B, D, L Plasminogen activator Cathepsin K Tumor necrosis factor (TNF)-α Interleukin (IL-1)α and β IL-6, oncostatin M (OSM) IL-17, IL-18 RANKL IL-8 MCP-1 RANTES SDF-1

Anti-inflammatory or anticatabolic

Tissue inhibitor of MMPs (TIMP-1) Plasminogen activator inhibitor (PAI-1)

Anti-IL-1α IL-1Ra IL-4, IL-10 IL-13 Osteoprotegerin (OPG)

The systemic markers of inflammation, such as CRP and ESR, are indicative of disease activity and disease progression. The other markers listed are generally present in synovial fluids of RA patients and some may be detected in serum as outlined in this chapter. They may be produced by synovial cells, chondrocytes, or bone cells and their activities have been defined by their influence on bone and cartilage metabolism.

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lower in the RA subjects with or without glucocorticoid treatment compared to control subjects. Urinary DPD and CTX-I were elevated in glucocorticoid-treated subjects with RA compared to the untreated RA patients and controls. After hormone replacement therapy, the urinary CTX-I excretion decreased significantly in both the steroid-treated and non-treated group and these changes correlated with 2-year changes in BMD. More recently, Saag et al. [123] examined the effects of alendronate therapy on the prevention and treatment of glucocorticoid-induced osteoporosis in a heterogeneous group of 477 patients, including a large number of individuals with RA. They demonstrated that the urinary excretion of NTX-I decreased by 60% and serum alkaline phosphatase concentrations decreased by 27% in alendronate-treated subjects. These findings correlated with an increase in bone mass and a decrease in fracture risk.

C. Seronegative Spondyloarthropathies The seronegative spondyloarthropathies include ankylosing spondylitis, reactive arthritis, Reiter’s syndrome, spondylitis, and arthritis associated with psoriasis or inflammatory bowel disease, and juvenile-onset spondyloarthropathy. These disorders encompass a broad range of multisystem inflammatory disorders in which the spine, peripheral joints, periarticular structures, or all three sites are involved. The joint inflammation that accompanies these disorders exhibits many of the same histopathological features that characterize the joint pathology in RA, including synovial hyperplasia, lymphoid infiltration, and pannus formation. Joint margin and subchondral erosions and cartilage destruction are often seen, although the inflammation may be restricted to only a few isolated joints. The pattern of distribution is typically asymmetrical, affecting proximal as well as distal joints such as the distal interphalangeal joints that are not typically affected in RA. Inflammation of the enthesis (sites of tendon or ligament attachment to bone), especially in the axial spine, is the pathological hallmark of the spondyloarthropathies, and this feature distinguishes this group of disorders from the other inflammatory arthritides. Examination of synovial tissues from patients with psoriatic arthritis reveals the presence of osteoclast-like cells at the synovial–bone interface. Interestingly, peripheral blood mononuclear cells from patients with active joint disease exhibit a significant increase in the CD11b+hi osteoclast precursor cell pool that can be induced to form osteoclasts in culture. When these patients are given anti-TNF-α therapy, clinical improvement is associated with a reduced number of these CD11b+hi osteoclast precursor cells in the peripheral blood [124].

McGonagle and co-workers [125–128] have utilized magnetic resonance imaging and anatomic and histopathological analysis to confirm that the enthesis is the initial site of inflammation in the spondyloarthropathies. At sites in which the entheseal inflammation is proximate to the joint capsule, the inflammatory process may then extend to the synovial lining of the joint cavity and to the adjacent tenosynovium. The earliest focal joint pathology in patients with spondyloarthropathies most often presents as a localized zone of bone resorption that initially affects the insertion sites of juxta-articular tendon or ligamentous insertions. These enthesopathic erosions are detectable radiographically as a localized zone of radiolucency. In contrast to the joint pathology in RA, in which the inflammatory process ultimately leads to progressive bone loss with the absence of evidence of bone repair in patients with spondyloarthropathies, the initial destructive bone lesion may lead to calcification and ossification of the enthesis in the later stages of the disease process. Braun et al. [129] have used immunohistological and in situ hybridization techniques to study the inflamed sacroiliac joints of patients with spondylitis to gain insights into the biochemical and cellular mechanisms involved in the tendency of the joint inflammation to lead to excessive bone formation. In these studies, computer-assisted tomography was used to obtain biopsies of the sacroiliac joints in five patients with ankylosing spondylitis. Analysis of synovial tissue revealed dense infiltrates of T lymphocytes, consisting of both CD4-positive and CD8-positive cells, and CD14-positive macrophages. Localized nodules containing active foci of endochondral ossification were noted within the pannus, a finding not observed in synovial tissues from patients with RA. In situ hybridization identified abundant message for TNF-α (but not IL1-β) within the inflammatory cells. Of interest, with respect to the tendency for the development of ossification at the sites of joint inflammation, the message for TGF-β2 was noted in close proximity to the regions of new bone formation. More recently, Lories et al. [130] examined synovial biopsies from individuals with RA, spondyloarthropathy and controls for the expression of bone morphogenic proteins (BMPs). They detected elevated levels of BMP-2 and BMP-6 in synovial tissues from the patients with spondyloarthropathy but not the controls without inflammatory arthritis. These growth factors were localized by immunostaining to synovial fibroblasts and macrophages. They speculated that the bone growth factors could contribute to the enhanced bone formation seen at sites of inflammation in individuals with spondyloarthropathies. The observation that patients with RA also had elevated levels of these bone growth factors was somewhat surprising in light of the absence of new bone formation at sites of synovial inflammation in RA. The differential pattern of

Chapter 51 Rheumatoid Arthritis and other Inflammatory Joint Pathologies

bone formation in RA and the spondyloarthropathies could in part be related to the anatomic site of the inflammatory process, which occurs when the spondyloarthropathies involve the enthesis and adjacent periosteum. These findings suggest the possibility that local production of growth factors by the inflammatory tissues may induce the new bone formation that gives rise to dactylitis and ossification and eventual bony ankylosis of the sacroiliac joints that occur in some individuals with spondyloarthropathies. A similar process could account for the development of syndesmophytes that characterize the spinal pathology. In the spine, the initial lesion consists of inflammatory granulation tissue at the junction of the annulus fibrosis of the intervertebral disk and the margin of the vertebral bone. Replacement of the outer annular fibers of the intervertebral disks with bone by mechanisms similar to those described by Braun and Lories [129, 130] could lead eventually to formation of syndesmophytes, with resultant ankylosis of the adjoining vertebral bodies. That the local joint and entheseal inflammation may be reflected in systemic markers of bone formation is suggested by the studies of Franck et al. [131]. They investigated the systemic rate of bone formation in a series of patients with psoriatic arthritis compared to controls and individuals with psoriasis without joint involvement. Serum osteocalcin and alkaline phosphatase levels were elevated in the patients with psoriatic arthritis compared to the controls and patients with psoriasis. There was also a significant correlation between clinical activity and increased bone formation in the patients with psoriatic arthritis. Although individuals with spondyloarthropathies exhibit a tendency towards the development of localized regions of new bone formation at sites of enthesopathic peripheral joint and spine inflammation, many patients exhibit evidence of marked osteopenia of the spinal column. The bone loss that occurs following bony ankylosis of the spine is a common feature that is often ascribed to spinal immobility [132]. Will et al. [133] evaluated 25 patients with early ankylosing spondylitis using dual-photon absorptiometry and compared them to age- and sex-matched controls. They observed a significant reduction in bone mineral density in the lumbosacral spine and hip in the spondylitis cohort. Of interest, these changes occurred even in the absence of bony ankylosis. Based on these observations, they concluded that bone loss occurred early in disease before the development of spinal immobility. They speculated that local adverse effects of inflammatory mediators released from the sites of joint pathology, rather than immobilization, were responsible for the progressive bone loss. Frediani et al. [134] examined the BMD in 100 control subjects and 186 patients with psoriatic arthritis who lacked axial skeletal involvement. Decreased BMD was detected in the spine and hip in two-thirds of the patients

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with psoriatic arthritis. The decreased BMD was not correlated with indices of inflammation or disease duration, although there was an association with the functional status as assessed by the Health Assessment Questionnaire (HAQ) score, as well as age and the number of years since menopause. These results add further support to the concept that decreased bone density in the spine in patients with spondyloarthropathies may occur even in the absence of spondylitis. Ralston et al. [135] studied the pathogenesis of spontaneous spinal insufficiency fractures in patients with ankylosing spondylitis. Of 111 patients with ankylosing spondylitis evaluated prospectively, 15 developed radiographic evidence of vertebral compression fractures. Interestingly, bone mineral density of the appendicular skeleton as assessed by single-photon absorptiometry was normal. These findings suggest that osteoporosis in these patients is primarily localized to the axial spine. These observations are supported by the findings of Devogelaer et al. [136] using a variety of techniques, including standard radiographs, single- and dual-energy absorptiometry, and quantitative computer tomography. Of interest, studies by Marzo-Ortega et al. [137] demonstrated that BMD improves in patients with spondyloarthropathies who show clinical responses to the TNF-α blocker, etanercept.

D. Juvenile Rheumatoid Arthritis (JRA) Numerous authors have noted the often-dramatic adverse effects of JRA and related forms of inflammatory arthritis on linear growth and acquisition of skeletal mass in children [138, 139]. Hillman et al. [140] analyzed biochemical markers of bone remodeling and total body calcium in 44 children with active polyarticular or oligoarticular JRA. Their findings indicated a low bone formation rate with an overall reduction in bone turnover. There was no evidence for an increase in bone resorption rates. Falcini et al. [141] reported similar results. Several investigators have reported reduced bone mass in children with JRA [142–144]. Hopp et al. [142] studied spinal bone density in 20 children with active JRA using dual-photon absorptiometry to assess bone mass. They noted that the bone density was reduced in postpubertal girls compared with healthy controls. These observations provide good evidence of the adverse effect of inflammation on skeletal accretion during puberty. Because of their normal rapid increase in skeletal mass, adolescents with active joint disease may be particularly vulnerable to the impact of the inflammatory process on their capacity to achieve expected peak bone mass. Surprisingly, when prepubertal girls with inflammatory joint disease were evaluated, they did not differ from controls at any of the sites measured. This could be related to effects of treatment

852 on the activity of the disease in this group. These results contrast to those of French et al. [145] who examined a population-based cohort of adults with a history of JRA. Although many of the subjects had T scores within the normal range, 41% were osteopenic (T score < − 1) at the lumbar spine or femoral neck, suggesting they would be at future risk for fracture. Reed and co-workers [146] have evaluated radial BMD and serum osteocalcin levels in children with JRA. Over a 3-year period, they noted that improvement in the disease activity was associated with an increase of serum osteocalcin to normal levels. In contrast, in children with persistent joint inflammation, serum osteocalcin and BMD remained below normal. These findings provide further evidence of an adverse systemic effect of the underlying inflammatory joint disease on skeletal remodeling. The changes in serum osteocalcin levels are consistent with a suppression of bone formation, and the findings suggest that the effects of the suppression of bone formation in children, especially during the pubertal growth period, may have an adverse effect on the achievement of normal peak bone mass. That this decrease in bone mass predisposes to an increased risk of fracture in adulthood is supported by several studies [147, 148]. Several proinflammatory cytokines and their receptors have been implicated in the disturbance of local as well as generalized bone remodeling in children with JRA. Serum levels of the soluble receptor for TNF-α, the soluble form of the IL-2 receptor, and IL-6 have been shown to correlate with markers of inflammation (ESR, CRP) indicating an association of these markers with disease activity. The highest levels of these mediators were observed in patients with active systemic disease or with polyarthritis [149]. Similar observations have also been made in adults with RA and related forms of inflammatory arthritis.

IV. EFFECTS OF JOINT INFLAMMATION ON CARTILAGE REMODELING The subchondral bone is covered by articular cartilage that consists of chondrocytes, which are somewhat unique to this tissue, embedded in a specialized extracellular matrix. The cartilage matrix consists of highly crosslinked fibrils of type II collagen molecules associated with several cartilage-specific collagens, including types IX and XI, the large aggregating proteoglycan aggrecan, small proteoglycans, including decorin, biglycan, fibromodulin, and lumican, and other collagens (VI, XII, and XIV) and noncollagenous matrix proteins [1, 150, 151] (see Part I and Chapter 25). Within the tissue there are no blood vessels, nerves, or lymphatics and nutrition for the chondrocytes

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is derived by diffusion from both the blood supply in the subchondral bone and from the synovial fluid. Normally, chondrocyte proliferation is limited and penetration of other cell types from the joint space or subchondral bone is restricted. Under physiologic conditions, the chondrocytes maintain a stable equilibrium between the synthesis and the degradation of matrix components with a half-life of over 100 years for type II collagen [152] and a half-life for aggrecan subfractions in the range of 3–24 years [153]. During aging and in inflammatory and degenerative joint diseases, such as RA and osteoarthritis (OA), this stable equilibrium is disrupted and the rate of loss of collagens and proteoglycans from the matrix exceeds the rate of deposition of newly synthesized molecules [151]. This state of disequilibrium leads to the progressive loss of articular cartilage and disruption of normal joint function.

A. Cartilage Loss in RA Cartilage destruction in RA occurs primarily in areas contiguous with the proliferating synovial pannus [154, 155], as shown in Figure 2. In early RA, the loss of proteoglycan occurs throughout the cartilage matrix and is not limited to regions covered by synovial pannus, and selective damage to type II collagen fibrils can be observed in middle and deep zones [156, 157]. In the cartilage–pannus junction, there is evidence of attachment of the synovial cells, including synovial fibroblasts and macrophages, to the cartilage surface. Both synovial cell types have been implicated in the degradation of the adjacent cartilage matrix via release of proteinases capable of digesting the cartilage matrix components [158]. Several authors have suggested a critical role for a distinctive fibroblast-like cell type, the so-called “pannocyte”, that is present within the inflamed RA synovium [3, 4, 159, 160]. These cells exhibit anchorage-independent growth and, in contrast to synovial fibroblasts from normal or OA synovial tissue, demonstrate the capacity to invade cartilage in the absence of an inflammatory environment [161]. In addition, there is evidence of loss of proteoglycan throughout the cartilage matrix, particularly in superficial zones that are in contact with the synovial fluid at sites that are not directly associated with the pannus–cartilage junction [162, 163]. These effects may be related to the release and activation of proteinases from polymorphonuclear leukocytes and other inflammatory cells present within the synovial fluid. The involvement of metalloproteinases (MMPs) in the degradation of cartilage collagens and proteoglycans in RA is well established (Table II) (see Chapter 11). The MMPs of the collagenase and stromelysin families have been given greatest attention because they specifically

Chapter 51 Rheumatoid Arthritis and other Inflammatory Joint Pathologies

degrade native collagens and proteoglycans, respectively, and active stromelysin also serves as an activator of latent collagenases. MMPs are localized in regions of cartilage degradation [164] and are elevated in synovial fluids [165–168] and sera [168–171] of RA patients. Of the collagen-degrading enzymes, collagenases-1, -2, and -3 (MMP-1, MMP-8, MMP-13, respectively) and membrane type I MMP (MT1)–MMP (MMP-14) are believed to play major roles in cartilage destruction in RA joints [172–174]. The levels of TIMP-1 are also increased in RA synovial fluids and sera [166, 167, 171], possibly reflecting an endogenous corrective response to the increased levels of active MMPs. Synovial fluid levels of collagenase correlate with the degree of synovial inflammation, but not with CRP [175]. In addition to MMP-1, -8 and -13, the gelatinases (MMP-2 and -9), MMP-14, and stromelysin (MMP-3) are elevated in the sera of RA patients, correlating with CRP and ESR, and are generally indicative of disease activity, but not necessarily of progression [176–178]. Synovial fluid levels of MMPs are more reliable indicators than the levels in sera [179, 180]. Of the MMPs detected in the serum, MMP-3 is the most reliable predictive of joint progression in early RA patients [178, 181, 182]. However, it is unclear whether serum MMP levels also reflect tissue destruction within synovial joints. Most of the MMP activity in the synovial fluid originates from the synovium in RA, although there is evidence of intrinsic chondrolytic activity by chondrocytes at the cartilage–pannus junction as well as in deeper zones of cartilage matrix in a minority of RA specimens [183]. Serum levels of both MMP-1 and MMP-3 are decreased by therapy with the anti-TNF-α antibody cA2, and MMP-3 levels correlate with CRP levels in RA patients [184, 185]. Although the presence of MMPs in synovial fluid or serum may not be a specific marker for cartilage degradation per se, antibodies that recognize specific cleavage epitopes in cartilage matrix molecules have been developed, as described below (Table II). Several of the MMPs, including MMP-3, MMP-8, and MMP-14 also have the capacity to degrade aggrecan. However, there is evidence that members of the reprolysinrelated proteinases of the ADAM (a disintegrin and metalloproteinase) family are the principal mediators of aggrecan degradation [186]. Aggrecanases-1 and -2 have been cloned and are characterized as ADAMTS (ADAM with thrombospondin-1 domains) -4 and -5 [187–189] and ADAMTS-1, identified originally as an inflammationassociated protein [190], is also expressed in cartilage [191, 192]. The presence in RA joint tissues of other proteinases that can degrade cartilage matrix components provides further complexity to the clinical picture [3, 193, 194]. Serine proteinase activities due to urokinase-type plasminogen

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activator and elastase, which also interact with the MMP proteolytic cascade, are elevated in RA synovial fluids [195, 196]. The cathepsins B, L, and D can be detected in joint tissues and synovial fluids in experimental and human RA [197–200]. Cathepsin K expression has been detected in synovial fibroblasts on the cartilage surface at the pannus–cartilage junction [201, 202]. Using in vitro cell culture techniques we have demonstrated that mature cathepsin K is secreted by primary cultures of synovial fibroblasts derived from RA synovium and that treatment of these cells with IL-1 or TNF-α markedly up-regulates cathepsin K production [202]. Among the known cathepsins, cathepsin K is the only proteinase that is capable of hydrolyzing types I and II collagens at multiple sites within the triple-helical regions, and its requirement for acidic pH may be provided by the microenvironment between the synovial pannus and the cartilage [203]. The increased elastase activity is found in synovial fluids from patients with knee joint synovitis and with RA may be associated with cartilage damage [204]. Other MMP and ADAMTS family members expressed in OA cartilage, including MMP-16, MMP-28, and ADAMTS2, 14, and 16 [205], may also play a role in cartilage damage in RA.

B. Systemic Markers of Cartilage Metabolism in Rheumatoid Arthritis Molecular markers in body fluids have been developed for identifying local and systemic metabolic changes, monitoring the effects of cytokines and therapeutic agents on cartilage metabolism, and predicting the rate of joint destruction in RA [20, 21, 206] (see Table I). Proteolytic degradation products of matrix components have been considered both as diagnostic markers of cartilage damage and as potential autoantigens in the induction and maintenance of RA and related inflammatory joint diseases [207–210]. Molecules originating from the articular cartilage, including aggrecan fragments, which contain chondroitin sulfate and keratan sulfate, type II collagen fragments, collagen pyridinoline crosslinks, and cartilage oligomeric protein (COMP), are usually released as degradation products as a result of catabolic processes (see Chapters 26 and 27). Degradation products of type II collagen [211, 212], as well as aggrecan [213, 214], can be detected in the synovial fluid of RA patients. Cartilage proteoglycan components, including the keratin-sulfaterich domain and G1 and G2 globular domains of aggrecan and link protein, are higher in synovial fluids than in sera of patients with inflammatory joint diseases such as RA, psoriasis, ankylosing spondylitis, and JRA and after injury [23, 24, 213–219]. In these disorders, cellular immunity

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to these matrix components may also be detected [220, 221]. However, since both cartilage catabolism and the matrix synthetic activity may be affected by inflammatory processes in the joint, the levels of cartilage matrix fragments in synovial fluid, serum, and urine may represent the net result of these two processes. To overcome potential problems in interpretation of such analyses, specific antibodies that detect either synthetic or cleavage epitopes have been developed to study biological markers of cartilage metabolism in body fluids. 1. TYPE II COLLAGEN

Antibodies that recognize different epitopes on type II collagen have been used to follow both synthesis and degradation of cartilage collagen (Table I). The carboxyterminal collagen II-propeptide (PIICP or CPII) is released by the activity of C-propeptidase during extracellular fibril formation, and after a short half-life in cartilage (~12 h), it releases into the circulation. PIICP concentrations in body fluids thus reflect the rate of synthesis of cartilage collagen. RA patients have abnormally elevated serum levels and synovial fluid levels are greater than those in serum [23]. In established RA, PIICP levels are inversely related to keratan sulfate levels, a marker of aggrecan turnover. The N-terminal propeptide (PIINP) exists in two forms, PIIANP and PIIBNP, which reflect the production of type IIA collagen expressed primarily during skeletal development and type IIB collagen, the major form in adult cartilage. ELISA analysis detected decreased serum levels of PIIANP in patients with established RA [222]. The first evidence of increased degradation of type II collagen in RA articular cartilage in situ was provided by Dodge and Poole [163]. More recently, specific antibodies that detect cleavage of the triple helix of type II collagen have been developed [223, 224]. Immunoassay of the collagenase-generated cleavage neoepitope, using the C2C antibody (previously known as Col2-3/4CLong mono), detects increased type II collagen degradation in experimental models of RA [27, 225, 226]. The levels of type II collagen neoepitope detected by C2C in RA synovial fluids are variable, but correlate with the measures of local and systemic inflammation and with levels of MMP-1 and TNF-α, but not MMP-3 [28, 227]. In a recent study, an ELISA using the C2C antibody detected increased levels of neoepitope in both urine and sera from RA patients with moderate disease activity [228]. In contrast, in ankylosing spondylitis patients, the serum levels of C2C are not significantly different from controls, whereas the CPII (PIINP) levels are significantly elevated and the CPII:C2C ratio correlates with CRP in patients with active disease and responsiveness to anti-TNF-α therapy [229].

Collagen degradation has also been studied by monitoring the urinary excretion of pyridinoline crosslinks. The hydroxylysyl pyridinoline, referred to as PYD, is the most common crosslink of cartilage, but it is also present in bone, which also contains lysyl pyridinoline (DPD) crosslinks. There are reports that the levels of PYD, but not DPD, are elevated in RA urine and serum, and that they are greatest in severe or late RA than in inactive disease or OA [114, 118, 119, 230–232]. In one study, high levels of PYD were found in synovial fluids from RA patients, but DPD was not detected [121]. Although it is unclear whether the PYD:DPD ratio can be used as a marker for which tissue is under attack [233], it is of potential use for identifying the severity of destruction of cartilage and/or adjacent bone and the response to therapy [234]. For example, a report that tranexamic acid, an inhibitor of plasminogen activation, reduces collagen crosslink formation in RA indicates that this measure is clinically relevant [235]. However, the most clinically relevant data have been generated using urinary CTX-II, a fragment of the C-telopeptide of type II collagen [26]. In a study of early RA patients that had not undergone methotrexate treatment, high baseline levels of urinary CTX-II, along with serum CTX-I, a marker of bone degradation, and urinary Glc-Gal-PYD, a marker of synovial inflammation, were associated with a high risk of progression [21, 115]. The high urinary CTX-II in early RA patients followed prospectively in the COBRA clinical trial, along with the high urinary CTX-I levels mentioned earlier, predicted increased risk of radiological progression over 4 years, independent of RF or ESR [25]. CTX-II has also been used effectively to predict the effect of disease-modifying treatment on long-term radiographic progression in patients with RA [236]. 2. AGGRECAN

Fragments of the large aggregating proteoglycan, aggrecan, and several of its epitopes can now be measured in synovial fluid and serum (Table I). The highest levels of the chondroitin sulfate region of the core protein are observed in the synovial fluid of patients with reactive arthritis [237]. Another aggrecan epitope, the hyaluronic acid-binding region (HABr) or G1 domain, is released in small amounts in this condition. In RA an inverse relationship exists between these two epitopes with dominance of aggrecan release in early, and the prevalence of HABr in late, destructive disease [238]. Monoclonal antibodies that have been developed to recognize subtle differences in the glycosaminoglycans chondroitin sulfate (CS) and keratan sulfate (KS) have been used to investigate the sera and synovial fluids of RA patients. The CS epitope in the G3 domain of aggrecan recognized by monoclonal

Chapter 51 Rheumatoid Arthritis and other Inflammatory Joint Pathologies

antibody 846 is almost absent in normal cartilage but it is a useful serum marker, since it is released in pathologic conditions [227, 239]. Fragments of aggrecan bearing the 846 epitope reflect the turnover of newly synthesized aggrecan molecules and serum levels are frequently elevated in patients with chronic RA [24]. However, very low levels are found in RA patients with rapid erosive disease where the 846 epitope appears to represent increased aggrecan synthesis, thereby reflecting an attempt at repair and a possibly favorable prognosis [23, 28, 227]. An inverse correlation between the 846 epitope and indices of inflammation, CRP and ESR, suggests an indirect relationship between cartilage repair and inflammation [23]. Similarly, the monoclonal antibodies, 3B3(−) and 7D4, detect CS neoepitopes in synovial fluid that correlate inversely with the extent of articular cartilage damage in RA, OA, and acute knee injury [240–242]. In contrast, patients with ankylosing spondylitis also have increased serum levels of the 846 epitope, which tend to correlate with TGF-β levels and may be related to the increased biosynthetic turnover in articular and intervertebral disk cartilage [229]. The circulating levels of KS are generally decreased in RA patients, reflecting turnover of newly synthesized aggrecan [24, 218]. Although serum KS levels correlate with the severity of articular damage in RA measured radiographically [215, 243], synovial fluid levels correlate inversely and may reflect a decrease in cartilage mass in chronic RA [241]. KS, measured by monoclonal antibody 5D4 and ratios of chondroitin-6- to chondroitin4-sulfate [244], are lower in synovial fluids from RA patients than in those from healthy subjects or OA patients [241, 245]. The levels of the KS epitope in sera and synovial fluids are related inversely to TNF-α, serum acute-phase protein, MMP-3, and PMN number in synovial fluids or sera in RA patients and with CD68+ macrophage infiltration in the synovium [24, 28, 218, 246]. Recent attention has focused on the enzymes responsible for aggrecan degradation [247]. Patients with early-stage RA who exhibit high concentrations of aggrecan fragments in synovial fluid progress more rapidly towards joint destruction than patients with low concentrations [207]. At least seven specific cleavage sites have been defined in the protein core of aggrecan. Highly specific antipeptide antibodies that recognize catabolic sites within the interglobular G1 domain of aggrecan have been developed as sensitive markers for following disease course and activity and response to therapy (Table I). Antibodies have been developed to recognize the aggrecan G1 neoepitopes resulting from cleavage by MMPs between amino acid residues Asn341 and Phe342 (VIDIPEN) [248, 249] and cleavage by aggrecanases between Glu373 and Ala 374 (N-terminal ARGSV and C-terminal NITEGE [250, 251].

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Both MMP-8 [252] and membrane type I MMP (MT1MMP) [253] are capable of cleaving aggrecan at both sites. Antibodies that detect N- or C-terminal epitopes on either MMP- or aggrecanase-generated G1 fragments have been used to show elevated levels of fragments resulting from both activities in RA cartilage [254] and synovial fluids [255]. These studies confirm earlier work showing elevated levels of aggrecanase-generated fragments in RA synovial fluids [256]. The combined evidence indicates that both MMPs and aggrecanase activities may be important in aggrecan degradation, since fragments generated by both types of activities are found in RA synovial fluids, and that fragments produced by one type of activity are resistant to further cleavage by the other activity [255]. 3. OTHER CARTILAGE-DERIVED PROTEINS

Several novel cartilage matrix proteins discovered in recent years are not unique to cartilage, but have been investigated as markers of tissue damage in RA (Table I). Cartilage oligomeric matrix protein (COMP) is a pentameric extracellular matrix protein with subunit size of 100–110 kDa. It is primarily localized in the chondrocyte territorial matrix, as well as in synovium, tendon, and ligament and is synthesized by synovial fibroblasts and articular chondrocytes [257–259]. Studies in vitro showed that TGF-β1 increased COMP mRNA synthesis, and IL-1β counteracted this increase in human synovial fibroblasts and articular chondrocytes [257]. Adhesion of synovial fibroblasts to COMP via αvβ3 integrin induces synthesis of intracellular galectin-3, which is expressed at sites of joint erosion in RA synovium [260]. Like aggrecan, COMP is released in the synovial fluid in reactive arthritis and RA but it is more concentrated in the serum, thus giving a synovial fluid:serum ratio of 10:1. High serum levels in early RA may distinguish an aggressive destructive form of the disease [23, 261]. Several studies have shown that COMP is a measure of processes in cartilage that are distinct from inflammatory aspects of the disease [262–264], particularly cartilage turnover [265, 266]. A recent prospective study in a cohort of RA patients examined after 5 and 10 years has shown that serum COMP has additive predictive value in relation to inflammatory indices and autoantibodies, including IgA RF, ESR, CRP, antibodies against cyclic citrullinated peptide (CCP), and naturally occurring anti-IL-1α [29]. COMP may be particularly useful for following the effects of therapy on cartilage turnover in RA patients [267, 268]. YKL-40, a chitinase 3-like protein that is also termed human cartilage glycoprotein-39 (HC-gp39), is a major secretory product of articular chondrocytes, as well as synovial cells [269–271]. Similar to COMP, the synovial fluid:serum ratio is 10 and elevated levels are observed in

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patients with active inflammatory joint disease [270, 272–276]. The increased levels of YKL-40 in the synovial fluid and serum from patients with active RA correlate with CRP and with serum PIIINP (procollagen type III N propeptide), a marker of synovial inflammation, and the highest correlation is with the Larsen wrist index of bone erosion [270, 277]. Increased levels of YKL-40 (two-fold compared to controls) are found in patients with early RA, but progression occurs only in patients with sustained, elevated levels [278]. Thus, YKL-40 has not been found to be predictive of radiographic damage in prospective large cohort studies [11]. However, serum YKL-40 levels are responsive to treatment with prednisolone and/or antiTNF-α, although DMARDs are less effective, suggesting that the effective therapies may have direct effects on cellular expression of YKL-40 [274, 277, 279]. Since synovial cells, chondrocytes, and neutrophils from the joints, as well as other tissues, can contribute to the pool of YKL-40 released into the circulation, it cannot be considered as a specific marker of joint tissue turnover. Recent studies indicate that YKL-40/gp-39 in immune complexes with HLA-DR4 is a specific histological marker in inflamed RA synovium [280], but that the immune response to this antigen in healthy individuals is biased towards regulatory, suppressor T-cell phenotype that is shifted from an anti-inflammatory to a proinflammatory phenotype in RA patients [281]. Its primary role in cartilage may be to stimulate chondrocyte proliferation and matrix synthesis [282–284]. Recent work also indicates that a related member of the chitinase family, YKL-39, may be a more specific serum marker as cartilage-derived autoantigen [285, 286]. Another novel marker is the cartilage-derived retinoic-acid-sensitive protein (CD-RAP), also known as melanoma inhibitory activity, which is found at high levels in synovial fluids from patients with mild RA and decreases with disease progression [287].

C. Local Markers of Cartilage Metabolism in Rheumatoid Arthritis In addition to the direct action of proteinases released from the pannus, the RA synovial tissues contribute to cartilage matrix degradation by releasing cytokines and other mediators that act on chondrocytes to regulate catabolic and synthetic functions (see Table II). The resultant disequilibrium in remodeling likely contributes to the rapid loss of cartilage matrix components characteristic of the RA joint lesion. There is also evidence that the chondrocytes may participate in this destructive process not only by responding to the proinflammatory cytokines released from the synovium, but may themselves be the source of proinflammatory and catabolic cytokines that,

via autocrine or paracrine mechanisms, contribute to cartilage matrix loss [288–291]. Our understanding of basic cellular mechanisms regulating chondrocyte responses to inflammatory mediators has been inferred from numerous studies in vitro using cultures of cartilage fragments or isolated chondrocytes. Much of this information is also relevant to OA and is supported by studies in animal models [18, 292, 293]. The impacts of these cytokines on chondrocyte function, particularly with respect to the various roles in OA pathogenesis, have been reviewed recently [290, 294]. Less information has been derived from direct analysis of cartilage or chondrocytes obtained from RA patients where cartilage damage is extensive. 1. CYTOKINES AND CYTOKINE-INDUCED MARKERS

The alterations in products of cartilage matrix turnover and levels of matrix degrading proteinases and inhibitors described above are accompanied by changes in the levels of various cytokines in the rheumatoid synovial fluids (see Table II). The prominent role of proinflammatory cytokines in destruction of cartilage matrix in RA is well supported by evidence accumulated during the last 20 years [18, 295–298]. Of the cytokines released in the RA synovial joint, IL-1 and TNF-α are considered to be of major importance in cartilage destruction. In addition to inducing the synthesis of MMPs and other proteinases by chondrocytes and synovial cells, IL-1 and TNF-α increase the synthesis of prostaglandin E2 (PGE2) by increasing the expression or activities of cyclooxygenase (COX)-2, microsomal PGE synthase-1 (mPGES-1), and soluble phospholipase A2 (sPLA2). They also up-regulate the production of nitric oxide via inducible nitric oxide synthetase (iNOS), other proinflammatory cytokines such as IL-6, leukemia inhibitory factor (LIF), IL-17, and IL-18, and chemokines, including IL-8. IL-17, a recently discovered T-lymphocyte product, also stimulates the production of proinflammatory cytokines and has effects on chondrocytes similar to IL-1 [299]. The concentrations of IL-1 in plasma and synovial fluids of patients with RA are significantly higher than in healthy individuals and correlate with disease activity and radiographic and histological features [300–302]. Several other proinflammatory cytokines, including TNF-α, IL-6, oncostatin M (OSM), and IL-17, and chemokines such as IL-8 and monocyte chemoattractant protein-1 (MCP-1) are present in RA synovial fluids and sera at higher levels than in patients with other arthritides [303–311]. Recent work has focused on correlating cytokine levels in inflammatory body fluids with other markers of cartilage matrix turnover. For example, the levels of TNF-α and OSM, both of which can synergize with IL-1, correlate with markers of collagen and aggrecan degradation in RA synovial fluids, as well as with IL-6 levels [246].

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Transition from early to established inflammatory arthritis is associated with rising levels of TNF-α in the synovial fluid and a marked reduction of immunosuppressive IL-10, associated with the impaired T-helper (Th) cytokine response [28]. In contrast to the levels of cytokines themselves, the levels of autoantibodies or receptor antagonists may be useful prognostic indicators, particularly in combination with markers of inflammation and cartilage turnover (see Table II). For example, naturally occurring anti-IL-1α antibodies, which bind with high affinity to IL-1α and neutralize its activity, are detected at higher levels in sera of patients with less-destructive disease [29, 312, 313] and correlate inversely with COMP, an indicator of more severe joint damage, in one study [29]. Although the levels of IL-1 receptor antagonist (IL-1Ra) in RA synovial fluids are elevated compared with those in OA, the ratio of IL-1Ra to IL-1 is decreased in RA patients, suggesting that the levels may not be sufficient to prevent IL-1 receptor binding [314]. The synovial fluid levels of other inhibitory cytokines such as IL-4 and IL-10, which increase IL-1Ra production, decrease the production and actions of proinflammatory cytokines, and are chondroprotective in vivo [315, 316], are probably reflective of the synovitis and the imbalance in the Th1 and Th2 cytokine environment [2, 317]. The different profiles of cytokine production by mononuclear cells in the peripheral blood and synovial fluid also serve to distinguish patients with acute and chronic arthritis and determine suitability for anticytokine therapy [318]. Anti-IL-6 receptor therapy has been proposed for juvenile idiopathic arthritis, since these patients have high circulating levels of IL-6 and anti-TNF-α therapy has proven ineffective [30]. Adipocytokines, which can affect cartilage metabolism [319], have received some attention in recent years. Leptin, for example, correlates negatively with inflammatory markers in RA sera [320] and has been proposed to serve as a link between the neuroendocrine and immune systems [321]. 2. CHEMOKINES

The role of chemokines in RA tissues, where they play roles in neutrophil activation and chemotaxis, is well established, but their potential contribution to cartilage metabolism has been recognized only recently [294, 322] (Table II). IL-8, probably the most potent and abundant chemotactic agent in RA synovial fluids, and other chemokines, such as MCP-1 and RANTES, are produced primarily by the synovium and serve as indicators of synovitis [323]. Chondrocytes, when activated by IL-1 and TNF-α, express several chemokines, including IL-8, MCP-1, MIP-1α, MIP-1β, RANTES, and GROα, which are present at high levels in arthritic joints, and also have receptors that enable responses to some of these chemokines [324–326].

Although high levels of stromal cell-derived factor 1 (SDF-1) are detected in RA synovial fluids, its receptor, CXCR4, is expressed by chondrocytes but not synovial fibroblasts [327], suggesting that this chemokine could reflect cartilage damage. 3. ADHESION MOLECULES

In addition to the requirement of chemokines for the recruitment of T-lymphocytes and other inflammatory cells to the subsynovial lining, adhesion receptors must be available on synovial blood vessels for binding the circulating leukocytes and other cell types with which they interact in the inflamed tissue, including macrophages, dendritic cells, and fibroblasts [323, 328]. Principal families of adhesion molecules involved are the selectins, the integrins, the immunoglobulin supergene family, and variants of the CD44 family. Although many of these molecules are common to different inflammatory sites, as is their expression on circulating cells in different rheumatic conditions, tissue-specific differences in expression of certain molecules have been found. Many of the prominent adhesion proteins expressed in the inflamed rheumatoid synovium are also expressed in cartilage. For example, vascular cell adhesion molecule (VCAM)-1 and intercellular adhesion molecule (ICAM)-1, which are members of the immunoglobulin family, are expressed by human articular chondrocytes, as well as synovial and endothelial cells, and their expression is upregulated by cytokines. The serum levels of soluble forms of these adhesion molecules, sICAM-1 and sVCAM-1, as well as E-selectin, are increased in RA patients compared with OA patients [329] and sICAM-1, in particular, serves as a prognostic marker of progression for following anti-TNF-α therapy [279]. VCAM-1, as well as VEGF, FGF, and TNF-α, contribute to angiogenesis during synovitis and to activation of chondrocytes during cartilage degradation [330]. Serum levels of VEGF, FGF, and other angiogenic factors are elevated in RA patients, although the elevated synovial fluid levels cannot be used to distinguish RA from other forms of inflammatory arthritis [329, 331, 332]. Serum VEGF, which is upregulated by inflammatory cytokines and hypoxia, correlates with disease activity and radiographic progression [333]. Specific roles for the hyaluronan receptor, CD44 [334] and anchorin CII (annexin V or annexin 5A) in cell–matrix interactions in cartilage, have also been determined. Several members of the integrin family are expressed by chondrocytes. For example, the α1β1 integrin functions as a receptor for types II and VI collagen, and the α5β1 integrin serves as a chondrocyte fibronectin receptor [335]. However, these integrins and the other adhesion molecules described above are not unique to chondrocytes, and therefore, their use as specific markers of cartilage metabolism is limited. CD44 expression is upregulated on chondrocytes

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in articular cartilage from RA patients and is higher on lymphocytes from RA synovial fluid than on peripheral blood leukocytes [336–339]. Furthermore, induction of MMP-specific cleavage of type II collagen and nitric oxide production by the heparin-binding fragment of fibronectin is mediated by CD44 [340, 341]. Anti-CD44 therapy has been shown to reduce inflammation on a murine model of RA [342] and to inhibit cartilage destruction in an in vitro model [343]. Interestingly, β2 integrins, but not β1 integrins, are elevated in peripheral blood monocytes in RA patients and the levels decrease with prednisolone therapy [344]. 4. INTRACELLULAR MEDIATORS

Signal transduction molecules and transcription factors activated by inflammatory mediators in chondrocytes and synovial cells are not likely to serve as accessible diagnostic or prognostic biomarkers, but they may serve as therapeutic targets. For example, NF-κB is a “master switch” of the inflammatory cascade, through which many beneficial therapeutic agents against RA may act [345], and the signaling intermediates in the p38 and JNK pathways have also been targeted for future therapeutic development [346]. In addition to NF-κB, members of the C/EBP, STAT, ETS, and AP-1 families are important for expression of IL-1- and TNF-α-inducible genes, such as MMPs, COX-2, and iNOS [347–351], and have been localized in rheumatoid tissues [352–355]. IL-1-induced transcription factors also suppress the expression of a number of genes associated with the differentiated chondrocyte phenotype, including type II collagen (COL2A1), aggrecan, and CD-RAP [356, 357]. Transcription factors, such as Sox9, might serve as more specific markers of chondrocyte phenotype, since they are important regulators of chondrogenesis during development [358]; however, they have not been studied in the context of the rheumatoid joint. Other markers found in RA synovial fluids remain to be assessed in relation to cartilage damage, including the advanced glycation end products [359, 360], heat shock proteins [361], the small calcium-binding protein S100A8/A9 [362], and other proteins elucidated by proteomic analysis [363]. Antibodies against heat shock proteins in sera or synovial fluids may also correlate directly or inversely with joint damage [11, 364].

V. CONCLUSION The characterization of the molecular markers in synovial fluids and sera, and in some cases the urine, from RA patients and their association with bone and cartilage loss has led to the development of new strategies for

monitoring disease activity and disease progression. Recent work has defined combinations of molecular markers of turnover of bone, cartilage, and synovium that complement measures of inflammatory disease activity, such as CRP and ESR, and radiographic progression. Local markers of tissue inflammation and catabolism, such as proteinases, cytokines, and chemokines are indicators of immunomodulation and synovial cell function, but they may also reflect bone and cartilage damage. Their validation as tissue-specific diagnostic tools or therapeutic indicators has included analysis of expression and activities in cartilage and bone in situ, although few have proven useful for routine clinical evaluations. Nevertheless, laboratory investigations have led to the development of novel therapies, including cytokine antagonists, cytokine receptor-blocking antibodies, proteinase inhibitors, and inhibitors of signal transduction [295, 298, 365–374]. Recent work indicates that combinations of systemic and local markers have the potential of identifying early RA patients at risk of rapid joint destruction before joint damage is detected by radiography. The rapid responsiveness of biological markers is likely to prove valuable for monitoring therapy against disease progression.

References 1. Goldring, S. R., and Goldring, M. B. (2004). Biology of the normal joint. In “Kelley’s Textbook of Rheumatology” (E. D. Harris, S. Ruddy, C. B. Sledge, J. S. Sergent, and R. C. Budd, eds.), pp. 1–34. WB Saunders, Philadelphia, PA. 2. Tak, P. P., and Bresnihan, B. (2000). The pathogenesis and prevention of joint damage in rheumatoid arthritis: advances from synovial biopsy and tissue analysis. Arthritis Rheum. 43, 2619–2633. 3. Yamanishi, Y., and Firestein, G. S. (2001). Pathogenesis of rheumatoid arthritis: the role of synoviocytes. Rheum. Dis. Clin. North Am. 27, 355–371. 4. Firestein, G. S. (2003). Evolving concepts of rheumatoid arthritis. Nature 423, 356–361. 5. Harris, E. D. (2004). In “Kelley’s Textbook of Rheumatology” (E. D. Harris, S. Ruddy, C. B. Sledge, J. S. Sergent, and R. C. Budd, eds.). WB Saunders, Philadelphia, PA. 6. Fassbender, H. G. (1983). Histomorphological basis of articular cartilage destruction in rheumatoid arthritis. Coll. Rel. Dis. 3, 141–155. 7. Krane, S. M., Conca, W., Stephenson, M. L., Amento, E. P., and Goldring, M. B. (1990). Mechanisms of matrix degradation in rheumatoid arthritis. Ann. NY Acad. Sci. 580, 340–354. 8. Harris, E. D. J. (1990). Rheumatoid arthritis. Pathophysiology and implications for therapy. N. Engl. J. Med. 322, 1277–1289. 9. Choy, E. H., and Panayi, G. S. (2001). Cytokine pathways and joint inflammation in rheumatoid arthritis. N. Engl. J. Med. 344, 907–916. 10. Corrigall, V. M., and Panayi, G. S. (2002). Autoantigens and immune pathways in rheumatoid arthritis. Crit. Rev. Immunol. 22, 281–293. 11. Combe, B., Dougados, M., Goupille, P., Cantagrel, A., Eliaou, J. F., Sibilia, J., Meyer, O., Sany, J., Daures, J. P., and Dubois, A. (2001). Prognostic factors for radiographic damage in early rheumatoid arthritis: a multiparameter prospective study. Arthritis Rheum. 44, 1736–1743.

Chapter 51 Rheumatoid Arthritis and other Inflammatory Joint Pathologies 12. Jansen, L. M., van der Horst-Bruinsma, I. E., van Schaardenburg, D., Bezemer, P. D., and Dijkmans, B. A. (2001). Predictors of radiographic joint damage in patients with early rheumatoid arthritis. Ann. Rheum. Dis. 60, 924–927. 13. van der Heijde, D. M., van Riel, P. L., van Leeuwen, M. A., van‘t Hof, M. A., van Rijswijk, M. H., and van de Putte, L. B. (1992). Prognostic factors for radiographic damage and physical disability in early rheumatoid arthritis. A prospective follow-up study of 147 patients. Br. J. Rheumatol. 31, 519–525. 14. Mottonen, T., Paimela, L., Leirisalo-Repo, M., Kautiainen, H., Ilonen, J., and Hannonen, P. (1998). Only high disease activity and positive rheumatoid factor indicate poor prognosis in patients with early rheumatoid arthritis treated with “sawtooth” strategy. Ann. Rheum. Dis. 57, 533–539. 15. Goronzy, J. J., Matteson, E. L., Fulbright, J. W., Warrington, K. J., Chang-Miller, A., Hunder, G. G., Mason, T. G., Nelson, A. M., Valente, R. M., Crowson, C. S., Erlich, H. A., Reynolds, R. L., Swee, R. G., O’Fallon, W. M., and Weyand, C. M. (2004). Prognostic markers of radiographic progression in early rheumatoid arthritis. Arthritis Rheum. 50, 43–54. 16. Mulherin, D., Fitzgerald, O., and Bresnihan, B. (1996). Clinical improvement and radiological deterioration in rheumatoid arthritis: evidence that the pathogenesis of synovial inflammation and articular erosion may differ. Br. J. Rheumatol. 35, 1263–1268. 17. Kirwan, J., Byron, M., and Watt, I. (2001). The relationship between soft tissue swelling, joint space narrowing and erosive damage in hand X-rays of patients with rheumatoid arthritis. Rheumatology (Oxford) 40, 297–301. 18. van den Berg, W. B. (2001). Uncoupling of inflammatory and destructive mechanisms in arthritis. Semin. Arthritis Rheum. 30, 7–16. 19. Graudal, N. (2004). The natural history and prognosis of rheumatoid arthritis: association of radiographic outcome with process variables, joint motion and immune proteins. Scand. J. Rheumatol. Suppl. 1–38. 20. Garnero, P., Rousseau, J. C., and Delmas, P. D. (2000). Molecular basis and clinical use of biochemical markers of bone, cartilage, and synovium in joint diseases. Arthritis Rheum. 43, 953–968. 21. Garnero, P., Geusens, P., and Landewe, R. (2003). Biochemical markers of joint tissue turnover in early rheumatoid arthritis. Clin. Exp. Rheumatol. 21, S54–58. 22. Garnero, P., and Delmas, P. D. (2004). Noninvasive techniques for assessing skeletal changes in inflammatory arthritis: bone biomarkers. Curr. Opin. Rheumatol. 16, 428–434. 23. Mansson, B., Carey, D., Alini, M., Ionescu, M., Rosenberg, L. C., Poole, A. R., Heinegard, D., and Saxne, T. (1995). Cartilage and bone metabolism in rheumatoid arthritis. Differences between rapid and slow progression of disease identified by serum markers of cartilage metabolism. J. Clin. Invest. 95, 1071–1077. 24. Poole, A. R., Ionescu, M., Swan, A., and Dieppe, P. A. (1994). Changes in cartilage metabolism in arthritis are reflected by altered serum and synovial fluid levels of the cartilage proteoglycan aggrecan. Implications for pathogenesis. J. Clin. Invest. 94, 25–33. 25. Garnero, P., Landewe, R., Boers, M., Verhoeven, A., Van Der Linden, S., Christgau, S., Van Der Heijde, D., Boonen, A., and Geusens, P. (2002). Association of baseline levels of markers of bone and cartilage degradation with long-term progression of joint damage in patients with early rheumatoid arthritis: the COBRA study. Arthritis Rheum. 46, 2847–2856. 26. Christgau, S., Garnero, P., Fledelius, C., Moniz, C., Ensig, M., Gineyts, E., Rosenquist, C., and Qvist, P. (2001). Collagen type II C-telopeptide fragments as an index of cartilage degradation. Bone 29, 209–215. 27. Kojima, T., Mwale, F., Yasuda, T., Girard, C., Poole, A. R., and Laverty, S. (2001). Early degradation of type IX and type II collagen with the onset of experimental inflammatory arthritis. Arthritis Rheum. 44, 120–127. 28. Fraser, A., Fearon, U., Billinghurst, R. C., Ionescu, M., Reece, R., Barwick, T., Emery, P., Poole, A. R., and Veale, D. J. (2003). Turnover of

29.

30.

31.

32.

33.

34.

35.

36.

37.

38. 39.

40.

41.

42.

43.

44.

45.

859

type II collagen and aggrecan in cartilage matrix at the onset of inflammatory arthritis in humans: relationship to mediators of systemic and local inflammation. Arthritis Rheum. 48, 3085–3095. Lindqvist, E., Eberhardt, K., Bendtzen, K., Heinegard, D., and Saxne, T. (2005). Prognostic laboratory markers of joint damage in rheumatoid arthritis. Ann. Rheum. Dis. 64, 196–201. Deodhar, A. A., and Woolf, A. D. (1996). Bone mass measurement and bone metabolism in rheumatoid arthritis: a review. Br. J. Rheumatol. 35, 309–322. Goldring, S. R. (1996). Osteoporosis and rheumatic diseases. In “Primer on the Metabolic Bone Diseases and Disorders of Mineral Metabolism, 3rd Edition” (M. J. Favus, ed.), pp. 299–301. Lippincott-Raven, Philadelphia, PA. Goldring, S. R., and Polisson, R. P. (1998). Bone disease in rheumatological disorders. In “Metabolic Bone Disease, 2nd Edition” (L. Avioli, and S. M. Krane, eds.), pp. 621–635. Academic Press, San Diego, CA. Bromley, M., and Woolley, D. E. (1984). Chondroclasts and osteoclasts at subchondral sites of erosions in the rheumatoid joint. Arthritis Rheum. 27, 968–975. Bromley, M., and Woolley, D. E. (1984). Histopathology of the rheumatoid lesion; identification of cell types at sites of cartilage erosion. Arthritis Rheum. 27, 857–863. Allard, S. A., Muirden, K. D., and Maini, R. N. (1991). Correlation of histopathological features of pannus with patterns of damage in different joints in rheumatoid arthritis. Ann. Rhenum. Dis. 50, 278–283. Ashton, B. A., Ashton, I. K., Marshall, M. J., and Butler, R. C. (1993). Localization of vitronectin receptor immunoreactivity and tartrate resistant acid phosphatase activity in synovium from patients with inflammatory and degenrative arthritis. Ann. Rheum. Dis. 52, 133–137. Chang, J. S., Quinn, J. M., Demaziere, A., Bulstrode, C. J., Francis, M. J., Duthie, R. B., and Athanasou, N. A. (1992). Bone resorption by cells isolated from rheumatoid synovium. Ann. Rheum. Dis. 51, 1223–1229. Chambers, T. J., Thomson, B. M., and Fuller, K. (1984). Resorption of bone by isolated rabbit osteoclasts. J. Cell. Sci. 66, 383. Hummel, K. M., Petrow, P. K., Franz, J. K., Muller-Ladner, U., Aicher, W. K., Gay, R. E., Bromme, D., and Gay, S. (1998). Cysteine proteinase cathepsin K mRNA is expressed in synovium of patients with rheumatoid arthritis and is detected at sites of synovial bone destruction. J. Rheumatol. 25, 1887–1894. Hattersley, G., and Chambers, T. J. (1989). Calcitonin receptors as markers for osteoclastic differentiation: correlation between generation of bone-resorptive cells and cells that express calcitonin receptors in mouse bone marrow cultures. Endocrinology 125, 1606–1612. Fujikawa, Y., Sabokbar, A., Neale, S., and Athanasou, N. A. (1993). Human osteoclast formation and bone resorption by monocytes and synovial macrophages in rheumatoid arthritis. Ann. Rheum. Dis. 55, 816–822. Itonaga, I., Fujikawa, Y., Sabokbar, A., Murray, D. W., and Athanasou, N. A. (2000). Rheumatoid arthritis synovial macrophageosteoclast differentiation is osteoprotegerin ligand-dependent. J. Pathol. 192, 97–104. Takayanagi, H., Iizuka, H., Juji, T., Nakagawa, T., Yamamoto, A., Miyazaki, T., Koshihara, Y., Oda, H., Nakamura, K., and Tanaka, S. (2000). Involvement of receptor activator of nuclear factor κB ligand/ osteoclast differentiation factor in osteoclastogenesis from synoviocytes in rheumatoid arthritis. Arthritis Rheum. 43, 259–269. Haynes, D. R., Crotti, T. N., Loric, M., Bain, G. I., Atkins, G. J., and Findlay, D. M. (2001). Osteoprotegerin and receptor activator of nuclear factor κB ligand (RANKL) regulate osteoclast formation by cells in the human rheumatoid arthritic joint. Rheumatology (Oxford) 40, 623–630. Suzuki, Y., Tsutsumi, Y., Nakagawa, M., Suzuki, H., Matsushita, K., Beppu, M., Aoki, H., Ichikawa, Y., and Mizushima, Y. (2001). Osteoclastlike cells in an in vitro model of bone destruction by rheumatoid synovium. Rheumatology (Oxford) 40, 673–682.

860 46. Gravallese, E. M., Harada, Y., Wang, J.-T., Gorn, A., Thornhill, T., and Goldring, S. R. (1998). Identification of cell types responsible for bone resorption in rheumatoid arthritis and juvenile rheumatoid arthritis. Am. J. Pathol. 152, 943–951. 47. Gorn, A. H., Lin, H. Y., Yamin, M., Auron, P. E., Flannery, M. R., Tapp, D. R., Manning, C. A., Lodish, H. F., Krane, S. M., and Goldring, S. R. (1992). The cloning, characterization and expression of a human calcitonin receptor from an ovarian carcinoma cell line. J. Clin. Invest. 90, 1726–1735. 48. Gorn, A. H., Rudolph, S. M., Flannery, M. R., Morton, C., Weremowicz, S., Wang, J.-T., Krane, S. M., and Goldring, S. R. (1995). Expression of two human skeletal calcitonin receptor isoforms cloned from a giant cell tumor of bone. J. Clin. Invest. 95, 2680–2691. 49. Lee, S. K., Goldring, S. R., and Lorenzo, J. A. (1995). Expression of the calcitonin receptor in bone marrow cell cultures and in bone: a specific marker of the differentiated osteoclast that is regulated by calcitonin. Endocrinology 136, 4572–4581. 50. Rodan, G. A., and Martin, T. J. (1981). Role of osteoblasts in hormonal control of bone resorption – a hypothesis. Calcif. Tissue Int. 33, 349–351. 51. Raisz, L. G. (1988). Local and systemic factors in the pathogenesis of osteoporosis. N. Engl. J. Med. 318, 818–828. 52. Suda, T., Takahashi, N., and Martin, T. J. (1992). Modulation of osteoclast differentiation. Endocr. Rev. 13, 66–80. 53. Pettit, A. R., Ji, H., von Stechow, D., Muller, R., Goldring, S. R., Choi, Y., Benoist, C., and Gravallese, E. M. (2001). TRANCE/RANKL knockout mice are protected from bone erosion in a serum transfer model of arthritis. Am. J. Pathol. 159, 1689–1699. 54. Kong, Y. Y., Yoshida, H., Sarosi, I., Tan, H. L., Timms, E., Capparelli, C., Morony, S., Oliveira-dos-Santos, A. J., Van, G., Itie, A., Khoo, W., Wakeham, A., Dunstan, C. R., Lacey, D. L., Mak, T. W., Boyle, W. J., and Penninger, J. M. (1999). OPGL is a key regulator of osteoclastogenesis, lymphocyte development and lymph-node organogenesis. Nature 397, 315–323. 55. Korganow, A., Weber, J. C., Martin, T. (1999). Animal models and autoimmune diseases. Rev. Med. Interne. 20, 283–286. 56. Matsumoto, I., Staub, A., Benoist, C., and Mathis, D. (1999). Arthritis provoked by linked T and B cell recognition of a glycolytic enzyme. Science 286, 1732–1735. 57. Matsumoto, I., Maccioni, M., Lee, D. M., Maurice, M., Simmons, B., Brenner, M., Mathis, D., and Benoist, C. (2002). How antibodies to a ubiquitous cytoplasmic enzyme may provoke joint-specific autoimmune disease. Nat. Immunol. 3, 360–365. 58. Redlich, K., Hayer, S., Ricci, R., David, J., Tohidast-Akrad, M., Kollias, G., Steiner, G., Smolen, J., Wagner, E., and Schett, G. (2002). Osteoclasts are essential for TNF-α-mediated joint destruction. J. Clin. Invest. 110, 1419–1427. 59. Teitelbaum, S. (2000). Bone resorption by osteoclasts. Science 289, 1504–1508. 60. Grigoriadis, A. E., Wang, Z. Q., Cecchini, M. G., Hofstetter, W., Felix, R., Fleisch, H. A., and Wagner, E. F. (1994). c-Fos: a key regulator of osteoclast-macrophage lineage determination and bone remodeling. Science 266, 443–448. 61. Gravallese, E. M., and Goldring, S. R. (2000). Cellular mechanisms and the role of cytokines in bone erosions in rheumatoid arthritis. Arthritis Rheum. 43, 2143–2151. 62. Feldmann, M., and Maini, R. N. (1999). The role of cytokines in the pathogenesis of rheumatoid arthritis. Rheumatology 38, 3–7. 63. Funk, J. L., Cordaro, L. A., Wei, H., Benjamin, J. B., and Yocum, D. E. (1998). Synovium as a source of increased amino-terminal parathyroid hormone-related protein expression in rheumatoid arthritis; a possible role for locally produced parathyroid hormone-related protein in the pathogenesis of rheumatoid arthritis. J. Clin. Invest. 101, 1362–1371. 64. Chabaud, M., Lubberts, E., Joosten, L., van Den Berg, W., and Miossec, P. (2001). IL-17 derived from juxta-articular bone and

Steven R. Goldring and Mary B. Goldring

65.

66.

67. 68.

69.

70.

71.

72.

73.

74.

75.

76.

77.

78.

synovium contributes to joint degradation in rheumatoid arthritis. Arthritis Res. 3, 168–177. Ogata, Y., Kukita, A., Kukita, T., Komine, M., Miyahara, A., Miyazaki, S., and Kohashi, O. (1999). A novel role of IL-15 in the development of osteoclasts: inability to replace its activity with IL-2. J. Immunol. 162, 2754–2760. Okano, K., Tsukazaki, T., Ohtsuru, A., Namba, H., Osaki, M., Iwasaki, K., and Yamashita, S. (1996). Parathyroid hormone-related peptide in synovial fluid and disease activity of rheumatoid arthritis. Br. J. Rheumatol. 35, 1056–1062. Romas, E., and Martin, T. J. (1997). Cytokines in the pathogenesis of osteoporosis. Osteoporos. Int. 7, S47–S53. Horwood, N. J., Kartsogiannis, V., Quinn, J. M., Romas, E., Martin, T. J., and Gillespie, M. T. (1999). Activated T lymphocytes support osteoclast formation in vitro. Biochem. Biophys. Res. Commun. 265, 144–150. Wong, B. R., Josien, R., Lee, S. Y., Sauter, B., Li, H. L., Steinman, R. M., and Choi, Y. (1997). TRANCE (tumor necrosis factor [TNF]-related activation-induced cytokine), a new TNF family member predominantly expressed in T cells, is a dendritic cell-specific survival factor. J. Exp. Med. 186, 2075–2080. Wong, B. R., Rho, J., Arron, J., Robinson, E., Orlinick, J., Chao, M., Kalachikov, S., Cayani, E., Bartlett, F. S., 3rd, Frankel, W. N., Lee, S. Y., and Choi, Y. (1997). TRANCE is a novel ligand of the tumor necrosis factor receptor family that activates c-Jun N-terminal kinase in T cells. J. Biol. Chem. 272, 25190–25194. Josien, R., Wong, B. R., Li, H. L., Steinman, R. M., and Choi, Y. (1999). TRANCE, a TNF family member, is differentially expressed on T cell subsets and induces cytokine production in dendritic cells. J. Immunol. 162, 2562–2568. Anderson, D. M., Maraskovsky, E., Billingsley, W. L., Dougall, W. C., Tometsko, M. E., Roux, E. R., Teepe, M. C., DuBose, R. F., Cosman, D., and Galibert, L. (1997). A homologue of the TNF receptor and its ligand enhance T-cell growth and dendritic-cell function. Nature 390, 175–179. Lacey, D. L., Timms, E., Tan, H. L., Kelley, M. J., Dunstan, C. R., Burgess, T., Elliott, R., Colombero, A., Elliott, G., Scully, S., Hsu, H., Sullivan, J., Hawkins, N., Davy, E., Capparelli, C., Eli, A., Qian, Y. X., Kaufman, S., Sarosi, I., Shalhoub, V., Senaldi, G., Guo, J., Delaney, J., and Boyle, W. J. (1998). Osteoprotegerin ligand is a cytokine that regulates osteoclast differentiation and activation. Cell 93, 165–176. Yasuda, H., Shima, N., Nakagawa, N., Mochizuki, S. I., Yano, K., Fujise, N., Sato, Y., Goto, M., Yamaguchi, K., Kuriyama, M., Kanno, T., Murakami, A., Tsuda, E., Morinaga, T., and Higashio, K. (1998). Identity of osteoclastogenesis inhibitory factor (OCIF) and osteoprotegerin (OPG): a mechanism by which OPG/OCIF inhibits osteoclastogenesis in vitro. Endocrinology 139, 1329–1337. Bucay, N., Sarosi, I., Dunstan, C. R., Morony, S., Tarpley, J., Capparelli, C., Scully, S., Tan, H. L., Xu, W., Lacey, D. L., Boyle, W. J., and Simonet, W. S. (1998). Osteoprotegerin-deficient mice develop early onset osteoporosis and arterial calcification. Genes Dev. 12, 1260–1268. Emery, J. G., McDonnell, P., Burke, M. B., Deen, K. C., Lyn, S., Silverman, C., Dul, E., Appelbaum, E. R., Eichman, C., DiPrinzio, R., Dodds, R. A., James, I. E., Rosenberg, M., Lee, J. C., and Young, P. R. (1998). Osteoprotegerin is a receptor for the cytotoxic ligand TRAIL. J. Biol. Chem. 273, 14363–14367. Simonet, W. S., Lacey, D. L., Dunstan, C. R., Kelley, M., Chang, M. S., Luthy, R., Nguyen, H. Q., Wooden, S., Bennett, L., Boone, T., Shimamoto, G., DeRose, M., Elliott, R., Colombero, A., Tan, H. L., Trail, G., Sullivan, J., Davy, E., Bucay, N., Renshaw-Gegg, L., Hughes, T. M., Hill, D., Pattison, W., Campbell, P., Boyle, W. J., et al. (1997). Osteoprotegerin: a novel secreted protein involved in the regulation of bone density. Cell 89, 309–319. Romas, E., Gillespie, M. T., and Martin, T. J. (2002). Involvement of receptor activator of NFκB ligand and tumor necrosis factor-κ in bone destruction in rheumatoid arthritis. Bone 30, 340–346.

Chapter 51 Rheumatoid Arthritis and other Inflammatory Joint Pathologies 79. Redlich, K., Hayer, S., Maier, A., Dunstan, C., Tohidast-Akrad, M., Lang, S., Turk, B., Pietschmann, P., Woloszczuk, W., Haralambous, S., Kollias, G., Steiner, G., Smolen, J., and Schett, G. (2002). Tumor necrosis factor-α-mediated joint destruction is inhibited by targeting osteoclasts with osteoprotegerin. Arthritis Rheum. 46, 785–792. 80. Weitzmann, M. N., Cenci, S., Rifas, L., Brown, C., and Pacifici, R. (2000). Interleukin-7 stimulates osteoclast formation by up-regulating the T- cell production of soluble osteoclastogenic cytokines. Blood 96, 1873–1878. 81. Weitzmann, M. N., Cenci, S., Rifas, L., Haug, J., Dipersio, J., and Pacifici, R. (2001). T cell activation induces human osteoclast formation via receptor activator of nuclear factor κB ligand-dependent and -independent mechanisms. J. Bone Miner. Res. 16, 328–337. 82. Kotake, S., Udagawa, N., Hakoda, M., Mogi, M., Yano, K., Tsuda, E., Takahashi, K., Furuya, T., Ishiyama, S., Kim, K. J., Saito, S., Nishikawa, T., Takahashi, N., Togari, A., Tomatsu, T., Suda, T., and Kamatani, N. (2001). Activated human T cells directly induce osteoclastogenesis from human monocytes: possible role of T cells in bone destruction in rheumatoid arthritis patients. Arthritis Rheum. 44, 1003–1012. 83. Stewart, A., Mackenzie, L. M., Black, A. J., and Reid, D. M. (2004). Predicting erosive disease in rheumatoid arthritis. A longitudinal study of changes in bone density using digital X-ray radiogrammetry: a pilot study. Rheumatology (Oxford) 43, 1561–1564. 84. Shimizo, S., Shiozawa, S., Shiozawa, K., Imura, S., and Fugita, T. (1985). Quantitative histological studies on the pathogenesis of periarticular osteoporosis in rheumatoid arthritis. Arthritis Rheum. 28, 25–31. 85. Deleuran, B. W., Chu, C. Q., Field, M., Brennan, F. M., Katsikis, P. M. F., and Maini, R. N. (1992). Localization of interleukin-1α, type 1 interleukin-1 receptor and interleukin-1 receptor antagonis in the synovial membrane and cartilage/pannus junction in rhumatoid arthritis. Br. J. Rheumatol. 31, 801–809. 86. Firestein, G. S., Alcaro-Garcia, J. M., and Maki, R. (1990). Quantitative analysis of cytokine gene expression in rheumatoid arthritis. J. Immunol. 144, 3347–3353. 87. Chu, C. Q., Field, M., Feldman, M., and Maini, R. N. (1991). Localization of tumour necrosis factor-a in synovial tissue and at the cartilage-pannus junction in patients with rheumatoid arthritis. Arthritis Rheum. 34, 1125–1132. 88. Chu, C. Q. M. F., Allard, S., Abney, E., Feldmann, M., and Maini, R. N. (1992). Detection of cytokines at the cartilage/pannus junction in patients with rheumatoid arthritis; implications for the role of cytokines in cartilage destruction and repair. Br. J. Rheumatol. 32, 653–661. 89. Mazess, R. B., and Whedon, G. D. (1983). Immobilization and bone. Calcif. Tiss. Int. 32, 265. 90. Joffe, I., and Epstein, S. (1991). Osteoporosis associated with rheumatoid arthritis: pathogenesis and management. Semin. Arthritis Rheum. 20, 256–272. 91. Spector, T. D., Hall, G. M., McCloskey, E. V., and Kanis, J. A. (1993). Risk of vertebral fracture in women with rheumatoid arthritis. BMJ 306, 558. 92. Gough, A. K., Lilley, J., Eyre, S., Holdin, R., and Emery, P. (1994). Generalized bone loss in patients with rheumatoid arthritis. Lancet 344, 23–27. 93. Hooyman, J. R., Melton, L. J., Nelson, A. M., O’Fallon, W. M., and Riggs, B. L. (1984). Fractures after rheumatoid arthritis; a population based study. Arthritis Rheum. 27, 1353–1361. 94. Beat, A. M., Bloch, D. A., and Fries, J. F. (1991). Predictors of fractures in early rheumatoid arthritis. J. Rheumatol. 18, 804–808. 95. Haugeberg, G., Orstavik, R. E., and Kvien, T. K. (2003). Effects of rheumatoid arthritis on bone. Curr. Opin. Rheumatol. 15, 469–475. 96. Forslind, K., Keller, C., Svensson, B., and Hafstrom, I. (2003). Reduced bone mineral density in early rheumatoid arthritis is associated with radiological joint damage at baseline and after 2 years in women. J. Rheumatol. 30, 2590–2596.

861

97. Orstavik, R. E., Haugeberg, G., Mowinckel, P., Hoiseth, A., Uhlig, T., Falch, J. A., Halse, J. I., McCloskey, E., and Kvien, T. K. (2004). Vertebral deformities in rheumatoid arthritis: a comparison with population-based controls. Arch. Intern. Med. 164, 420–425. 98. Orstavik, R. E., Haugeberg, G., Uhlig, T., Mowinckel, P., Falch, J. A., Halse, J. I., and Kvien, T. K. (2005). Incidence of vertebral deformities in 255 female rheumatoid arthritis patients measured by morphometric X-ray absorptiometry. Osteoporos. Int. 16, 35–42. 99. Lodder, M. C., de Jong, Z., Kostense, P. J., Molenaar, E. T., Staal, K., Voskuyl, A. E., Hazes, J. M., Dijkmans, B. A., and Lems, W. F. (2004). Bone mineral density in patients with rheumatoid arthritis: relation between disease severity and low bone mineral density. Ann. Rheum. Dis. 63, 1576–1580. 100. Lane, N. E., and Goldring, S. R. (1998). Bone loss in rheumatoid arthritis: what role does inflammation play. J. Rheumatol. 7, 1251–1253. 101. Sambrook, P., Ansel, B., Foster, S., et al. (1985). Bone turnover in early rheumatoid arthritis; longitudinal bone density studies. Ann. Rheum. Dis. 44, 580. 102. Sambrook, P., Eisman, J., Yeates, M., et al. (1986). Osteoporosis in rheumatoid arthritis: safety of low-dose corticosteroids. Ann. Rheum. Dis. 45, 950. 103. Sambrook, P. N., Eisman, A., Champion, G., Yeates, M. G., Pocock, N. A., and Eberl, S. (1987). Determinants of axial bone loss in rheumatoid arthritis. Arthritis Rheum. 30, 721–728. 104. Sambrook, P., Birmingham, J., Kelly, P., Kempler, S., Freund, J., and Eisman, J. (1992). Postmenopausal bone loss in rheumatoid arthritis: effects of estrogens and androgens. J. Rheum. 19, 357–361. 105. Peel, N. F., Eastell, R., and Russell, R. G. G. (1991). Osteoporosis in rheumatoid arthritis – the laboratory perspective. Br. J. Rheum. 30, 84–85. 106. Woolf, A. D. (1991). Osteoporosis in rheumatoid arthritis – the clinical viewpoint. Br. J. Rheum. 30, 82–84. 107. Compston, J. E., Vedi, S., Croucher, P. I., Garrahan, N. J., and O’Sullivan, M. M. (1994). Bone turnover in non-steroid treated rheumatoid arthritis. Ann. Rheum. Dis. 53, 163–166. 108. Sambrook, P., Spector, T., Seeman, E., Bellamy, N., Russel, R. D. D. N. M., Prince, R., Owen, E., Silman, A., and Eisman, J. (1995). Osteoporosis in rheumatoid arthritis: a monozygotic co-twin control study. Arthritis Rheum. 38, 806–809. 109. Laan, R. F., van-Riel, P. L., and van-de-Putte, L. B. (1992). Bone mass in patients with rheumatoid arthritis. Ann. Rheum. Dis. 51, 826–832. 110. Als, O. S., Gotfredsen, A., Riis, B. J., and Christiansen, C. (1985). Are disease duration and degree of functional impairment determinanats of bone loss in rheumatoid arthritis? Ann. Rhem. Dis. 44, 406–411. 111. Kroger, H., Arnala, I., and Alhava, E. M. (1991). Bone remodeling in osteoporosis associated with rheumatoid arthritis. Calcif. Tiss. Int. 49, S90. 112. Mellish, R. W. E., O’Sullivan, M. M., Garrahan, N. J., and Compston, J. E. (1987). Iliac crest trabecular bone mass and structure in patients with non-steroid treated rheumatoid arthritis. Ann. Rheum. Dis. 46, 830–836. 113. Hall, G. M., Spector, T. D., and Delmas, P. D. (1995). Markers of bone metabolism in postmenopausal women with rheumatoid arthritis. Effects of corticosteroids and hormone replacement therapy. Arthritis Rheum. 38, 902–906. 114. Gough, A. K., Peel, N. F., Eastell, R., Holder, R. L., Lilley, J., and Emery, P. (1994). Excretion of pyridinium crosslinks correlates with disease activity and appendicular bone loss in early rheumatoid arthritis. Ann. Rheum. Dis. 53, 14–17. 115. Garnero, P., Gineyts, E., Christgau, S., Finck, B., and Delmas, P. D. (2002). Association of baseline levels of urinary glucosyl-galactosylpyridinoline and type II collagen C-telopeptide with progression of joint destruction in patients with early rheumatoid arthritis. Arthritis Rheum. 46, 21–30.

862 116. Iwamoto, J., Takeda, T., and Ichimura, S. (2003). Urinary cross-linked N-telopeptides of type I collagen levels in patients with rheumatoid arthritis. Calcif. Tissue Int. 72, 491–497. 117. Gough, A., Sambrook, P., Devlin, J., Huissoon, A., Njeh, C., Robbins, S., Nguyen, T., and Emery, P. (1998). Osteoclastic activation is the principal mechanism leading to secondary osteoporosis in rheumatoid arthritis. J. Rheumatol. 7, 1282–1289. 118. Black, D., Marabani, M., Sturrock, R. D., and Robins, S. P. (1989). Urinary excretion of the hydroxypyridinium cross links of collagen in patients with rheumatoid arthritis. Ann. Rheum. Dis. 48, 641–644. 119. Robins, S. P., Stewart, P., Astbury, C., and Bird, H. A. (1986). Measurement of the cross linking compound, pyridinoline, in urine as an index of collagen degradation in joint disease. Ann. Rheum. Dis. 45, 969–973. 120. Siebel, M. J., Duncan, A., and Robins, S. P. (1989). Urinary hydroxypyridinium crosslinks provide indices of cartilage and bone involvement in arthritis disease. J. Rheumatol. 16, 964–970. 121. Sinigaglia, L., Varenna, M., Binelli, L., Bartucci, F., Arrigoni, M., Ferrara, R., and Abbiati, G. (1995). Urinary and synovial pyridinium crosslink concentrations in patients with rheumatoid arthritis and osteoarthritis. Ann. Rheum. Dis. 54, 144–147. 122. Garnero, P., Jouvenne, P., Buchs, N., Delmas, P. D., and Miossec, P. (1999). Uncoupling of bone metabolism in rheumatoid arthritis patients with or without joint destruction: assessment with serum type I collagen breakdown products. Bone 24, 381–385. 123. Saag, K. G., Emkey, R., Schnitzer, T. J., Brown, J. P., Hawkins, F., Goemaere, S., Thamsborg, G., Liberman, U. A., Delmas, P. D., Malice, P., Czachur, M., and Daifotis, A. G. (1998). Alendronate for the prevention and treatment of glucocorticoid-induced osteoporosis. N. Engl. J. Med. 339, 292–299. 124. Ritchlin, C. T., Haas-Smith, S. A., Li, P., Hicks, D. G., and Schwarz, E. M. (2003). Mechanisms of TNF-α- and RANKLmediated osteoclastogenesis and bone resorption in psoriatic arthritis. J. Clin. Invest. 111, 821–831. 125. McGonagle, D., Benjamin, M., Marzo-Ortega, H., and Emery, P. (2002). Advances in the understanding of entheseal inflammation. Curr. Rheumatol. Rep. 4, 500–506. 126. Benjamin, M., and McGonagle, D. (2001). The anatomical basis for disease localisation in seronegative spondyloarthropathy at entheses and related sites. J. Anat. 199, 503–526. 127. Benjamin, M., Moriggl, B., Brenner, E., Emery, P., McGonagle, D., and Redman, S. (2004). The “enthesis organ” concept: why enthesopathies may not present as focal insertional disorders. Arthritis Rheum. 50, 3306–3313. 128. McGonagle, D. (2005). Imaging the joint and enthesis: insights into pathogenesis of psoriatic arthritis. Ann. Rheum. Dis. 64 Suppl 2, ii58–60. 129. Braun, J., Bollow, M., Neure, L., Seipelt, E., Seyrekbasan, F., Herbst, H., Eggens, U., Distler, A., and Sieper, J. (1995). Use of immunohistologic and in situ hybridization techniques in the examination of sacroiliac joint biopsy specimens from patients with ankylosing spondylitis. Arthritis Rheum. 4, 499–505. 130. Lories, R. J., Derese, I., Ceuppens, J. L., and Luyten, F. P. (2003). Bone morphogenetic proteins 2 and 6, expressed in arthritic synovium, are regulated by proinflammatory cytokines and differentially modulate fibroblast-like synoviocyte apoptosis. Arthritis Rheum. 48, 2807–2818. 131. Franck, H., and Ittel, T. (2000). Serum osteocalcin levels in patients with psoriatic arthritis: an extended report. Rheumatol. Int. 19, 161–164. 132. Spencer, D., Park, W., Dick, H., et al. (1979). Radiologic manifestations in 200 patients with ankylosing spondylitis; correlations with clinical features and HLA-B27. J. Rheumatol. 6, 305. 133. Will, R., Bhalla, A., Palmer, R., Ring, F., and Calin, A. (1989). Osteoporosis in early ankylosing spondylitis; a primary pathological event? Lancet 23, 1483–1485.

Steven R. Goldring and Mary B. Goldring 134. Frediani, B., Allegri, A., Falsetti, P., Storri, L., Bisogno, S., Baldi, F., Filipponi, P., and Marcolongo, R. (2001). Bone mineral density in patients with psoriatic arthritis. J. Rheumatol. 28, 138–143. 135. Ralston, S. H., Urquhart, G. D., Brzeski, M., and Sturrock, R. D. (1990). Prevalence of vertebral compression fractures due to osteoporosis in ankylosing spondylitis. BMJ 300, 563–565. 136. Devogelaer, J.-P., Maldague, B., Malghem, J., and de Deuxchaisnes, C. N. (1992). Appendicular and vertebral bone mass in ankylosing spondylitis. A comparison of plain radiographs with single- and dual-photon absorptiometry and with quantitative computed tomography. Arthritis Rheum. 35, 1062–1067. 137. Marzo-Ortega, H., McGonagle, D., Haugeberg, G., Green, M. J., Stewart, S. P., and Emery, P. (2003). Bone mineral density improvement in spondyloarthropathy after treatment with etanercept. Ann. Rheum. Dis. 62, 1020–1021. 138. Ansell, B. M., and Bywaters, E. G. L. (1956). Growth in Still’s Disease. Ann. Rheum. Dis. 15, 295–318. 139. Bernstein, B. H., Stobie, D., Singsen, B. H., Koster-King, K., Kornreich, H. K., and Hanson, V. (1977). Growth retardation in juvenile rheumatoid arthritis (JRA). Arthritis Rheum. 20, 212–216. 140. Hillman, L., Cassidy, J. T., Johnson, L., Lee, D., and Allen, S. (1994). Vitamin D metabolism and bone mineralization in children with juvenile rheumatoid arthritis. J. Pediatr. 124, 910–916. 141. Falcini, F., Ermini, M., and Bagnoli, F. (1998). Bone turnover is reduced in children with juvenile rheumatoid arthritis. J. Endocrinol. Invest. 21, 31–36. 142. Hopp, R., Degan, J. A., Gallagher, J. C., and Cassidy, J. T. (1991). Estimation of bone mineral density in children with juvenile rheumatoid arthritis. J. Rheumatol. 18, 1235–1239. 143. Henderson, C. J., Specker, B. L., Sierra, R. I., Campaigne, B. N., and Lovell, D. J. (2000). Total-body bone mineral content in noncorticosteroid-treated postpubertal females with juvenile rheumatoid arthritis: frequency of osteopenia and contributing factors. Arthritis Rheum. 43, 531–540. 144. Rabinovich, C. E. (2000). Bone mineral status in juvenile rheumatoid arthritis. J. Rheumatol. Suppl. 58, 34–37. 145. French, A. R., Mason, T., Nelson, A. M., Crowson, C. S., O’Fallon, W. M., Khosla, S., and Gabriel, S. E. (2002). Osteopenia in adults with a history of juvenile rheumatoid arthritis. A population based study. J. Rheumatol. 29, 1065–1070. 146. Reed, A., Haugen, M., Pachman, L., and Langman, C. (1993). Repair of osteopenia in children with juvenile rheumatoid arthritis. J. Pediatr. 122, 693–696. 147. Badley, B. W. D., and Ansell, B. M. (1960). Fracture’s in Still’s disease. Ann. Rheum. Dis. 19, 135–138. 148. Varonos, S., Ansell, B. M., and Reeve, J. (1987). Vertebral collapse in juvenile chronic arthritis; its relationship with glucocorticoid therapy. Calcif. Tiss. Int. 41, 75–78. 149. Reeve, J., Loftus, J., Hesp, R., Ansell, B. M., Wright, D. J., and Woo, P. M. (1993). Biochemical prediction of changes in spinal bone mass in juvenile chronic (or rheumatoid) arthritis treated with glucocorticoids. J. Rheum. 20, 1189–1195. 150. Poole, A. R., Kojima, T., Yasuda, T., Mwale, F., Kobayashi, M., and Laverty, S. (2001). Composition and structure of articular cartilage: a template for tissue repair. Clin. Orthop. S26–33. 151. Poole, A. R. (2001). Cartilage in health and disease. In “Arthritis and Allied Conditions: a Textbook of Rheumatology” (W. Koopman, ed.), pp. 226–284. Lippincott Williams & Wilkins, Philadelphia, PA. 152. Verzijl, N., DeGroot, J., Thorpe, S. R., Bank, R. A., Shaw, J. N., Lyons, T. J., Bijlsma, J. W., Lafeber, F. P., Baynes, J. W., and TeKoppele, J. M. (2000). Effect of collagen turnover on the accumulation of advanced glycation end products. J. Biol. Chem. 275, 39027–39031. 153. Maroudas, A., Bayliss, M. T., Uchitel-Kaushansky, N., Schneiderman, R., and Gilav, E. (1998). Aggrecan turnover in human

Chapter 51 Rheumatoid Arthritis and other Inflammatory Joint Pathologies

154.

155.

156.

157.

158.

159.

160.

161.

162. 163.

164.

165.

166.

167.

168.

169.

170.

articular cartilage: use of aspartic acid racemization as a marker of molecular age. Arch. Biochem. Biophys. 350, 61–71. Kobayashi, I., and Ziff, M. (1975). Electron microscopic studies of the cartilage-pannus junction in rheumatoid arthritis. Arthritis Rheum. 18, 475–483. Woolley, D. E., Crossley, M. J., and Evanson, J. M. (1977). Collagenase at sites of cartilage erosion in the rheumatoid joint. Arthritis Rheum. 20, 1231–1239. Mitchell, N. S., and Shepard, N. (1978). Changes in proteoglycan and collagen in cartilage in rheumatoid arthritis. J. Bone Joint Surg. 60A, 349–354. Dodge, G. R., Pidoux, I., and Poole, A. R. (1991). The degradation of type II collagen in rheumatoid arthritis: an immunoelectron microscopic study. Matrix 11, 330–338. Edwards, J. C. W. (2000). Fibroblast biology. Development and differentiation of synovial fibroblasts in arthritis. Arthritis Res. 2, 344–347. Zvaifler, N. J., Tsai, V., Alsalameh, S., von Kempis, J., Firestein, G. S., and Lotz, M. (1997). Pannocytes: distinctive cells found in rheumatoid arthritis articular cartilage erosions. Am. J. Pathol. 150, 1125–1138. Zvaifler, N. J., and Firestein, G. S. (1994). Pannus and pannocytes. Alternative models of joint destruction in rheumatoid arthritis. Arthritis Rheum. 37, 783–789. Muller-Ladner, U., Kriegsmann, J., Franklin, B. N., Matsumoto, S., Geiler, T., Gay, R. E., and Gay, S. (1996). Synovial fibroblasts of patients with rheumatoid arthritis attach to and invade normal human cartilage when engrafted into SCID mice. Am. J. Pathol. 149, 1607–1615. Kimura, H., Tateishi, H. J., and Ziff, M. (1977). Surface ultrastructure of rheumatoid articular cartilage. Arthritis Rheum. 20, 1085–1098. Dodge, G. R., and Poole, A. R. (1989). Immunohistochemical detection and immunochemical analysis of type II collagen degradation in human normal, rheumatoid, and osteoarthritic articular cartilages and in explants of bovine articular cartilage cultured with interleukin 1. J. Clin. Invest. 83, 647–661. Hembry, R. M., Bagga, M. R., Reynolds, J. J., and Hamblen, D. L. (1995). Immunolocalisation studies on six matrix metalloproteinases and their inhibitors, TIMP-1 and TIMP-2, in synovia from patients with osteo- and rheumatoid arthritis. Ann. Rheum. Dis. 54, 25–32. Walakovits, L. A., Moore, V. L., Bhardwaj, N., Gallick, G. S., and Lark, M. W. (1992). Detection of stromelysin and collagenase in synovial fluid from patients with rheumatoid arthritis and posttraumatic knee injury. Arthritis Rheum. 35, 35–42. Clark, I. M., Powell, L. K., Ramsey, S., Hazleman, B. L., and Cawston, T. E. (1993). The measurement of collagenase, tissue inhibitor of metalloproteinases (TIMP), and collagenase-TIMP complex in synovial fluids from patients with osteoarthritis and rheumatoid arthritis. Arthritis Rheum. 36, 372–379. Ahrens, D., Koch, A. E., Pope, R. M., Stein-Picarella, M., and Niedbala, M. J. (1996). Expression of matrix metalloproteinase 9 (96-kd gelatinase B) in human rheumatoid arthritis. Arthritis Rheum. 39, 1576–1587. Ishiguro, N., Ito, T., Obata, K., Fujimoto, N., and Iwata, H. (1996). Determination of stromelysin-1, 72 and 92 kDa type IV collagenase, tissue inhibitor of metalloproteinase-1 (TIMP-1), and TIMP-2 in synovial fluid and serum from patients with rheumatoid arthritis. J. Rheumatol. 23, 1599–1604. Taylor, D. J., Cheung, N. T., and Dawes, P. T. (1994). Increased serum proMMP-3 in inflammatory arthritides: a potential indicator of synovial inflammatory monokine activity. Ann. Rheum. Dis. 53, 768–772. Manicourt, D. H., Fujimoto, N., Obata, K., and Thonar, E. J. (1995). Levels of circulating collagenase, stromelysin-1, and tissue inhibitor of matrix metalloproteinases 1 in patients with rheumatoid arthritis.

171.

172.

173.

174.

175.

176.

177.

178.

179.

180.

181.

182.

183.

184.

863

Relationship to serum levels of antigenic keratan sulfate and systemic parameters of inflammation [see comments]. Arthritis Rheum. 38, 1031–1039. Yoshihara, Y., Obata, K., Fujimoto, N., Yamashita, K., Hayakawa, T., and Shinmei, M. (1995). Increased levels of stromelysin-1 and tissue inhibitor of metalloproteinases-1 in sera from patients with rheumatoid arthritis. Arthritis Rheum. 38, 969–975. Tetlow, L. C., and Woolley, D. E. (1998). Comparative immunolocalization studies of collagenase 1 and collagenase 3 production in the rheumatoid lesion, and by human chondrocytes and synoviocytes in vitro. B. J. Rheumatol. 37, 64–70. Cunnane, G., Fitzgerald, O., Beeton, C., Cawston, T. E., and Bresnihan, B. (2001). Early joint erosions and serum levels of matrix metalloproteinase 1, matrix metalloproteinase 3, and tissue inhibitor of metalloproteinases 1 in rheumatoid arthritis. Arthritis Rheum. 44, 2263–2274. Cunnane, G., FitzGerald, O., Hummel, K. M., Youssef, P. P., Gay, R. E., Gay, S., and Bresnihan, B. (2001). Synovial tissue protease gene expression and joint erosions in early rheumatoid arthritis. Arthritis Rheum. 44, 1744–1753. Maeda, S., Sawai, T., Uzuki, M., Takahashi, Y., Omoto, H., Seki, M., and Sakurai, M. (1995). Determination of interstitial collagenase (MMP-1) in patients with rheumatoid arthritis. Ann. Rheum. Dis. 54, 970–975. Yoshihara, Y., Nakamura, H., Obata, K., Yamada, H., Hayakawa, T., Fujikawa, K., and Okada, Y. (2000). Matrix metalloproteinases and tissue inhibitors of metalloproteinases in synovial fluids from patients with rheumatoid arthritis or osteoarthritis. Ann. Rheum. Dis. 59, 455–461. Goldbach-Mansky, R., Lee, J. M., Hoxworth, J. M., Smith, D., 2nd, Duray, P., Schumacher, R. H., Jr., Yarboro, C. H., Klippel, J., Kleiner, D., and El-Gabalawy, H. S. (2000). Active synovial matrix metalloproteinase-2 is associated with radiographic erosions in patients with early synovitis. Arthritis Res. 2, 145–153. Tchetverikov, I., Lard, L. R., DeGroot, J., Verzijl, N., TeKoppele, J. M., Breedveld, F. C., Huizinga, T. W., and Hanemaaijer, R. (2003). Matrix metalloproteinases-3, -8, -9 as markers of disease activity and joint damage progression in early rheumatoid arthritis. Ann. Rheum. Dis. 62, 1094–1099. Tchetverikov, I., Ronday, H. K., Van El, B., Kiers, G. H., Verzijl, N., TeKoppele, J. M., Huizinga, T. W., DeGroot, J., and Hanemaaijer, R. (2004). MMP profile in paired serum and synovial fluid samples of patients with rheumatoid arthritis. Ann. Rheum. Dis. 63, 881–883. Giannelli, G., Erriquez, R., Iannone, F., Marinosci, F., Lapadula, G., and Antonaci, S. (2004). MMP-2, MMP-9, TIMP-1 and TIMP-2 levels in patients with rheumatoid arthritis and psoriatic arthritis. Clin. Exp. Rheumatol. 22, 335–338. Yamanaka, H., Matsuda, Y., Tanaka, M., Sendo, W., Nakajima, H., Taniguchi, A., and Kamatani, N. (2000). Serum matrix metalloproteinase 3 as a predictor of the degree of joint destruction during the six months after measurement, in patients with early rheumatoid arthritis. Arthritis Rheum. 43, 852–858. Posthumus, M. D., Limburg, P. C., Westra, J., van Leeuwen, M. A., and van Rijswijk, M. H. (2003). Serum matrix metalloproteinase 3 levels in comparison to C-reactive protein in periods with and without progression of radiological damage in patients with early rheumatoid arthritis. Clin. Exp. Rheumatol. 21, 465–472. Woolley, D. E., and Tetlow, L. C. (1997). Observations on the microenvironmental nature of cartilage degradation in rheumatoid arthritis. Ann. Rheum. Dis. 56, 151–161. Brennan, F. M., Browne, K. A., Green, P. A., Jaspar, J. M., Maini, R. N., and Feldmann, M. (1997). Reduction of serum matrix metalloproteinase 1 and matrix metalloproteinase 3 in rheumatoid arthritis patients following anti-tumour necrosis factor-α (cA2) therapy. Br. J. Rheumatol. 36, 643–650.

864 185. Vandooren, B., Kruithof, E., Yu, D. T., Rihl, M., Gu, J., De Rycke, L., Van Den Bosch, F., Veys, E. M., De Keyser, F., and Baeten, D. (2004). Involvement of matrix metalloproteinases and their inhibitors in peripheral synovitis and down-regulation by tumor necrosis factor α blockade in spondylarthropathy. Arthritis Rheum. 50, 2942–2953. 186. Arner, E. C. (2002). Aggrecanase-mediated cartilage degradation. Curr. Opin. Pharmacol. 2, 322–329. 187. Tortorella, M. D., Burn, T. C., Pratta, M. A., Abbaszade, I., Hollis, J. M., Liu, R., Rosenfeld, S. A., Copeland, R. A., Decicco, C. P., Wynn, R., Rockwell, A., Yang, F., Duke, J. L., Solomon, K., George, H., Bruckner, R., Nagase, H., Itoh, Y., Ellis, D. M., Ross, H., Wiswall, B. H., Murphy, K., Hillman, M. C., Jr., Hollis, G. F., Arner, E. C., et al. (1999). Purification and cloning of aggrecanase-1: a member of the ADAMTS family of proteins. Science 284, 1664–1666. 188. Hurskainen, T. L., Hirohata, S., Seldin, M. F., and Apte, S. S. (1999). ADAM-TS5, ADAM-TS6, and ADAM-TS7, novel members of a new family of zinc metalloproteases. General features and genomic distribution of the ADAM-TS family. J. Biol. Chem. 274, 25555–25563. 189. Abbaszade, I., Liu, R. Q., Yang, F., Rosenfeld, S. A., Ross, O. H., Link, J. R., Ellis, D. M., Tortorella, M. D., Pratta, M. A., Hollis, J. M., Wynn, R., Duke, J. L., George, H. J., Hillman, M. C., Jr., Murphy, K., Wiswall, B. H., Copeland, R. A., Decicco, C. P., Bruckner, R., Nagase, H., Itoh, Y., Newton, R. C., Magolda, R. L., Trzaskos, J. M., Burn, T. C., et al. (1999). Cloning and characterization of ADAMTS11, an aggrecanase from the ADAMTS family. J. Biol. Chem. 274, 23443–23450. 190. Kuno, K., Terashima, Y., and Matsushima, K. (1999). ADAMTS-1 is an active metalloproteinase associated with the extracellular matrix. J. Biol. Chem. 274, 18821–18826. 191. Kuno, K., Okada, Y., Kawashima, H., Nakamura, H., Miyasaka, M., Ohno, H., and Matsushima, K. (2000). ADAMTS-1 cleaves a cartilage proteoglycan, aggrecan. FEBS Lett. 478, 241–245. 192. Flannery, C. R., Little, C. B., Hughes, C. E., and Caterson, B. (1999). Expression of ADAMTS homologues in articular cartilage. Biochem. Biophys. Res. Commun. 260, 318–322. 193. Nagase, H., and Kashiwagi, M. (2003). Aggrecanases and cartilage matrix degradation. Arthritis Res. Ther. 5, 94–103. 194. Okada, Y. (2004). Proteases and matrix degradation. In “Kelley’s Textbook of Rheumatology” (E. D. Harris, S. Ruddy, C. B. Sledge, J. S. Sergent, and R. C. Budd, eds.), pp. 63–81. WB Saunders, Philadelphia, PA. 195. Chevalier, X., Groult, N., Texier, J. M., Larget-Piet, B., and Hornebeck, W. (1996). Elastase activity in cartilage extracts and synovial fluids from subjects with osteoarthritis or rheumatoid arthritis: the prominent role of metalloproteinases. Clin. Exp. Rheumatol. 14, 235–241. 196. Saxne, T., Lecander, I., and Geborek, P. (1993). Plasminogen activators and plasminogen activator inhibitors in synovial fluid. Difference between inflammatory joint disorders and osteoarthritis. J. Rheumatol. 20, 91–96. 197. Keyszer, G. M., Heer, A. H., Kriegsmann, J., Geiler, T., Trabandt, A., Keysser, M., Gay, R. E., and Gay, S. (1995). Comparative analysis of cathepsin L, cathepsin D, and collagenase messenger RNA expression in synovial tissues of patients with rheumatoid arthritis and osteoarthritis, by in situ hybridization. Arthritis Rheum. 38, 976–984. 198. Schurigt, U., Stopfel, N., Huckel, M., Pfirschke, C., Wiederanders, B., and Brauer, R. (2005). Local expression of matrix metalloproteinases, cathepsins, and their inhibitors during the development of murine antigen-induced arthritis. Arthritis Res. Ther. 7, R174–188. 199. Taubert, H., Riemann, D., Kehlen, A., Meye, A., Bartel, F., John, V., Brandt, J., Bache, M., Wurl, P., Schmidt, H., and Weber, E. (2002). Expression of cathepsin B, D and L protein in juvenile idiopathic arthritis. Autoimmunity 35, 221–224. 200. Ikeda, Y., Ikata, T., Mishiro, T., Nakano, S., Ikebe, M., and Yasuoka, S. (2000). Cathepsins B and L in synovial fluids from patients with

Steven R. Goldring and Mary B. Goldring

201.

202.

203.

204.

205.

206. 207.

208.

209. 210.

211.

212.

213.

214.

215.

216.

rheumatoid arthritis and the effect of cathepsin B on the activation of pro-urokinase. J. Med. Invest. 47, 61–75. Hou, W. S., Li, Z., Gordon, R. E., Chan, K., Klein, M. J., Levy, R., Keysser, M., Keyszer, G., and Bromme, D. (2001). Cathepsin K is a critical protease in synovial fibroblast-mediated collagen degradation. Am. J. Pathol. 159, 2167–2177. Hou, W. S., Li, W., Keyszer, G., Weber, E., Levy, R., Klein, M. J., Gravallese, E. M., Goldring, S. R., and Bromme, D. (2002). Comparison of cathepsins K and S expression within the rheumatoid and osteoarthritic synovium. Arthritis Rheum. 46, 663–674. Garnero, P., Borel, O., Byrjalsen, I., Ferreras, M., Drake, F. H., McQueney, M. S., Foged, N. T., Delmas, P. D., and Delaisse, J. M. (1998). The collagenolytic activity of cathepsin K is unique among mammalian proteinases. J. Biol. Chem. 273, 32347–32352. Elsaid, K. A., Jay, G. D., and Chichester, C. O. (2003). Detection of collagen type II and proteoglycans in the synovial fluids of patients diagnosed with non-infectious knee joint synovitis indicates early damage to the articular cartilage matrix. Osteoarthritis Cartilage 11, 673–680. Kevorkian, L., Young, D. A., Darrah, C., Donell, S. T., Shepstone, L., Porter, S., Brockbank, S. M., Edwards, D. R., Parker, A. E., and Clark, I. M. (2004). Expression profiling of metalloproteinases and their inhibitors in cartilage. Arthritis Rheum. 50, 131–141. Poole, A. R., and Dieppe, P. (1994). Biological markers in rheumatoid arthritis. Sem. Arthritis Rheum. 23, 17–31. Saxne, T., and Heinegard, D. (1995). Matrix proteins: potentials as body fluid markers of changes in the metabolism of cartilage and bone in arthritis. J. Rheumatol. 43 (Suppl.), 71–74. Leroux, J. Y., Guerassimov, A., Cartman, A., Delaunay, N., Webber, C., Rosenberg, L. C., Banerjee, S., and Poole, A. R. (1996). Immunity to the G1 globular domain of the cartilage proteoglycan aggrecan can induce inflammatory erosive polyarthritis and spondylitis in BALB/c mice but immunity to G1 is inhibited by covalently bound keratan sulfate in vitro and in vivo. J. Clin. Invest. 97, 621–632. Kingsley, G., and Panayi, G. S. (1997). Joint destruction in rheumatoid arthritis: biological bases. Clin. Exp. Rheumatol. 15, S3–14. Cremer, M. A., Rosloniec, E. F., and Kang, A. H. (1998). The cartilage collagens: a review of their structure, organization, and role in the pathogenesis of experimental arthritis in animals and in human rheumatic disease. J. Molec. Med. 76, 275–288. Cheung, H. S., Ryan, L. M., Kozin, F., and McCarty, D. J. (1980). Identification of collagen subtypes in synovial fluid sediments from arthritic patients. Am. J. Med. 68, 73–79. Moreland, L. W., Stewart, T., Gay, R. E., Huang, G. Q., McGee, N., and Gay, S. (1989). Immunohistologic demonstration of type II collagen in synovial fluid phagocytes of osteoarthritis and rheumatoid arthritis patients. Arthritis Rheum. 32, 1458–1464. Saxne, T., Heinegard, D., and Wollheim, F. A. (1987). Cartilage proteoglycans in synovial fluid and serum in patients with inflammatory joint disease. Relation to systemic treatment. Arthritis Rheum. 30, 972–979. Witter, J., Roughley, P. J., Webber, C., Roberts, N., Keystone, E., and Poole, A. R. (1987). The immunologic detection and characterization of cartilage proteoglycan degradation products in synovial fluids of patients with arthritis. Arthritis Rheum. 30, 519–529. Thonar, E. J., Lenz, M. E., Klintworth, G. K., Caterson, B., Pachman, L. M., Glickman, P., Katz, R., Huff, J., and Kuettner, K. E. (1985). Quantification of keratan sulfate in blood as a marker of cartilage catabolism. Arthritis Rheum. 28, 1367–1376. Ratcliffe, A., Doherty, M., Maini, R. N., and Hardingham, T. E. (1988). Increased concentrations of proteoglycan components in the synovial fluids of patients with acute but not chronic joint disease. Ann. Rheum. Dis. 47, 826–832.

Chapter 51 Rheumatoid Arthritis and other Inflammatory Joint Pathologies 217. Lohmander, L. S., Dahlberg, L., Ryd, L., and Heinegard, D. (1989). Increased levels of proteoglycan fragments in knee joint fluid after injury. Arthritis Rheum. 32, 1434–1442. 218. Poole, A. R., Witter, J., Roberts, N., Piccolo, F., Brandt, R., Paquin, J., and Baron, M. (1990). Inflammation and cartilage metabolism in rheumatoid arthritis. Studies of the blood markers hyaluronic acid, orosomucoid, and keratan sulfate. Arthritis Rheum. 33, 790–799. 219. Silverman, B., Cawston, T. E., Page Thomas, D. P., Dingle, J. T., and Hazleman, B. L. (1990). The sulphated glycosaminoglycan levels in synovial fluid aspirates in patients with acute and chronic joint disease. Br. J. Rheumatol. 29, 340–344. 220. Guerassimov, A., Duffy, C., Zhang, Y., Banerjee, S., Leroux, J. Y., Reimann, A., Webber, C., Delaunay, N., Vipparti, V., Ronbeck, L., Cartman, A., Arsenault, L., Rosenberg, L. C., and Poole, A. R. (1997). Immunity to cartilage link protein in patients with juvenile rheumatoid arthritis. J. Rheumatol. 24, 959–964. 221. Guerassimov, A., Zhang, Y., Banerjee, S., Cartman, A., Leroux, J. Y., Rosenberg, L. C., Esdaile, J., Fitzcharles, M. A., and Poole, A. R. (1998). Cellular immunity to the G1 domain of cartilage proteoglycan aggrecan is enhanced in patients with rheumatoid arthritis but only after removal of keratan sulfate. Arthritis Rheum. 41, 1019–1025. 222. Rousseau, J. C., Zhu, Y., Miossec, P., Vignon, E., Sandell, L. J., Garnero, P., and Delmas, P. D. (2004). Serum levels of type IIA procollagen amino terminal propeptide (PIIANP) are decreased in patients with knee osteoarthritis and rheumatoid arthritis. Osteoarthritis Cartilage 12, 440–447. 223. Billinghurst, R. C., Dahlberg, L., Ionescu, M., Reiner, A., Bourne, R., Rorabeck, C., Mitchell, P., Hambor, J., Diekmann, O., Tschesche, H., Chen, J., Van Wart, H., and Poole, A. R. (1997). Enhanced cleavage of type II collagen by collagenases in osteoarthritic articular cartilage. J. Clin. Invest. 99, 1534–1545. 224. Hollander, A. P., Heathfield, T. F., Webber, C., Iwata, Y., Bourne, R., Rorabeck, C., and Poole, A. R. (1994). Increased damage to type II collagen in osteoarthritic articular cartilage detected by a new immunoassay. J. Clin. Invest. 93, 1722–1732. 225. Stoop, R., van der Kraan, P. M., Buma, P., Hollander, A. P., Poole, A. R., and van den Berg, W. B. (1999). Denaturation of type II collagen in articular cartilage in experimental murine arthritis. Evidence for collagen degradation in both reversible and irreversible cartilage damage. J. Pathol. 188, 329–337. 226. Song, X., Zeng, L., Jin, W., Thompson, J., Mizel, D. E., Lei, K., Billinghurst, R. C., Poole, A. R., and Wahl, S. M. (1999). Secretory leukocyte protease inhibitor suppresses the inflammation and joint damage of bacterial cell wall-induced arthritis. J. Exp. Med. 190, 535–542. 227. Ishiguro, N., Ito, T., Oguchi, T., Kojima, T., Iwata, H., Ionescu, M., and Poole, A. R. (2001). Relationships of matrix metalloproteinases and their inhibitors to cartilage proteoglycan and collagen turnover and inflammation as revealed by analyses of synovial fluids from patients with rheumatoid arthritis. Arthritis Rheum. 44, 2503–2511. 228. Poole, A. R., Ionescu, M., Fitzcharles, M. A., and Billinghurst, R. C. (2004). The assessment of cartilage degradation in vivo: development of an immunoassay for the measurement in body fluids of type II collagen cleaved by collagenases. J. Immunol. Methods. 294, 145–153. 229. Kim, T. H., Stone, M., Payne, U., Zhang, X., Ionescu, M., Lobanok, T., King, L., Poole, A. R., and Inman, R. D. (2005). Cartilage biomarkers in ankylosing spondylitis: Relationship to clinical variables and treatment response. Arthritis Rheum. 52, 885–891. 230. Seibel, M. J., Duncan, A., and Robins, S. P. (1989). Urinary hydroxypyridinium crosslinks provide indices of cartilage and bone involvement in arthritic diseases. J. Rheumatol. 16, 964–970. 231. Astbury, C., Bird, H. A., McLaren, A. M., and Robins, S. P. (1994). Urinary excretion of pyridium crosslinks of collagen correlated with joint damage in arthritis. Br. J. Rheumatol. 33, 11–15.

865

232. Hein, G., Franke, S., Muller, A., Braunig, E., Eidner, T., and Stein, G. (1997). The determination of pyridinium crosslinks in urine and serum as a possible marker of cartilage degradation in rheumatoid arthritis. Clin. Rheumatol. 16, 167–172. 233. Takahashi, M., Kushida, K., Hoshino, H., Suzuki, M., Sano, M., Miyamoto, S., and Inoue, T. (1996). Concentrations of pyridinoline and deoxypyridinoline in joint tissues from patients with osteoarthritis or rheumatoid arthritis. Ann. Rheum. Dis. 55, 324–327. 234. Greenwald, R. A. (1996). Monitoring collagen degradation in patients with arthritis. The search for suitable surrogates. Arthritis Rheum. 39, 1455–1465. 235. Ronday, H. K., Te Koppele, J. M., Greenwald, R. A., Moak, S. A., De Roos, J. A., Dijkmans, B. A., Breedveld, F. C., and Verheijen, J. H. (1998). Tranexamic acid, an inhibitor of plasminogen activation, reduces urinary collagen cross-link excretion in both experimental and rheumatoid arthritis. Br. J. Rheumatol. 37, 34–38. 236. Landewe, R., Geusens, P., Boers, M., van der Heijde, D., Lems, W., te Koppele, J., van der Linden, S., and Garnero, P. (2004). Markers for type II collagen breakdown predict the effect of disease-modifying treatment on long-term radiographic progression in patients with rheumatoid arthritis. Arthritis Rheum. 50, 1390–1399. 237. Saxne, T., Glennas, A., Kvien, T. K., Melby, K., and Heinegard, D. (1993). Release of cartilage macromolecules into the synovial fluid in patients with acute and prolonged phases of reactive arthritis. Arthritis Rheum. 36, 20–25. 238. Saxne, T., and Heinegard, D. (1992). Synovial fluid analysis of two groups of proteoglycan epitopes distinguishes early and late cartilage lesions. Arthritis Rheum. 35, 385–390. 239. Rizkalla, G., Reiner, A., Bogoch, E., and Poole, A. R. (1992). Studies of the articular cartilage proteoglycan aggrecan in health and osteoarthritis. Evidence for molecular heterogeneity and extensive molecular changes in disease. J. Clin. Invest. 90, 2268–2277. 240. Slater, R. R., Jr., Bayliss, M. T., Lachiewicz, P. F., Visco, D. M., and Caterson, B. (1995). Monoclonal antibodies that detect biochemical markers of arthritis in humans. Arthritis Rheum. 38, 655–659. 241. Sharif, M., Osborne, D. J., Meadows, K., Woodhouse, S. M., Colvin, E. M., Shepstone, L., and Dieppe, P. A. (1996). The relevance of chondroitin and keratan sulphate markers in normal and arthritic synovial fluid. Br. J. Rheumatol. 35, 951–957. 242. Bello, A. E., Garrett, W. E., Jr., Wang, H., Lohnes, J., DeLong, E., Caterson, B., and Kraus, V. B. (1997). Comparison of synovial fluid cartilage marker concentrations and chondral damage assessed arthroscopically in acute knee injury. Osteoarthritis Cartilage 5, 419–426. 243. Haraoui, B., Thonar, E. J., Martel-Pelletier, J., Goulet, J. R., Raynauld, J. P., Ouellet, M., and Pelletier, J. P. (1994). Serum keratan sulfate levels in rheumatoid arthritis: inverse correlation with radiographic staging. J. Rheumatol. 21, 813–817. 244. Shinmei, M., Miyauchi, S., Machida, A., and Miyazaki, K. (1992). Quantitation of chondroitin-4-sulfate and chondroitin-6-sulfate in pathological joint fluid. Arthritis Rheum. 35, 1304–1308. 245. Belcher, C., Yaqub, R., Fawthrop, F., Bayliss, M., and Doherty, M. (1997). Synovial fluid chondroitin and keratan sulphate epitopes, glycosaminoglycans, and hyaluronan in arthritic and normal knees. Ann. Rheum. Dis. 56, 299–307. 246. Manicourt, D. H., Poilvache, P., Van Egeren, A., Devogelaer, J. P., Lenz, M. E., and Thonar, E. J. (2000). Synovial fluid levels of tumor necrosis factor α and oncostatin M correlate with levels of markers of the degradation of crosslinked collagen and cartilage aggrecan in rheumatoid arthritis but not in osteoarthritis. Arthritis Rheum. 43, 281–288. 247. Lark, M. W., Bayne, E. K., and Lohmander, L. S. (1995). Aggrecan degradation in osteoarthritis and rheumatoid arthritis. Acta Orthopaed. Scand. Supplementum, 92–97.

866 248. Lark, M. W., Williams, H., Hoernner, L. A., Weidner, J., Ayala, J. M., Harper, C. F., Christen, A., Olszewski, J., Konteatis, Z., Webber, R., et al. (1995). Quantification of a matrix metalloproteinase-generated aggrecan G1 fragment using monospecific anti-peptide serum. Biochem. J. 307, 245–252. 249. Fosang, A. J., Last, K., Gardiner, P., Jackson, D. C., and Brown, L. (1995). Development of a cleavage-site-specific monoclonal antibody for detecting metalloproteinase-derived aggrecan fragments: detection of fragments in human synovial fluids. Biochem. J. 310, 337–343. 250. Hughes, C. E., Caterson, B., Fosang, A. J., Roughley, P. J., and Mort, J. S. (1995). Monoclonal antibodies that specifically recognize neoepitope sequences generated by ‘aggrecanase’ and matrix metalloproteinase cleavage of aggrecan: application to catabolism in situ and in vitro. Biochem. J. 305, 799–804. 251. Lark, M. W., Gordy, J. T., Weidner, J. R., Ayala, J., Kimura, J. H., Williams, H. R., Mumford, R. A., Flannery, C. R., Carlson, S. S., Iwata, M., et al. (1995). Cell-mediated catabolism of aggrecan. Evidence that cleavage at the “aggrecanase” site (Glu373-Ala374) is a primary event in proteolysis of the interglobular domain. J. Biol. Chem. 270, 2550–2556. 252. Fosang, A. J., Last, K., Neame, P. J., Murphy, G., Knauper, V., Tschesche, H., Hughes, C. E., Caterson, B., and Hardingham, T. E. (1994). Neutrophil collagenase (MMP-8) cleaves at the aggrecanase site E373-A374 in the interglobular domain of cartilage aggrecan. Biochem. J. 304, 347–351. 253. Buttner, F. H., Hughes, C. E., Margerie, D., Lichte, A., Tschesche, H., Caterson, B., and Bartnik, E. (1998). Membrane type 1 matrix metalloproteinase (MT1-MMP) cleaves the recombinant aggrecan substrate rAgg1mut at the ‘aggrecanase’ and the MMP sites. Characterization of MT1-MMP catabolic activities on the interglobular domain of aggrecan. Biochem. J. 333, 159–165. 254. Lark, M. W., Bayne, E. K., Flanagan, J., Harper, C. F., Hoerrner, L. A., Hutchinson, N. I., Singer, II, Donatelli, S. A., Weidner, J. R., Williams, H. R., Mumford, R. A., and Lohmander, L. S. (1997). Aggrecan degradation in human cartilage. Evidence for both matrix metalloproteinase and aggrecanase activity in normal, osteoarthritic, and rheumatoid joints. J. Clin. Invest. 100, 93–106. 255. Fosang, A. J., Last, K., and Maciewicz, R. A. (1996). Aggrecan is degraded by matrix metalloproteinases in human arthritis. Evidence that matrix metalloproteinase and aggrecanase activities can be independent. J. Clin. Invest. 98, 2292–2299. 256. Lohmander, L. S., Neame, P. J., and Sandy, J. D. (1993). The structure of aggrecan fragments in human synovial fluid. Evidence that aggrecanase mediates cartilage degradation in inflammatory joint disease, joint injury, and osteoarthritis. Arthritis Rheum. 36, 1214–1222. 257. Recklies, A. D., Baillargeon, L., and White, C. (1998). Regulation of cartilage oligomeric matrix protein synthesis in human synovial cells and articular chondrocytes. Arthritis Rheum. 41, 997–1006. 258. Dodge, G. R., Hawkins, D., Boesler, E., Sakai, L., and Jimenez, S. A. (1998). Production of cartilage oligomeric matrix protein (COMP) by cultured human dermal and synovial fibroblasts. Osteoarthritis Cartilage 6, 435–440. 259. Hummel, K. M., Neidhart, M., Vilim, V., Hauser, N., Aicher, W. K., Gay, R. E., Gay, S., and Hauselmann, H. J. (1998). Analysis of cartilage oligomeric matrix protein (COMP) in synovial fibroblasts and synovial fluids. Br. J. Rheumatol. 37, 721–728. 260. Neidhart, M., Zaucke, F., von Knoch, R., Jungel, A., Michel, B. A., Gay, R. E., and Gay, S. (2005). Galectin-3 is induced in rheumatoid arthritis synovial fibroblasts after adhesion to cartilage oligomeric matrix protein. Ann. Rheum. Dis. 64, 419–424. 261. Forslind, K., Eberhardt, K., Jonsson, A., and Saxne, T. (1992). Increased serum concentrations of cartilage oligomeric matrix protein. A prognostic marker in early rheumatoid arthritis. Br. J. Rheumatol. 31, 593–598.

Steven R. Goldring and Mary B. Goldring 262. Joosten, L. A., Helsen, M. M., Saxne, T., van De Loo, F. A., Heinegard, D., and van Den Berg, W. B. (1999). IL-1 α β blockade prevents cartilage and bone destruction in murine type II collageninduced arthritis, whereas TNF-α blockade only ameliorates joint inflammation. J. Immunol. 163, 5049–5055. 263. Roux-Lombard, P., Eberhardt, K., Saxne, T., Dayer, J. M., and Wollheim, F. A. (2001). Cytokines, metalloproteinases, their inhibitors and cartilage oligomeric matrix protein: relationship to radiological progression and inflammation in early rheumatoid arthritis. A prospective 5-year study. Rheumatology (Oxford) 40, 544–551. 264. Larsson, E., Erlandsson Harris, H., Lorentzen, J. C., Larsson, A., Mansson, B., Klareskog, L., and Saxne, T. (2002). Serum concentrations of cartilage oligomeric matrix protein, fibrinogen and hyaluronan distinguish inflammation and cartilage destruction in experimental arthritis in rats. Rheumatology (Oxford) 41, 996–1000. 265. Skoumal, M., Kolarz, G., and Klingler, A. (2003). Serum levels of cartilage oligomeric matrix protein. A predicting factor and a valuable parameter for disease management in rheumatoid arthritis. Scand. J. Rheumatol. 32, 156–161. 266. Soderlin, M. K., Kastbom, A., Kautiainen, H., Leirisalo-Repo, M., Strandberg, G., and Skogh, T. (2004). Antibodies against cyclic citrullinated peptide (CCP) and levels of cartilage oligomeric matrix protein (COMP) in very early arthritis: relation to diagnosis and disease activity. Scand. J. Rheumatol. 33, 185–188. 267. Crnkic, M., Mansson, B., Larsson, L., Geborek, P., Heinegard, D., and Saxne, T. (2003). Serum cartilage oligomeric matrix protein (COMP) decreases in rheumatoid arthritis patients treated with infliximab or etanercept. Arthritis Res. Ther. 5, R181–185. 268. Forsblad d’Elia, H., Christgau, S., Mattsson, L. A., Saxne, T., Ohlsson, C., Nordborg, E., and Carlsten, H. (2004). Hormone replacement therapy, calcium and vitamin D3 versus calcium and vitamin D3 alone decreases markers of cartilage and bone metabolism in rheumatoid arthritis: a randomized controlled trial [ISRCTN46523456]. Arthritis Res. Ther. 6, R457–468. 269. Hakala, B. E., White, C., and Recklies, A. D. (1993). Human cartilage gp-39, a major secretory product of articular chondrocytes and synovial cells, is a mammalian member of a chitinase protein family. J. Biol. Chem. 268, 25803–25810. 270. Johansen, J. S., Jensen, H. S., and Price, P. A. (1993). A new biochemical marker for joint injury. Analysis of YKL-40 in serum and synovial fluid. Br. J. Rheumatol. 32, 949–955. 271. Hu, B., Trinh, K., Figueira, W. F., and Price, P. A. (1996). Isolation and sequence of a novel human chondrocyte protein related to mammalian members of the chitinase protein family. J. Biol. Chem. 271, 19415–19420. 272. Johansen, J. S., Hvolris, J., Hansen, M., Backer, V., Lorenzen, I., and Price, P. A. (1996). Serum YKL-40 levels in healthy children and adults. Comparison with serum and synovial fluid levels of YKL-40 in patients with osteoarthritis or trauma of the knee joint. Br. J. Rheumatol. 35, 553–559. 273. Harvey, S., Whaley, J., and Eberhardt, K. (2000). The relationship between serum levels of YKL-40 and disease progression in patients with early rheumatoid arthritis. Scand. J. Rheumatol. 29, 391–393. 274. Volck, B., Johansen, J. S., Stoltenberg, M., Garbarsch, C., Price, P. A., Ostergaard, M., Ostergaard, K., Lovgreen-Nielsen, P., Sonne-Holm, S., and Lorenzen, I. (2001). Studies on YKL-40 in knee joints of patients with rheumatoid arthritis and osteoarthritis. Involvement of YKL-40 in the joint pathology. Osteoarthritis Cartilage 9, 203–214. 275. Vos, K., Steenbakkers, P., Miltenburg, A. M., Bos, E., van Den Heuvel, M. W., van Hogezand, R. A., de Vries, R. R., Breedveld, F. C., and Boots, A. M. (2000). Raised human cartilage glycoprotein-39 plasma levels in patients with rheumatoid arthritis and other inflammatory conditions. Ann. Rheum. Dis. 59, 544–548.

Chapter 51 Rheumatoid Arthritis and other Inflammatory Joint Pathologies 276. Matsumoto, T., and Tsurumoto, T. (2001). Serum YKL-40 levels in rheumatoid arthritis: correlations between clinical and laboratory parameters. Clin. Exp. Rheumatol. 19, 655–660. 277. Johansen, J. S., Stoltenberg, M., Hansen, M., Florescu, A., HorslevPetersen, K., Lorenzen, I., and Price, P. A. (1999). Serum YKL-40 concentrations in patients with rheumatoid arthritis: relation to disease activity. Rheumatology (Oxford) 38, 618–626. 278. Johansen, J. S., Kirwan, J. R., Price, P. A., and Sharif, M. (2001). Serum YKL-40 concentrations in patients with early rheumatoid arthritis: relation to joint destruction. Scand. J. Rheumatol. 30, 297–304. 279. den Broeder, A. A., Joosten, L. A., Saxne, T., Heinegard, D., Fenner, H., Miltenburg, A. M., Frasa, W. L., van Tits, L. J., Buurman, W. A., van Riel, P. L., van de Putte, L. B., and Barrera, P. (2002). Long term anti-tumour necrosis factor α monotherapy in rheumatoid arthritis: effect on radiological course and prognostic value of markers of cartilage turnover and endothelial activation. Ann. Rheum. Dis. 61, 311–318. 280. Baeten, D., Steenbakkers, P. G., Rijnders, A. M., Boots, A. M., Veys, E. M., and De Keyser, F. (2004). Detection of major histocompatibility complex/human cartilage gp-39 complexes in rheumatoid arthritis synovitis as a specific and independent histologic marker. Arthritis Rheum. 50, 444–451. 281. van Bilsen, J. H., van Dongen, H., Lard, L. R., van der Voort, E. I., Elferink, D. G., Bakker, A. M., Miltenburg, A. M., Huizinga, T. W., de Vries, R. R., and Toes, R. E. (2004). Functional regulatory immune responses against human cartilage glycoprotein-39 in health vs. proinflammatory responses in rheumatoid arthritis. Proc. Natl. Acad. Sci. USA 101, 17180–17185. 282. De Ceuninck, F., Gaufillier, S., Bonnaud, A., Sabatini, M., Lesur, C., and Pastoureau, P. (2001). YKL-40 (cartilage gp-39) induces proliferative events in cultured chondrocytes and synoviocytes and increases glycosaminoglycan synthesis in chondrocytes. Biochem. Biophys. Res. Commun. 285, 926–931. 283. Recklies, A. D., White, C., and Ling, H. (2002). The chitinase 3-like protein human cartilage glycoprotein 39 (HC-gp39) stimulates proliferation of human connective-tissue cells and activates both extracellular signal-regulated kinase- and protein kinase B-mediated signalling pathways. Biochem. J. 365, 119–126. 284. Ling, H., and Recklies, A. D. (2004). The chitinase 3-like protein human cartilage glycoprotein 39 inhibits cellular responses to the inflammatory cytokines interleukin-1 and tumour necrosis factor-α. Biochem. J. 380, 651–659. 285. Sekine, T., Masuko-Hongo, K., Matsui, T., Asahara, H., Takigawa, M., Nishioka, K., and Kato, T. (2001). Recognition of YKL-39, a human cartilage related protein, as a target antigen in patients with rheumatoid arthritis. Ann. Rheum. Dis. 60, 49–54. 286. Knorr, T., Obermayr, F., Bartnik, E., Zien, A., and Aigner, T. (2003). YKL-39 (chitinase 3-like protein 2), but not YKL-40 (chitinase 3-like protein 1), is up regulated in osteoarthritic chondrocytes. Ann. Rheum. Dis. 62, 995–998. 287. Saito, S., Kondo, S., Mishima, S., Ishiguro, N., Hasegawa, Y., Sandell, L. J., and Iwata, H. (2002). Analysis of cartilage-derived retinoic-acid-sensitive protein (CD-RAP) in synovial fluid from patients with osteoarthritis and rheumatoid arthritis. J. Bone Joint. Surg. Br. 84, 1066–1069. 288. Goldring, S. R. (2002). Bone and joint destruction in rheumatoid arthritis: what is really happening? J. Rheumatol. Suppl. 65, 44–48. 289. Goldring, S. R. (2003). Pathogenesis of bone and cartilage destruction in rheumatoid arthritis. Rheumatology (Oxford) 42 Suppl 2, ii11–6. 290. Goldring, S. R., and Goldring, M. B. (2004). The role of cytokines in cartilage matrix degeneration in osteoarthritis. Clin. Orthop. Relat. Res. S27–36. 291. Goldring, M. B., and Berenbaum, F. (2004). The regulation of chondrocyte function by proinflammatory mediators: prostaglandins and nitric oxide. Clin. Orthop. S37–46.

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292. Goldring, M. B., and Berenbaum, F. (1999). Human chondrocyte culture models for studying cyclooxygenase expression and prostaglandin regulation of collagen gene expression. Osteoarthritis Cartilage 7, 386–388. 293. van de Loo, F. A., and van den Berg, W. B. (2002). Gene therapy for rheumatoid arthritis. Lessons from animal models, including studies on interleukin-4, interleukin-10, and interleukin-1 receptor antagonist as potential disease modulators. Rheum. Dis. Clin. North Am. 28, 127–129. 294. Goldring, M. B., and Goldring, S. R. (2002). Role of cytokines and chemokines in cartilage and bone destruction in arthritis. Curr. Opin. Orthopaed. 13, 351–362. 295. Abramson, S. B., and Amin, A. (2002). Blocking the effects of IL-1 in rheumatoid arthritis protects bone and cartilage. Rheumatology (Oxford) 41, 972–980. 296. Dayer, J. M. (2003). The pivotal role of interleukin-1 in the clinical manifestations of rheumatoid arthritis. Rheumatology (Oxford) 42(Suppl. 2), ii3–ii10. 297. Dayer, J. M. (2004). The process of identifying and understanding cytokines: from basic studies to treating rheumatic diseases. Best Pract. Res. Clin. Rheumatol. 18, 31–45. 298. Kay, J., and Calabrese, L. (2004). The role of interleukin-1 in the pathogenesis of rheumatoid arthritis. Rheumatology (Oxford) 43 Suppl 3, iii2–iii9. 299. Lubberts, E., Koenders, M. I., and van den Berg, W. B. (2005). The role of T cell interleukin-17 in conducting destructive arthritis: lessons from animal models. Arthritis Res. Ther. 7, 29–37. 300. Eastgate, J. A., Symons, J. A., Wood, N. C., Grinlinton, F. M., di Giovine, F. S., and Duff, G. W. (1988). Correlation of plasma interleukin 1 levels with disease activity in rheumatoid arthritis. Lancet 2, 706–709. 301. Rooney, M., Symons, J. A., and Duff, G. W. (1990). Interleukin 1 β in synovial fluid is related to local disease activity in rheumatoid arthritis. Rheumatol. Int. 10, 217–219. 302. Kahle, P., Saal, J. G., Schaudt, K., Zacher, J., Fritz, P., and Pawelec, G. (1992). Determination of cytokines in synovial fluids: correlation with diagnosis and histomorphological characteristics of synovial tissue. Ann. Rheum. Dis. 51, 731–734. 303. Houssiau, F. A., Devogelaer, J. P., Van Damme, J., de Deuxchaisnes, C. N., and Van Snick, J. (1988). Interleukin-6 in synovial fluid and serum of patients with rheumatoid arthritis and other inflammatory arthritides. Arthritis Rheum. 31, 784–788. 304. Manicourt, D. H., Triki, R., Fukuda, K., Devogelaer, J. P., Nagant de Deuxchaisnes, C., and Thonar, E. J. (1993). Levels of circulating tumor necrosis factor α and interleukin-6 in patients with rheumatoid arthritis. Relationship to serum levels of hyaluronan and antigenic keratan sulfate. Arthritis Rheum. 36, 490–499. 305. Neidel, J., Schulze, M., and Lindschau, J. (1995). Association between degree of bone-erosion and synovial fluid-levels of tumor necrosis factor α in the knee-joints of patients with rheumatoid arthritis. Inflamm. Res. 44, 217–221. 306. Okamoto, H., Yamamura, M., Morita, Y., Harada, S., Makino, H., and Ota, Z. (1997). The synovial expression and serum levels of interleukin-6, interleukin-11, leukemia inhibitory factor, and oncostatin M in rheumatoid arthritis. Arthritis Rheum. 40, 1096–1105. 307. Hui, W., Bell, M., and Carroll, G. (1997). Detection of oncostatin M in synovial fluid from patients with rheumatoid arthritis. Ann. Rheum. Dis. 56, 184–187. 308. Brennan, F. M., Zachariae, C. O., Chantry, D., Larsen, C. G., Turner, M., Maini, R. N., Matsushima, K., and Feldmann, M. (1990). Detection of interleukin 8 biological activity in synovial fluids from patients with rheumatoid arthritis and production of interleukin 8 mRNA by isolated synovial cells. Eur. J. Immunol. 20, 2141–2144.

868 309. Koch, A. E., Kunkel, S. L., Harlow, L. A., Johnson, B., Evanoff, H. L., Haines, G. K., Burdick, M. D., Pope, R. M., and Strieter, R. M. (1992). Enhanced production of monocyte chemoattractant protein-1 in rheumatoid arthritis. J. Clin. Invest. 90, 772–779. 310. Ziolkowska, M., Koc, A., Luszczykiewicz, G., KsiezopolskaPietrzak, K., Klimczak, E., Chwalinska-Sadowska, H., and Maslinski, W. (2000). High levels of IL-17 in rheumatoid arthritis patients: IL-15 triggers in vitro IL-17 production via cyclosporin A-sensitive mechanism. J. Immunol. 164, 2832–2838. 311. McNearney, T., Baethge, B. A., Cao, S., Alam, R., Lisse, J. R., and Westlund, K. N. (2004). Excitatory amino acids, TNF-α, and chemokine levels in synovial fluids of patients with active arthropathies. Clin. Exp. Immunol. 137, 621–627. 312. Jouvenne, P., Fossiez, F., Banchereau, J., and Miossec, P. (1997). High levels of neutralizing autoantibodies against IL-1 α are associated with a better prognosis in chronic polyarthritis: a follow-up study. Scand. J. Immunol. 46, 413–418. 313. Graudal, N. A., Svenson, M., Tarp, U., Garred, P., Jurik, A. G., and Bendtzen, K. (2002). Autoantibodies against interleukin 1α in rheumatoid arthritis: association with long term radiographic outcome. Ann. Rheum. Dis. 61, 598–602. 314. Chikanza, I. C., Roux-Lombard, P., Dayer, J. M., and Panayi, G. S. (1995). Dysregulation of the in vivo production of interleukin-1 receptor antagonist in patients with rheumatoid arthritis. Pathogenetic implications. Arthritis Rheum. 38, 642–648. 315. van Roon, J. A. G., Lafeber, F. P. J. G., and Bijlsma, J. W. J. (2001). Synergistic activity of interleukin-4 and interleukin-10 in suppression of inflammation and joint destruction in rheumatoid arthritis. Arthritis Rheum. 44, 3–12. 316. Johansson, A. C., Hansson, A. S., Nandakumar, K. S., Backlund, J., and Holmdahl, R. (2001). IL-10-deficient B10.Q mice develop more severe collagen-induced arthritis, but are protected from arthritis induced with anti-type II collagen antibodies. J. Immunol. 167, 3505–3512. 317. Hata, H., Sakaguchi, N., Yoshitomi, H., Iwakura, Y., Sekikawa, K., Azuma, Y., Kanai, C., Moriizumi, E., Nomura, T., Nakamura, T., and Sakaguchi, S. (2004). Distinct contribution of IL-6, TNF-α, IL-1, and IL-10 to T cell-mediated spontaneous autoimmune arthritis in mice. J. Clin. Invest. 114, 582–588. 318. Butrimiene, I., Jarmalaite, S., Ranceva, J., Venalis, A., Jasiuleviciute, L., and Zvirbliene, A. (2004). Different cytokine profiles in patients with chronic and acute reactive arthritis. Rheumatology (Oxford) 43, 1300–1304. 319. Loeser, R. F. (2003). Systemic and local regulation of articular cartilage metabolism: where does leptin fit in the puzzle? Arthritis Rheum. 48, 3009–3012. 320. Popa, C., Netea, M. G., Radstake, T. R., van Riel, P. L., Barrera, P., and van der Meer, J. W. (2005). Markers of inflammation are negatively correlated with serum leptin in rheumatoid arthritis. Ann. Rheum. Dis. 64, 1195–1198. 321. Otero, M., Lago, R., Lago, F., Casanueva, F. F., Dieguez, C., GomezReino, J. J., and Gualillo, O. (2005). Leptin, from fat to inflammation: old questions and new insights. FEBS Lett. 579, 295–301. 322. Koch, A. E. (2005). Chemokines and their receptors in rheumatoid arthritis: Future targets? Arthritis Rheum. 52, 710–721. 323. Szekanecz, Z., Kim, J., and Koch, A. E. (2003). Chemokines and chemokine receptors in rheumatoid arthritis. Semin. Immunol. 15, 15–21. 324. Borzi, R. M., Mazzetti, I., Cattini, L., Uguccioni, M., Baggiolini, M., and Facchini, A. (2000). Human chondrocytes express functional chemokine receptors and release matrix-degrading enzymes in response to C-X-C and C-C chemokines. Arthritis Rheum. 43, 1734–1741. 325. Yuan, G. H., Masuko-Hongo, K., Sakata, M., Tsuruha, J., Onuma, H., Nakamura, H., Aoki, H., Kato, T., and Nishioka, K. (2001). The role of C-C chemokines and their receptors in osteoarthritis. Arthritis Rheum. 44, 1056–1070.

Steven R. Goldring and Mary B. Goldring 326. Alaaeddine, N., Olee, T., Hashimoto, S., Creighton-Achermann, L., and Lotz, M. (2001). Production of the chemokine RANTES by articular chondrocytes and role in cartilage degradation. Arthritis Rheum. 44, 1633–1643. 327. Kanbe, K., Takagishi, K., and Chen, Q. (2002). Stimulation of matrix metalloprotease 3 release from human chondrocytes by the interaction of stromal cell-derived factor 1 and CXC chemokine receptor 4. Arthritis Rheum. 46, 130–137. 328. Morel, J. C., Park, C. C., Woods, J. M., and Koch, A. E. (2001). A novel role for interleukin-18 in adhesion molecule induction through NF κ B and phosphatidylinositol (PI) 3-kinase-dependent signal transduction pathways. J. Biol. Chem. 276, 37069–37075. 329. Klimiuk, P. A., Sierakowski, S., Latosiewicz, R., Cylwik, J. P., Cylwik, B., Skowronski, J., and Chwiecko, J. (2002). Soluble adhesion molecules (ICAM-1, VCAM-1, and E-selectin) and vascular endothelial growth factor (VEGF) in patients with distinct variants of rheumatoid synovitis. Ann. Rheum. Dis. 61, 804–809. 330. Szekanecz, Z., and Koch, A. E. (2001). Update on synovitis. Curr. Rheumatol. Rep. 3, 53–63. 331. Fearon, U., Griosios, K., Fraser, A., Reece, R., Emery, P., Jones, P. F., and Veale, D. J. (2003). Angiopoietins, growth factors, and vascular morphology in early arthritis. J. Rheumatol. 30, 260–268. 332. Gudbjornsson, B., Christofferson, R., and Larsson, A. (2004). Synovial concentrations of the angiogenic peptides bFGF and VEGF do not discriminate rheumatoid arthritis from other forms of inflammatory arthritis. Scand. J. Clin. Lab. Invest. 64, 9–15. 333. Taylor, P. C. (2005). Serum vascular markers and vascular imaging in assessment of rheumatoid arthritis disease activity and response to therapy. Rheumatology (Oxford) 44, 721–728. 334. Knudson, W., Aguiar, D. J., Hua, Q., and Knudson, C. B. (1996). CD44-anchored hyaluronan-rich pericellular matrices: an ultrastructural and biochemical analysis. Exp. Cell Res. 228, 216–228. 335. Loeser, R. F., Carlson, C. S., and McGee, M. P. (1995). Expression of β1 integrins by cultured articular chondrocytes and in osteoarthritic cartilage. Exp. Cell Res. 217, 248–257. 336. Brennan, F. R., Mikecz, K., Glant, T. T., Jobanputra, P., Pinder, S., Bavington, C., Morrison, P., and Nuki, G. (1997). CD44 expression by leucocytes in rheumatoid arthritis and modulation by specific antibody: implications for lymphocyte adhesion to endothelial cells and synoviocytes in vitro. Scand. J. Immunol. 45, 213–220. 337. Koller, M., Aringer, M., Kiener, H., Erlacher, L., Machold, K., Eberl, G., Studnicka-Benke, A., Graninger, W., and Smolen, J. (1999). Expression of adhesion molecules on synovial fluid and peripheral blood monocytes in patients with inflammatory joint disease and osteoarthritis. Ann. Rheum. Dis. 58, 709–712. 338. Takagi, T., Okamoto, R., Suzuki, K., Hayashi, T., Sato, M., Kurosaka, N., and Koshino, T. (2001). Up-regulation of CD44 in rheumatoid chondrocytes. Scand. J. Rheumatol. 30, 110–113. 339. Naor, D., and Nedvetzki, S. (2003). CD44 in rheumatoid arthritis. Arthritis Res. Ther. 5, 105–115. 340. Yasuda, T., Poole, A. R., Shimizu, M., Nakagawa, T., Julovi, S. M., Tamamura, H., Fujii, N., and Nakamura, T. (2003). Involvement of CD44 in induction of matrix metalloproteinases by a COOH-terminal heparin-binding fragment of fibronectin in human articular cartilage in culture. Arthritis Rheum. 48, 1271–1280. 341. Yasuda, T., Kakinuma, T., Julovi, S. M., Yoshida, M., Hiramitsu, T., Akiyoshi, M., and Nakamura, T. (2004). COOH-terminal heparin-binding fibronectin fragment induces nitric oxide production in rheumatoid cartilage through CD44. Rheumatology (Oxford) 43, 1116–1120. 342. Mikecz, K., Dennis, K., Shi, M., and Kim, J. H. (1999). Modulation of hyaluronan receptor (CD44) function in vivo in a murine model of rheumatoid arthritis. Arthritis Rheum. 42, 659–668. 343. Neidhart, M., Gay, R. E., and Gay, S. (2000). Anti-interleukin-1 and anti-CD44 interventions producing significant inhibition of cartilage

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344.

345. 346.

347.

348.

349.

350.

351.

352.

353.

354.

355.

356.

357.

destruction in an in vitro model of cartilage invasion by rheumatoid arthritis synovial fibroblasts. Arthritis Rheum. 43, 1719–1728. Torsteinsdottir, I., Arvidson, N. G., Hallgren, R., and Hakansson, L. (1999). Monocyte activation in rheumatoid arthritis (RA): increased integrin, Fc γ and complement receptor expression and the effect of glucocorticoids. Clin. Exp. Immunol. 115, 554–560. Firestein, G. S. (2004). NF-κB: Holy Grail for rheumatoid arthritis? Arthritis Rheum. 50, 2381–2386. Vincenti, M. P., and Brinckerhoff, C. E. (2002). Transcriptional regulation of collagenase (MMP-1, MMP-13) genes in arthritis: integration of complex signaling pathways for the recruitment of genespecific transcription factors. Arthritis Res. 4, 157–164. Thomas, B., Berenbaum, F., Humbert, L., Bian, H., Béréziat, G., Crofford, L., and Olivier, J. L. (2000). Critical role of C/EBPδ and C/EBPβ factors in the stimulation of cyclooxygenase-2 gene transcription by interleukin-1β in articular chondrocytes. Eur. J. Biochem. 267, 1–13. Mengshol, J. A., Vincenti, M. P., and Brinckerhoff, C. E. (2001). IL-1 induces collagenase-3 (MMP-13) promoter activity in stably transfected chondrocytic cells: requirement for Runx-2 and activation by p38 MAPK and JNK pathways. Nucleic Acids Res. 29, 4361–4372. Catterall, J. B., Carrere, S., Koshy, P. J., Degnan, B. A., Shingleton, W. D., Brinckerhoff, C. E., Rutter, J., Cawston, T. E., and Rowan, A. D. (2001). Synergistic induction of matrix metalloproteinase 1 by interleukin-1α and oncostatin M in human chondrocytes involves signal transducer and activator of transcription and activator protein 1 transcription factors via a novel mechanism. Arthritis Rheum. 44, 2296–2310. Liacini, A., Sylvester, J., Li, W. Q., and Zafarullah, M. (2002). Inhibition of interleukin-1-stimulated MAP kinases, activating protein-1 (AP-1) and nuclear factor κ B (NF-κ B) transcription factors down-regulates matrix metalloproteinase gene expression in articular chondrocytes. Matrix Biol. 21, 251–262. Legendre, F., Dudhia, J., Pujol, J. P., and Bogdanowicz, P. (2003). JAK/STAT but not ERK1/ERK2 pathway mediates interleukin (IL)-6/soluble IL-6R down-regulation of Type II collagen, aggrecan core, and link protein transcription in articular chondrocytes. Association with a down-regulation of SOX9 expression. J. Biol. Chem. 278, 2903–2912. Handel, M. L., McMorrow, L. B., and Gravallese, E. M. (1995). Nuclear factor-κB in rheumatoid synovium. Localization of p50 and p65. Arthritis Rheum. 38, 1762–1770. Grall, F., Gu, X., Tan, L., Cho, J.-Y., Inan, M. S., Pettit, A., Thamrongsak, U., Choy, B. K., Manning, C., Akbarali, Y., Zerbini, L., Rudders, S., Goldring, S. R., Gravallese, E. M., Oettgen, P., Goldring, M. B., and Libermann, T. A. (2003). Responses to the proinflammatory cytokines interleukin-1 and tumor necrosis factor α in cells derived from rheumatoid synovium and other joint tissues involve NF κB-mediated induction of the Ets transcription factor ESE-1. Arthritis Rheum. 48, 1249–1260. Redlich, K., Kiener, H. P., Schett, G., Tohidast-Akrad, M., Selzer, E., Radda, I., Stummvoll, G. H., Steiner, C. W., Groger, M., Bitzan, P., Zenz, P., Smolen, J. S., and Steiner, G. (2001). Overexpression of transcription factor Ets-1 in rheumatoid arthritis synovial membrane: regulation of expression and activation by interleukin-1 and tumor necrosis factor α. Arthritis Rheum. 44, 266–274. Koller, M., Hayer, S., Redlich, K., Ricci, R., David, J. P., Steiner, G., Smolen, J. S., Wagner, E. F., and Schett, G. (2005). JNK1 is not essential for TNF-mediated joint disease. Arthritis Res. Ther. 7, R166–173. Okazaki, K., Li, J., Yu, H., Fukui, N., and Sandell, L. J. (2002). CCAAT/ Enhancer-binding proteins β and δ mediate the repression of gene transcription of cartilage-derived retinoic acid-sensitive protein induced by interleukin-1β. J. Biol. Chem. 277, 31526–31533. Tan, L., Peng, H., Osaki, M., Choy, B. K., Auron, P. E., Sandell, L. J., and Goldring, M. B. (2003). Egr-1 mediates transcriptional repression

358.

359.

360.

361.

362.

363.

364.

365.

366.

367. 368. 369.

370. 371.

372. 373. 374.

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of COL2A1 promoter activity by interleukin-1β. J. Biol. Chem. 278, 17688–17700. Imamura, T., Imamura, C., Iwamoto, Y., and Sandell, L. J. (2005). Transcriptional co-activator CBP/p300 increase chondrocyte Cd-rap gene expression by multiple mechanisms including sequestration of the repressor CCAAT/enhancer-binding protein. J. Biol. Chem. 280, 8343–8350. Hein, G. E., Kohler, M., Oelzner, P., Stein, G., and Franke, S. (2004). The advanced glycation end product pentosidine correlates to IL-6 and other relevant inflammatory markers in rheumatoid arthritis. Rheumatol Int. 26, 137–141. Foell, D., Kane, D., Bresnihan, B., Vogl, T., Nacken, W., Sorg, C., Fitzgerald, O., and Roth, J. (2003). Expression of the pro-inflammatory protein S100A12 (EN-RAGE) in rheumatoid and psoriatic arthritis. Rheumatology (Oxford) 42, 1383–1389. Martin, C. A., Carsons, S. E., Kowalewski, R., Bernstein, D., Valentino, M., and Santiago-Schwarz, F. (2003). Aberrant extracellular and dendritic cell (DC) surface expression of heat shock protein (hsp)70 in the rheumatoid joint: possible mechanisms of hsp/DCmediated cross-priming. J. Immunol. 171, 5736–5742. Drynda, S., Ringel, B., Kekow, M., Kuhne, C., Drynda, A., Glocker, M. O., Thiesen, H. J., and Kekow, J. (2004). Proteome analysis reveals disease-associated marker proteins to differentiate RA patients from other inflammatory joint diseases with the potential to monitor anti-TNFα therapy. Pathol. Res. Pract. 200, 165–171. Sinz, A., Bantscheff, M., Mikkat, S., Ringel, B., Drynda, S., Kekow, J., Thiesen, H. J., and Glocker, M. O. (2002). Mass spectrometric proteome analyses of synovial fluids and plasmas from patients suffering from rheumatoid arthritis and comparison to reactive arthritis or osteoarthritis. Electrophoresis 23, 3445–3456. Hattori, T., von der Mark, K., Kawaki, H., Yutani, Y., Kubota, S., Nakanishi, T., Eberspaecher, H., de Crombrugghe, B., and Takigawa, M. (2005). Downregulation of rheumatoid arthritis-related antigen RA-A47 (HSP47/colligin-2) in chondrocytic cell lines induces apoptosis and cell-surface expression of RA-A47 in association with CD9. J. Cell Physiol. 202, 191–204. Evans, C. H., Ghivizzani, S. C., Herndon, J. H., Wasko, M. C., Reinecke, J., Wehling, P., and Robbins, P. D. (2000). Clinical trials in the gene therapy of arthritis. Clin. Orthop. S300–307. Bresnihan, B. (2001). The safety and efficacy of interleukin-1 receptor antagonist in the treatment of rheumatoid arthritis. Semin. Arthritis Rheum. 30 (Suppl 2), 17–20. Feldmann, M., and Maini, R. N. (2001). Anti-TNF α therapy of rheumatoid arthritis: what have we learned? Annu. Rev. Immunol. 19, 163–196. Dayer, J. M., and Bresnihan, B. (2002). Targeting interleukin-1 in the treatment of rheumatoid arthritis. Arthritis Rheum. 46, 574–578. Mengshol, J. A., Mix, K. S., and Brinckerhoff, C. E. (2002). Matrix metalloproteinases as therapeutic targets in arthritic diseases: bull’seye or missing the mark? Arthritis Rheum. 46, 13-20. Goldring, M. B. (2001). Anticytokine therapy for osteoarthritis. Expert Opin. Biol. Ther. 1, 817–829. Chernajovsky, Y., Vessillier, S., Adams, G., Gofur, Y., Subang, M. C., Annenkov, A., Favorov, P., Daly, G., and Gould, D. (2003). Development of new molecules, vectors and cells for therapy of arthritis. Joint Bone Spine 70, 474–476. Vilcek, J., and Feldmann, M. (2004). Historical review: Cytokines as therapeutics and targets of therapeutics. Trends Pharmacol. Sci. 25, 201–209. Sweeney, S. E., and Firestein, G. S. (2004). Signal transduction in rheumatoid arthritis. Curr. Opin. Rheumatol. 16, 231–237. Ishikawa, T., Nishigaki, F., Miyata, S., Hirayama, Y., Minoura, K., Imanishi, J., Neya, M., Mizutani, T., Imamura, Y., Naritomi, Y., Murai, H., Ohkubo, Y., Kagayama, A., and Mutoh, S. (2005). Prevention of progressive joint destruction in collagen-induced arthritis in rats by a novel matrix metalloproteinase inhibitor, FR255031. Br. J. Pharmacol. 144, 133–143.

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Chapter 52

Osteoarthritis and Degenerative Spine Pathologies Kristina Åkesson, MD, PhD

I. II. III. IV. V. VI.

Associate Professor, Department of Orthopedics, Malmö University Hospital, 205 02 Malmö, Sweden

VII. VIII. IX. X. XI.

Introduction Characteristics of OA Etiology Treatment Options Current Diagnostic Procedures Biochemical Aspects of Osteoarthritis

I. INTRODUCTION

insights into bone and cartilage composition and metabolism, and thereby also into the pathogenesis of osteoarthritis and related degenerative joint diseases. In this regard, it is now apparent that the early biochemical changes in osteoarthritis include increased release of degradation products and increased production of reparative molecules from bone and cartilage matrix. Consequently, it is conceivable that the release of these products may be indicators of abnormality in bone and cartilage, and biochemical assays of these compounds may be used to detect and/or monitor development of joint disease, including osteoarthritis. There are currently a number of biochemical assays that can readily detect bone and cartilage matrix molecules in serum, urine and synovial fluid. Some of these biochemical markers may have the potential to serve as diagnostic aids, to evaluate disease progress and response to treatment in joint diseases. However, despite the rapid development in this area, most marker assays are still at the level of research,

Osteoarthritis is one of the most common diseases affecting the musculoskeletal system, causing pain, decreased mobility, and reduced quality of life for the individual patient. It is a heterogeneous and complex joint disease, which, in severe cases, may lead to progressive joint destruction and ultimately, to joint replacement surgery in major joints. The onset of the disease is usually insidious, occurring long before it can be detected with current clinical and diagnostic methods. Subsequently, permanent and irreversible damage of the joint have often occurred by the time the clinical diagnosis is made. Thus, it is highly desirable to develop sensitive and reliable methods for early diagnosis and prognostication of disease progress and to design appropriate therapies to avoid the need for joint replacement surgery. Advances in the knowledge of bone and cartilage biochemistry and physiology have provided important Dynamics of Bone and Cartilage Metabolism

Markers of Bone Turnover Markers of Cartilage Metabolism Spine Degeneration and Markers Summary Conclusions References

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which again reflects the complexity of the problem and the difficulties in defining osteoarthritis [1]. In this chapter I will discuss the potential utility of biochemical markers to evaluate osteoarthritis patients, while detailed description of each marker is presented in previous chapters. To facilitate the understanding of the rationale and the value of biochemical markers in osteoarthritic joint disease, a brief background of epidemiology, etiology, and currently available diagnostic methods is included.

II. CHARACTERISTICS OF OA Osteoarthritis may affect any of the peripheral joints, as well as the joints of the axial skeleton. The small joints of the hands, the knee, and hip joints are most frequently involved, while other joints such as the shoulders, elbows, wrists, and ankles are more likely to be spared, even in generalized osteoarthritis [2]. In spinal osteoarthritis, changes occur both in the intervertebral disk space, with disk degeneration, and in the small facet joints. Thus, osteoarthritic changes may be limited to one single major joint or be described as generalized, commonly including multiple small joints with or without engagement of large joints. The occurrence of osteoarthritis usually begins in the fifth decade of life with increasing prevalence and incidence with advancing age. Radiographic changes in the finger joints are evident in 10% of individuals by 40–49 years of age, while as many as 70–90% of women above 70 years of age show signs of finger joint osteoarthritis. Most surveys of hip osteoarthritis indicate a prevalence of 3–4% in populations above 55 years of age, with a similar distribution between men and women. Bilateral changes occur in about one-third of the cases. Thus, osteoarthritis of the hip is a major contributor to the 250 000 total joint replacements of the hip undertaken in the US each year. Osteoarthritis of the knee joint is more common with a reported prevalence of radiographic changes ranging from 4% to 30% in populations above 45 years of age and with an even further age-related increase above age 75 (40–60%) and with a predominance of women suffering [2]. Only limited information is available on the natural course of osteoarthritis. This relates to the inconsistent correlation between radiographic changes and clinical symptoms, causing difficulties in the assessment of disease progression and to the fact that arthritic pain is accepted as part of aging. However, it is well known that osteoarthritis may lead to either rapidly deteriorating joint integrity or be observed as a fairly stable condition. In osteoarthritis of the hip radiographic progression was seen in two-thirds of the cases over an 11-year period [3], while radiographic

progression in a study of knee osteoarthritis was seen in about 30% of the patients during a similar observation period [4].

III. ETIOLOGY Ample evidence suggests a multifactorial etiology of osteoarthritis, by combinations of biomechanical, biochemical, and genetic factors. However, the initial event that triggers the pathological process is unclear and it is still being debated whether the initial changes occur in the synovium, the cartilage, the subchondral bone or concurrently in all of these compartments. Even if it is assumed that the synovial involvement is secondary, one can not rule out the possibility that synovial secretion of substances, including resorptive cytokines, could by themselves influence the disease process [5]. Factors interfering with the biomechanical properties of the joint are risk factors for the development of osteoarthritis in weight-bearing joints, particularly the hip and knee. Congenital and developmental deformities, although relatively rare, confer a high risk of osteoarthritis at early age. Obesity, joint trauma, and certain repetitive loading patterns (often occupation-related) have also been shown to be associated with an increase in osteoarthritis [6, 7]. Pertaining to this, the knee joint is for mechanical reasons more vulnerable to injury than the hip, and instability or incongruity from ligamentous or meniscus injury is related to an increased incidence of osteoarthritis [8]. Hereditary factors also play a role for the development of osteoarthritis, especially generalized osteoarthritis. Familial forms have been identified, and in some instances are related to specific mutations in genes that regulate matrix production. Indeed, a mutation in the gene encoding type II pro-collagen (COL2A1) has been linked to the disease in rare familial types of osteoarthritis [9] (see Chapter 38). However, considering the heterogeneity of the disease, it seems highly unlikely that alteration in one or even several genes can solely account for the overall genetic influence on the development of osteoarthritis. From a biochemical viewpoint, it is generally accepted that the osteoarthritic process includes alterations in the normal balance between synthesis and degradation of the articular cartilage and/or the subchondral bone (see Chapters 25 and 26). On the other hand, it is less known as to whether the primary deficiency is inadequate cellular response to normal tissue demand or insufficient cellular response to supranormal demand from mechanical loading or injury. Nevertheless, the metabolic disturbance of bone and/or cartilage turnover leads to increased release of degradation products primarily into the synovial fluid,

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Chapter 52 Osteoarthritis and Degenerative Spine Pathologies

but also into serum (see Chapters 26, 27, and 31). The subsequent repair response could induce elevated levels of anabolic molecules, some of which may not be incorporated into the matrix, but released into the joint space (i.e., synovial fluid) or the circulation. Therefore, measurements of the levels of these degradation products and/or repair molecules may yield important information regarding the progression of the disease (see also Chapters 27 and 31). Past clinical observations suggest that the impairment of the normal coupling between degradation and repair in bone and cartilage can be either continuously progressive or halted at any time from process initiation. Accordingly, in some patients radiographically evident osteoarthritis may stabilize with only marginal changes over observational periods of 10-years or more, while in another patient with similar baseline changes, the disease may rapidly progress to end-stage joint destruction within 6–12 months. This implies that regulatory and disease-limiting mechanisms are as complex as the initiating forces, and may involve various inter- and intracellular signal transduction mediators and/or local regulators.

IV. TREATMENT OPTIONS Currently available treatment options for osteoarthritis are focusing on symptom relief; whereas truly diseasemodifying agents or methods are lacking. Thus, the basic therapy includes common analgesics, non-steroidal antiinflammatory drugs, physical therapy, walking aids, and eventually in severe cases joint replacement surgery. However, future research perspectives encompass strategies, from alteration of disease progression and thereby limitation of joint damage by pharmacological interventions of local or systemic agents, to cartilage transplants.

V. CURRENT DIAGNOSTIC PROCEDURES In order to evaluate the degree of joint affection a number of diagnostic tools are available, however, in most cases these require rather advanced disease state for a reliable diagnosis and identification. Aside from clinical assessment, the most widely used and accepted method relies on standardized radiographic assessment of the joint space and of the secondary periarticular changes in peripheral joints [10, 11]. Lower extremity joints are preferably evaluated on weight-bearing films to enhance correct staging and sequential evaluation. Relative insensitivity during early stages of disease and high interobserver variation on radiographic

film interpretation are, nevertheless, major limitations using radiography diagnostically. Furthermore, evaluation of degenerative changes of the spine imposes specific considerations, since there is no distinct correlation between radiographically observed degenerative changes and clinical symptoms. To overcome these limitations, scintimetric methods using technetium-99–labeled diphosphonate may be valuable tools at least for peripheral joints, as was shown by Dieppe et al. [12]. In this study, the investigators showed a good correlation between subchondral technetium retention and disease progression over 5 years in patients with established OA. Several new technologies for osteoarthritis detection, such as digitalization of X-rays and magnetic resonance imaging, are rapidly evolving. These methodologies are extensively computerized techniques, which allow for postexposure manipulation to enhance the evaluation of pathologies. These methods have the potential of becoming important auxiliary or even primary diagnostic methods identifying also early cartilage deficiencies, but they need further validation to confirm the clinical and pathological significance of identified aberration [13, 14]. One new method to visualize early changes in the cartilage is delayed gadolinium enhanced MRI (deGEMRIC). The negatively charged contrast agent Gd-DTPA2 distributes in the cartilage inversely to the likewise negatively charged glucosaminoglycan (GAG) content [15–17]. In patients with arthroscopically verified early osteoarthritis and acute anterior cruciate ligament injury, the loss of GAG could be identified as well as increased cartilage hydration [18, 19]. Most major peripheral joints are today accessible to arthroscopy, with the knee joint most commonly examined. Despite the fact that arthroscopy is not primarily a diagnostic tool, but rather a treatment procedure, this method provides a direct means to identify both major and minor intra-articular lesions and pathological changes. Even so, the significance of the findings for future development of osteoarthritis is only partially known and factors such as previous meniscectomy, severe instability, and intraarticular fracture are linked to osteoarthritis [8, 20].

VI. BIOCHEMICAL ASPECTS OF OSTEOARTHRITIS The biochemical alterations of osteoarthritis involve the dynamic processes of two completely different but adjacent and partially interconnected tissues: bone and cartilage. Throughout the body, bone is the predominant tissue, constituting 20–25% of the body weight, while cartilagecovered joint surfaces comprise less than a few percent.

874 The normal skeletal bone turnover rate is relatively slow and estimated to be an average of about 10% per year with a higher activity in trabecular bone (15–20%) and a lower activity in cortical bone (3–5%) [21]; no such estimation for cartilage turnover is available. In contrast to cartilage, bone tissue is highly vascularized, promoting rapid exchange of nutrients, ions, and other substances. This also renders the bone tissue susceptible to systemic regulator, including various hormones. Bone remodeling is activated by factors such as PTH, 1,25-dihydroxy vitamin D, growth factors, cytokines, mechanical load, or microinjuries [22, 23]. Cartilage is dependent on diffusion for nutrition and for release of waste products. Normal exchange requires an intact cartilage surface and pressure variation from joint loading. Thus, local factors, in addition to mechanical loading such as local enzyme and cytokine release, are probably of greater importance than systemic regulators for cartilage turnover. However, putative locally acting regulators such as growth factors, including insulin-like growth factors and cytokines, do not uniquely affect either bone or cartilage turnover but affect both, while certain enzymes and their inhibitors are tissue-specific (see also Chapters 25 and 26). Bone consists mainly of collagen type I; cartilage mainly contains collagen type II, but additional collagen types in various amounts (type VI, IX, X, and XI). The collagen fibrils are similarly synthesized but the final structural assembly differs, giving each tissue-specific properties of tensile strength and ability to withstand loading forces. In bone, structural solidity is provided by matrix mineralization from precipitation of hydroxyapatite, whereas in cartilage the firm elasticity results from large proteoglycans responsive to osmotic and loading pressures. In addition, a number of other collagenous and non-collagenous proteins are incorporated into bone and cartilage matrix (see also Chapters 25 and 26). Bone resorption relies on osteoclasts creating a suitable environment by locally increasing the acidity, and by secreting proteinases. Osteoblasts, on the other hand, are specifically related to bone formation or restoration, and produce a number of essential proteins, in addition to collagen. In cartilage, the chondrocytes are responsible for both the production of new matrix proteins and enzymes related to cartilage degradation. The particular cellular production and thereby the cartilage composition (type of collagen and proteoglycan content) appears to vary with the distance from the cartilage surface, where the surface layers are also affected by enzymes and cytokines produced by synovial cells [24]. Assuming that bone and cartilage turnover under normal circumstances remains at a fairly steady state, products released during tissue breakdown, bone resorption, and cartilage degradation could be used as markers for the

Kristina Åkesson

catabolic events. During the anabolic phase, osteoblasts and chondrocytes are actively producing collagens and other proteins to be incorporated in the newly synthesized matrix. Surplus products or fragments released into serum or synovial fluid may then be used as markers of bone and cartilage formation. In states of diseases, alterations of marker levels should provide insight into the metabolic disturbances in a particular bone and joint disorder. With regard to osteoarthritis, the ideal biochemical marker or markers should; (1) detect joint damage at an earlier stage than conventional methods; (2) provide information on disease activity and progressive joint damage; (3) predict future illness and course of disease, and (4) provide estimates of intervention efficacy (see also Chapter 31). None of the available biochemical markers for bone or cartilage are able to supply this information, due to lack of tissue specificity and sensitivity and, in view of the complexity of the osteoarthritic disease, clinically useful biochemical markers are not expected to be found in the immediate future. Nevertheless, recent research on biochemical markers has increased our knowledge of the underlying metabolic changes occurring during joint injury and during joint destruction in both osteoarthritis and rheumatoid arthritis. Most promising in a future perspective is the use of combinations of multiple markers to increase the disease specificity and thereby the clinical utility. In the following section, each one of the potentially useful markers for bone and for cartilage will be reviewed separately (Table I) as will markers related to spinal degenerative disease.

VII. MARKERS OF BONE TURNOVER The rationale for using markers primarily regarded as markers of bone formation and bone resorption is obviously related to the fact that bone and cartilage form an integrated tissue compartment, where processes involving the interface affect both tissues. The pronounced juxta-articular skeletal involvement in advanced osteoarthritis also indicates an early affection of bone tissue, while observations of structural differences (i.e. bone mass, geometry, cortical width) may suggest a general skeletal difference in this population. In addition, it is possible that the initial changes in the osteoarthritis process may emanate from the subchondral bone.

A. Osteocalcin Osteocalcin is one of the most abundant non-collagenous proteins of bone matrix, synthesized by the osteoblasts [23]. Osteocalcin is incorporated in the extracellular matrix and

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Chapter 52 Osteoarthritis and Degenerative Spine Pathologies Table I. Primary tissue source Bone

Cartilage

Cartilage and/or synovium

Synovium

Potential Biochemical Markers of Bone and Joint Tissue Turnover in Osteoarthritis Marker

Synthesis/degradation

Osteocalcin Bone-specific alkaline phosphatase Type I collagen C- and N-propeptide (PICP and PINP) Pyridinoline and cross-links Pyridinoline (PYD) Deoxypyridinoline (DPD) C-terminal and N-terminal cross-linking telopeptide (CTX-I and NTX-I) Tartrate-resistant acid phosphatase isoenzyme 5b Bone sialoprotein Aggrecan and fragments Core protein fragments Keratan sulfate Chondroitin sulfate Type II collagen C-propeptide (PIICP and PIINP) Type II collagen C-terminal cross-linking telopeptide (CTX-II) Cartilage oligomeric matrix protein (COMP) Matrix metalloproteinases and inhibitors Stromelysin (MMP-3) Interstitial collagenase (MMP-1) Tissue inhibitors of metalloproteinases (TIMP) YLK-40 Hyaluronic acid Type III collagen N-propeptide (PIIINP)

Assayed in

Synthesis Synthesis Synthesis

Serum + sf Serum Serum

Degradation Degradation Degradation Degradation Degradation

Urine + serum + sf Urine + serum + sf Serum + urine Serum Serum

Degradation Degradation Synthesis Synthesis Degradation Degradation

sf + serum sf + serum sf + serum sf + serum Serum + sf + urine sf + serum

Synthesis Synthesis Synthesis Synthesis Synthesis Synthesis

sf + serum sf + serum sf + serum Serum + sf Serum sf + serum

sf = synovial fluid.

is regarded as bone-specific, as the small amount present in dentin is negligible, but its precise function remains unknown. Serum osteocalcin is most often used to evaluate bone turnover in osteoporosis and regarded as a valid marker of bone formation [25, 26] (see Chapter 3). It has been suggested that there is an inverse relationship between osteoarthritis and osteoporosis, with a higher bone mass in patients with osteoarthritis [27]. Subsequently, if OA represents the antipode of osteoporosis, a protective effect could possibly be inferred and a shift in bone turnover anticipated. In a study of postmenopausal women with generalized osteoarthritis, osteocalcin was moderately but significantly increased compared with age- and sexmatched controls [28]. Even so, the increase was similar to that of postmenopausal women with osteoporosis, despite the higher bone mineral density in women with osteoarthritis [29]. In contrast to this, Garnero et al. [30] found osteocalcin decreased by 36% in patients with knee OA. Radiographically progressive knee OA over 3 years could on the other hand be predicted by the 1-year increase in osteocalcin, whereas change in bone markers did not correlate with subjective and functional changes (WOMAC) [31]. Baseline severity was not associated with osteocalcin levels, a finding similar to that in Japanese women with knee OA [32]. In order to test bone metabolic differences as a possible underlying cause of the increased bone mass in osteoarthritis, the responsiveness to 1,25-dihydroxyvitamin D3, a stimulator of osteocalcin synthesis and bone formation,

were explored. The stimulatory effect of short-term 1,25-dihydroxyvitamin D3 on osteocalcin production was, however, similar in OA and osteoporosis [29]. Still, quantitative and qualitative differences are observed and Raymaekers et al. [33] showed increased osteocalcin content in iliac crest bone from patients with osteoarthritis, corresponding to shift towards higher densities in the mineralization profile. Few studies have been conducted comparing serum osteocalcin and synovial fluid osteocalcin levels in knee osteoarthritis and with diverging results [34–36]. In a study by Sharif et al. [37] synovial fluid concentrations of osteocalcin correlated with abnormalities on scintigraphic scans and the osteocalcin level was related to scintigraphically assessed severity of joint affection (Fig. 1). This is the only study linking osteocalcin to osteoarthritis disease severity, suggesting that subchondral bone involvement may lead to an increased release of osteocalcin into the synovial fluid. In addition, synovial fluid osteocalcin was almost completely gammacarboxylated in the osteoarthritis patients compared with patients with rheumatoid arthritis [38].

B. Serum Alkaline Phosphatase The skeletal isoform of alkaline phosphatase is a membrane-bound protein with enzymatic activity (see also Chapter 9). Despite this, the release mechanism and the function remain to be elucidated. Studies using

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diagnostic specificity and sensitivity in osteoporosis [44], but also in monitoring treatment affecting bone turnover [45]. None of the markers have been evaluated in relation to osteoarthritis except in minor studies. In one study, PICP was not found to be associated with the radiographic severity score of end-stage knee OA [46] and in one study using PINP as an indicator of bone formation, PINP increased significantly more in those patients with osteoarthritis developing heterotopic ossification after total hip replacement, than in those who did not [47].

D. Pyridinoline Cross-links

Figure 1

Synovial fluid osteocalcin (OC) levels in osteoarthritis patients by category of results on scintigraphy scan. Median concentration indicated by bars. (Reprinted with permission from [85].)

combinations of monoclonal antibodies show a 52% increase in serum bone-specific alkaline phosphatase after menopause [39]. Alkaline phosphatase has only briefly been assessed in studies of osteoarthritis. In the only study found assessing biochemical markers in patients with spinal osteoarthritis, Peel et al. found a decrease in bone-specific alkaline phosphatase in women with spinal osteoarthritis compared with women without spinal osteoarthritis in a population-based setting. The decrease in bone-specific alkaline phosphatase was 19%. In this study, spinal osteoarthritis was associated with a generalized increase in BMD [40]. In contrast to this, the bone content of alkaline phosphatase was significantly elevated in femoral heads from women with osteoarthritis of the hip compared with controls [41].

C. Type I Collagen Carboxy- and Amino-propeptides During collagen type I synthesis by the osteoblasts, the amino- and carboxyterminal extension peptides are split off. The intact terminal fragments are released into the circulation prior to extracellular fibril formation [42] (see Chapters 1 and 2). Studies indicate that PICP may be used as a marker of bone formation, but with a questionable sensitivity in identifying subtle alterations of bone turnover [43], whereas evidence suggests PINP to have greater

Pyridinoline and deoxypyridinoline represent specific degradation products of mature collagen. The pyridinium crosslinks interconnect collagen molecules within the extracellular matrix; thus, pyridinium crosslinks are present in most connective tissues [24]. However, the reported distribution varies depending on tissue type: pyridinoline is primarily present in cartilage, but also in bone collagen, whereas deoxypyridinoline is predominant in bone and dentine [48]. Theoretically, deoxypyridinoline should therefore be the more suitable marker of bone resorption; it has indeed been shown to correlate positively with other indices of bone resorption, and inversely with bone mass in osteoporosis [39] (see also Chapters 2 and 35). The crosslinking telopeptide regions, C-telopeptide to helix (CTX-I) and N-telopeptide to helix (NTX-I), are currently considered the most valid markers of bone resorption, measurable either in serum or urine. Evaluation and use of these markers is focused on the assessment of osteoporosis and antiresorptive therapies [49–51] (see also Chapters 2, 35, and 40), however increasing evidence confirms a role also in the assessment of arthritis, particularly rheumatoid arthritis [52, 53]. In a study assessing the relative amounts of pyridinoline and deoxypyridinoline in subchondral bone, articular cartilage, and synovium, the relative content of PYR:DPD was 3:1 in bone, 25:1 in synovium and 50:1 in cartilage [54]. These ratios were similar to those of patients with rheumatoid arthritis [54]. The only significant difference was a higher cartilage content of deoxypyridinoline in osteoarthritis. Most studies report increased urinary concentrations of pyridinoline and deoxypyridinoline in both single joint or generalized osteoarthritis compared to reported excretion in healthy subjects [55–59], however reports with contradictory findings, i.e. no difference from controls regardless of whether the patient suffered from hip, knee, or generalized osteoarthritis exists [60] (Fig. 2). Nevertheless, patients judged to have severe or end-stage osteoarthritis had

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Chapter 52 Osteoarthritis and Degenerative Spine Pathologies

Assays for C-terminal or N-terminal cross-linking telopeptide are also markers of bone resorption [49, 62]. In the Clingford study, CTX-I and NTX-I were evaluated in 1003 postmenopausal women assessed for knee OA and followed for 4 years [63]. In women with radiographically progressive knee OA the levels of NTX-I were increased and at the same level as in those with osteoporosis, whereas CTX-I was not significantly higher. In a similar population of 67 patients CTX-I levels were lower (as were other markers of bone turnover) than in controls [30], furthermore CTX-I did not correlate with disease severity [30, 32]. Thus, although the subchondral and periarticular bone is clearly involved in osteoarthritic disease, evaluation of disease severity or disease progression through serum or urine markers of bone resorption does not give uniform information. Synovial concentrations of pyridinoline and deoxypyridinoline were measured in 20 patients with osteoarthritis and pyridinoline was about 30% of urinary concentrations and deoxypyridinoline was below the detection limit in all samples. This finding of pyridinoline, but not deoxypyridinoline in synovial fluid, may suggest that in cases of joint disease, pyridinoline excretion could also derive from collagen breakdown other than bone collagen, possibly cartilage collagen [64].

E. Tartrate-resistant Acid Phosphatase Figure 2

Urinary excretion of pyridinoline (PYD) and deoxypyridinoline (DPD) in osteoarthritis subgroups (knee, n = 31; hip, n = 47; generalized, n = 36). (Reprinted with permission from [26].)

higher excretion of pyridinolines compared to patients with early osteoarthritis, while the PYR/DPD ratio remained unchanged. It would be desirable to use biochemical markers as a severity index and several studies are assessing pyridinoline excretion in relation to joint destruction. Pyridinoline and deoxypyridinoline were identified as weak markers of disease severity according to X-ray staging of knee osteoarthritis in 60–68-year-old women [32, 58]. Contrary to this, Astbury et al. [59] found no difference in pyridinoline and deoxypyridinoline levels relative to the degree of joint involvement in osteoarthritis, but a significant correlation to disease severity in rheumatoid arthritis. In other studies, patients with knee OA and simultaneously verified hand OA, suggestive of generalized disease, have higher excretion of DPD or PYD compared to patients with only knee OA or to patients with only generalized osteoarthritis or only hip osteoarthritis [32, 61]. However, neither PYD nor DPD correlated with clinical or subjective (WOMAC) indices of disease [31, 59].

Tartrate-resistant acid phosphatase (TRAP) is secreted by osteoclasts and the amount reaching the circulation and a suggested marker of bone resorption. TRAP is an enzyme expressed in osteoclasts and other cells of the monocyte-macrophage lineage [65, 66] (see Chapter 10). TRAP exists in two isoforms 5a and 5b, where type 5b is specifically secreted by osteoclasts [67, 68] correlated with the bone resorption rate [69] and fracture [51]. In osteoarthritis the TRAP 5b and total TRAP protein levels are normal and in contrast to rheumatoid arthritis where total TRAP protein is increased and significantly correlated with disease activity as measured with CRP [70], a finding suggestive of a greater importance of TRAP 5a and macrophage activity.

F. Bone Sialoprotein Bone sialoprotein (BSP) is phosphorylated glycoprotein, containing sialic acid, with suggested cell-adhesive properties [71]. BSP is specific for bone and enriched in the immediate subchondral bone. Serum BSP is increased in states of high turnover and in a recent study, shown to

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correlate with bone resorption markers [72] (see Chapters 4 and 35). In symptomatic hip osteoarthritis serum BSP increased significantly during the first year by 63%. This increase was correlated with osteophyte formation, sclerosis, and radiographic severity index, but not with joint space narrowing assessed by digitized image analyzer, which was intended as the measure of disease progression [73]. In patients with knee joint abnormalities detectable on bone scan, BSP was elevated as compared to scannegative patients, however, the serum level was not correlated with the extent of abnormalities [74]. Neither was BSP a useful tool in predicting disease progress from baseline values, but progressive disease resulted in significantly increased levels in progressors at follow-up 3 years later [75]. The inconsistency of the findings is not surprising in view of the heterogeneity of the disease and subsequently the study populations include a certain disparity even if using specific selection criteria defining osteoarthritis. Because the disorder includes both single-joint disease and polyarticular involvement, it is more likely that general joint and skeletal affection leads to larger effect on the measured biochemical markers measured in serum and urine as these markers relate to changes in bone turnover of the entire skeleton.

VIII. MARKERS OF CARTILAGE METABOLISM Acute, and in many circumstances, chronic joint damage induce an increase in synovial fluid production. Thus, synovial fluid should have a high concentration of degradation products from cartilage break down, both after acute trauma to the joint surface and during degenerative processes (see also Chapter 27). Subsequent activation of repair mechanisms may lead to release of unincorporated matrix molecules into the synovial space. Assessment of synovial fluid concentrations of such molecular markers of cartilage turnover may provide a unique insight into the local joint process. In addition, because of systemic elimination of molecules related to cartilage degeneration and regeneration, these molecules are detectable in both serum and urine, but may then signify more general changes in body cartilage turnover.

A. Aggrecan Aggrecan is the major aggregating proteoglycan present in cartilage tissue. Attached to the core protein are glucosaminoglycan side chains containing keratan sulfate

or chondroitin sulfate. Aggrecan is attached to hyaluronic acid through a link protein. Degradation includes cleavage of the chondroitin and keratan sulfate-containing regions, as well as cleavage of the core protein at different sites and release of these fragments into body fluids. Several different epitopes are used to identify core protein fragments as well as the glucosaminoglycan side chains. A few studies also suggest that some of these epitopes, especially those related to chondroitin sulfate, may instead be indicators of aggrecan neosynthesis [76, 77] (see Chapters 5, 25, and 26).

B. Aggrecan Protein Fragments Aggrecan protein fragments have been assessed in several studies, most often evaluating knee osteoarthritis with the possibility of studying both synovial fluid and serum levels of aggrecan fragments. Knee trauma usually invokes an acute increase in synovial fluid. Thus, using a trauma model, Lohmander et al. [78] showed a rapid increase in proteoglycan fragment concentration in synovial fluid immediately after trauma. After this initial increase, the levels of proteoglycan fragments fell within a few months, however, a significant elevation persisted in many patients several years after injury. Additional observations report increased release of proteoglycan fragments in early stages of osteoarthritis without radiographic changes and an inverse relationship to osteoarthritis scores have been noted [79, 80] (Fig. 3).

C. Keratan Sulfate Keratan sulfate epitopes have been proposed as additional markers for cartilage damage and aggrecan degradation, while the findings are not always consistent and depend on the different epitopes used (see Chapters 25, 26, and 39). The earliest study by Thonar et al. [81], using monoclonal antibody specific for keratan sulfate, showed increased levels of keratan sulfate in 24 patients with osteoarthritis compared to normal adults. Further studies have confirmed the results and in a comprehensive study including 125 well-characterized patients with knee osteoarthritis, serum keratan sulfate levels were increased, despite a wide variation in the serum levels in the osteoarthritis population (men having higher serum levels than women) [82]. Synovial fluid keratan sulfate was measured in knee aspirates and increased over the serum levels in patients with osteoarthritis; however, there was no correlation between serum and synovial fluid concentrations. Serum keratan sulfate correlated weakly with the number of joints

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Chapter 52 Osteoarthritis and Degenerative Spine Pathologies

D. Chondroitin Sulfate

Figure 3

The distribution of proteoglycan fragments in joint fluid. (A) 16 healthy knees, (B) 123 cruciate ligament injury, (C) menicus tear or rupture, (D) 43 pyrophosphate arthritis, (E) 135 knee osteoarthritis. (Reprinted with permission from [13].)

involved, whereas there was no correlation with clinical disease measures. Elevated serum levels of keratan sulfate were also associated with familial OA and progressive disease [83]. In an interesting study, evaluating a nonweight bearing joint, the gleno-humeral joint, synovial keratan sulfate was elevated in joints with moderate and advanced osteoarthritis as classified during arthroscopy [84]. Further attempts to assess the keratan sulfate in relation to disease activity or severity have been made, however, no such association was apparent assessing synovial levels of keratan sulfate when using scintigraphic abnormalities as an activity measure in knee osteoarthritis [37] and decreased levels of keratan sulfate have also been reported [77, 85]. Polyarticular involvement as compared to single joint osteoarthritis may also be a sign of more severe disease and subsequently a greater release of keratan sulfate could be expected. Nevertheless, in a study involving 137 women, the serum keratan sulfate concentration was unrelated to joint involvement and in contrast to other studies, the levels were comparable to those in the control group [86]. Thus, the value of keratan sulfate as a diagnostic or severity index remains questionable. In order to investigate the prognostic value of keratan sulfate, serum keratan sulfate was measured at baseline and after 5 years in patients with knee osteoarthritis. However, at follow-up no difference was obvious in the baseline levels between those who were to progress radiographically after 5 years and those who remained stationary [87].

Chondroitin sulfate epitopes are present on the largest or intact aggrecan molecules. One of the more recently identified epitopes, 846, on chondroitin sulfate, has been suggested to indicate increased cartilage turnover and cartilage neosynthesis, while epitopes more proximal to the globular domain presumably relate to degradation. The serum levels of chondroitin sulfate 846 were increased by 19% in osteoarthritis patients, whereas the elevation was 56% in patients with rheumatoid arthritis [77]. This finding may indicate a more systemic joint involvement in rheumatoid arthritis. Conversely, the synovial levels were highest in the osteoarthritis patients, particularly in those with the longest disease duration and most pronounced joint destruction [77, 88]. Chondroitin sulfate epitope 3B3 is not present in normal articular cartilage and synovial fluid, but is released early in the osteoarthritis process or in a close temporal relationship to joint injury, as demonstrated in human knee synovial fluid [61, 62, 89, 90]. Chondroitin sulfate epitope 3B3 has also been used to evaluate osteoarthritis of the glenohumeral joint. Synovial fluid concentrations of epitope 3B3 distinguished normal from diseased joints (osteoarthritis grades II–IV), but did not separate the different stages [58, 84]. On the other hand, no association to disease activity as judged by scintimetric abnormalities was found in knee osteoarthritis, using an antibody identifying the same epitope (3B3) in synovial fluid [37]. The sulfatation pattern of chondroitin sulfate is age, sex, and disease dependent and thus chondroitin sulfate includes the unsaturated disaccharides ∆di-CS6 and ∆di-CS4, which are released into the synovial fluid [85]. In patients with osteoarthritis ∆di-CS6 is lower compared to normal, as is the relative proportion of ∆di-CS6/ ∆di-CS4 [91, 92]. However, analyzing the unsaturated forms require high-performance liquid chromatography, a method not suitable for routine use.

E. Cartilage Oligomeric Matrix Protein Cartilage oligomeric matrix protein (COMP) is a noncollagenous protein present in the extracellular matrix, with the highest amounts present in articular cartilage [93]. Despite the unknown function of this protein, it has earned interest as a potential marker of tissue damage, because it is highly specific for cartilage (see Chapter 5). In an initial study by Saxne and Heinegård [94], normal serum concentrations of COMP were found in patients with osteoarthritis. In this study it was also noted that COMP was unrelated to age. The comparison of synovial

880 fluid content did not include controls; instead it was found that the different types of arthritis had similar joint levels of COMP, with the exception of elevated levels in advanced rheumatoid arthritis. Controlled studies have now been undertaken and increased COMP levels in joint fluid have been found in early-stage knee osteoarthritis, as compared to patients with advanced changes and healthy controls [30, 79, 95]. In a later report by Vilim et al. [96] it was shown that synovitis was significantly contributing to the increase. During the 3-year follow-up period, patients with initial knee pain progressing to radiographically evident osteoarthritis showed increasing concentrations of serum COMP, while non-progressors did not [75], however, no such association was evident in another 3-year study evaluating the clinical end-points of pain, stiffness, WOMAC, and joint space width [31]. In order to further evaluate the usefulness of COMP to monitor disease progression, 57 patients with knee osteoarthritis were followed for 5 years. The change in serum COMP during the first year of follow-up was significantly larger in subsequent progressors compared to non-progressors [97] (Fig. 4). Moreover, in a report on hip osteoarthritis, serum COMP also appears to be a predictor of progressive joint damage in the hip joint [98].

Figure 4 Changes in serum cartilage oligomeric matrix protein levels over 1 year in patients with progressive and non-progressive knee osteoarthritis. (Reprinted with permission from [84].)

Kristina Åkesson

F. YKL-40 – Human Cartilage Glycoprotein 39 YKL-40 is a member of the family 18 glycosyl hydroxylases, abundant in synovium and cartilage but absent in bone, and with unknown physiological function [99–101]. Positive staining for YKL-40 is related to the degree of synovitis and increased levels of YKL-40 are seen in both serum and synovial fluid of rheumatoid arthritis patients, with a greater increase in synovial fluid and it is also elevated in patients with knee OA but to a lesser degree [102]. YKL-40 was also significantly elevated in patients with knee effusions compared to those without [30]. The number of studies is scare and YKL-40 is considered only for clinical investigations.

G. Type II Collagen Carboxy- and Amino-propeptide As previously mentioned, during collagen class I synthesis the procollagen extension peptides are split off and released into body fluids prior to helix formation. Measurements of procollagen terminals should then represent newly produced collagen, and thereby reflect collagen neosynthesis. Increased collagen type II production may be part of the normal repair process of minor injuries or be an expression of excessive reparative response, as is possible in early osteoarthritis. In support of this, several studies have shown increased levels of PIICP in the joint fluid of patients with knee osteoarthritis [103–105]. The synovial increase in PIICP was most pronounced in early and midstage osteoarthritis and to knee angulation, but decreased in final-stage osteoarthritis [106]. In addition, PIICP correlated positively with BMI, another independent risk factor for osteoarthritis. PIICP may also be predictive of early-stage cartilage degeneration, as was indicated in a prospective study of early-middle-age women, where the baseline PIICP level was associated with radiographic progression after 4 years [107]. Fragments of procollagen are also released into the circulation from the newly synthesized molecule and should be directly related to cartilage content. Nevertheless, PICP measured in serum was reduced in patients with knee OA, despite a known increased tissue content in OA [108]. Similarly, when instead using an assay based on the N-terminal (PIINP) Garnero et al. [109] showed that patients with knee OA had decreased serum levels of PIINP and by combining this marker of synthesis with urinary excretion of CTX-II, a degradation marker, an uncoupling index was developed; an increased baseline uncoupling index was associated with greater progression of joint damage assessed by radiography, arthroscopy, or VAS.

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H. Type II Collagen Carboxy- and Amino-Telopeptide Collagen type II is the most abundant matrix molecule of cartilage and is essential for structural cartilage integrity (see Chapter 25). Improvement of molecular markers by necessity should include assessment of collagen destruction. Fragments from collagen type II degradation are released into synovial fluid, serum, and urine and through recent advances several immunoassays have been developed. Using a monoclonal urinary immunoassay for collagen type II C-telopeptide (CTX-II) it was shown that patients with OA had a 1.53-fold increase in CTX-II, with an even greater increase in patients with rheumatoid arthritis [110]. CTX-II was also able to predict joint damage and WOMAC index and it is thereby potentially a marker of severity [30]. In synovial fluid, CTX-II concentrations have been determined in patients with knee injury to the anterior cruciate ligament or meniscus and in patients with knee osteoarthritis, with both of these groups having elevated levels compared to controls [111] (Fig. 5). The CTX-II level increased within hours after trauma and remained elevated for many years, which is in line with experimental data of early OA [112], whereas CTX-II did not differ between different radiographic stages of OA.

Figure 5

I. Matrix Metalloproteinases and their Inhibitors The matrix metalloproteinases (MMPs) consist of a family of 13 enzymes, active under both physiological and pathological conditions of cartilage turnover (see Chapter 11). The various enzymes, synthesized both by chondrocytes and synovial cells, are activated during different steps in collagen degradation, including degradation of aggrecan and with collagenase having the unique ability to cleave the triple-helical type II collagen [113]. Other members of the family are gelatinase and stromelysin. Specific tissue inhibitors of metalloproteinases, TIMPs, regulate the activity of these potent enzymes (see Chapter 11). Collagenase (MMP-1), stromelysin (MMP-3), and TIMP release have been evaluated in a variety of synovial fluid samples, representing normal individuals and patients with different types of knee injury, as well as primary osteoarthritis. The concentrations of metalloproteinases were elevated in all types on knee pathology, with stromelysin showing the differentiating ability. Moreover, a relatively higher increase in the concentration of MMP, compared to the enzyme inhibitors TIMP, is suggested to indicate a potential risk for further cartilage destruction in both knee and hip osteoarthritis [92, 114, 115]. A persistent

Synovial fluid (SF) concentrations of C-telopeptide fragments of type II collagen (CTX-II) in the different cross-sectional diagnostic groups. Study groups included healthy-knee volunteers (reference group [REF]), and those with acute pyrophosphate arthritis (PPA), anterior cruciate ligament ruptures or meniscus tears (INJ), and primary knee osteoarthritis (POA). Boxes represent the 25th and 75th percentiles and symbols are outliers. Note the y-axis break to accommodate single outlier value. (Reprinted with permission of Wiley-Liss, Inc., a subsidiary of John Wiley & Sons, Inc., from [111].)

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elevation of joint fluid levels of stromelysin and TIMP, for many years, is also seen after acute knee injury [1, 116]. Furthermore, subchondral bone tissue from subjects suffering from osteoarthritis of the hip shows increased content of metalloproteinases (gelatinase A/MMP-2) as compared to normal or osteoporotic bone, again indicating the interaction between cellular activity of bone and cartilage [41].

J. Hyaluronic Acid Hyaluronic acid, a linear polysaccharide, is synthesized by a number of cells, including synovial lining cells. Most inflammatory mediators, such as cytokines, prostaglandins, cAMP and growth factors, enhance the synthesis. Subsequently, situations activating these factors induce increased production of hyaluronic acid. Thus, increased synovial fluid and serum levels can be expected after joint trauma and with degenerative changes [30]. The information on hyaluronic acid in relation to osteoarthritis is limited. Two studies assessing hyaluronic acid in association with disease progress suggest that hyaluronic acid may prognosticate outcome in terms of joint damage and WOMAC index in knee osteoarthritis [31, 87].

IX. SPINE DEGENERATION AND MARKERS Spinal problems are very common in the population, in particular, low back pain, with a lifetime prevalence of 60–85%. Most back pain is non-specific with only 5–10% related to specific causes. Spinal disk degeneration and the associated spondylarthrosis is highly prevalent in the elderly and regarded as a form of osteoarthritis; spinal osteoarthritis (spinal OA). There is still little knowledge about the causal correlation between mechanical loading, degenerative spine disease and clinically significant back pain, whereas it is likely that morphological and biochemical changes in the intervertebral disk play crucial roles. Furthermore, there are inherent problems in objectively staging the degree of degeneration and the degree of spondylarthropathy using common imaging methods and subsequently to link it to biochemical alterations. The inferior and superior endplates of the vertebral bodies are cartilaginous and form the boundaries for the intervertebral disk with functional similarities to a joint. The cartilage endplates and the intervertebral disk are of similar biochemical composition with aggrecan as the main proteoglycan and keratan sulfate as the major glucosaminoglycan in the disk [117]. Biochemical changes in degenerative

spine disorders have mainly been studied through in vitro experiments using either disks from diseased persons or animal models and indicating a decrease of proteoglycans and in pyridinoline crosslinks and an increase in matrix metalloproteinases [118–123]. Keratan sulfate is another potential marker, because of the high disk content [124]. Bone markers have been widely used to evaluate clinical vertebral compression fractures, subclinical vertebral deformities in osteoporosis, and other bone diseases (see Chapters 40–53). However, only limited information exists on degenerative spine disorders and their effect on human bone turnover. Two studies specifically evaluating bone turnover in spinal OA are available with both studies using bone-specific alkaline phosphatase as a marker of bone formation and deoxypyridinoline as a marker of bone resorption and both markers were lower in patients with spinal OA [40, 125]. It is of equal interest to evaluate cartilage markers because of the extensive involvement of the disk, the endplate cartilage and also the facet joints, however information is scarce. In a study by Garnero et al. [126], 324 postmenopausal women, age 58–94, were evaluated for disk space narrowing and osteophytes as indicators of degenerative spine disorder and urinary C-terminal crosslinking telopeptide collagen type II excretion. It was found that CTX-II was increased by 34% in those with lumbar spine degeneration, but not in those with degenerative changes in the thoracic spine (Fig. 6). In those with lumbar disk space narrowing in combination with knee osteoarthritis and clinical hand osteoarthritis the increase was even more pronounced (80%) (Fig.7). These findings suggest that serum markers related to collagen type II and osteoarthritis reflect disease activity of all joints and have been suggested for aggrecan degradation products such as keratan sulfate [81, 82]. This interpretation is consistent with the widely accepted fact that bone turnover markers reflect the metabolic activity of the entire skeleton but with a relatively greater contribution from specific regions at times of localized high activity, for example during fracture healing. In patients undergoing decompressive laminectomy because of degenerative lumbar spine disease with claudication, urinary NTX-I was measured before and 12 months after surgery [127]. NTX-I levels decreased significantly after surgery but whether this is related to suppression of bone resorption from increased physical activity postoperatively or directly related to the removal of highly active bone remains unclear. Degenerative disk disorder may also be genetically determined as is indicated by the association of a collagen type I polymorphism in the first intron and a higher risk of disk degeneration (OR 3.6 (95% CI 1.3–10)) in elderly persons over 65 years [128].

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Figure 6

Association of lumbar and thoracic disk degeneration with type II collagen degradation as assessed by urinary CTX-II. Untreated postmenopausal women were categorized according to the summary grade for spine disk degeneration (range 0–2), which included both disk space narrowing and osteophytes. Bars show the mean and SD level of urinary CTX-II for patients with each grade of spinal disk degeneration. *P values derived from post hoc tests and adjusted for age and body mass index; NS = not significant. (Reprinted with permission of Wiley-Liss, Inc., a subsidiary of John Wiley & Sons, Inc., from [126].).

X. SUMMARY In view of the heterogeneity of OA, the inconsistency of results when evaluating the disease by various biochemical markers is not surprising. First of all, a certain disparity in the study populations is inevitable; despite the use of specific diagnostic or selection criteria, the distinction

Figure 7

between single-joint or multiple-joint involvement may not always be obvious. Even so, osteoarthritis of a major joint may be a sign of a general skeletal engagement and a systemic bone and joint disorder, which at least, could possibly influence the levels of biochemical markers of bone turnover. In this respect, it should be remembered that markers assayed in serum and urine reflect the entire turnover

Association of urinary levels of C-terminal crosslinking telopeptide of type II collagen (CTX-II) with radiologic lumbar spine disc degeneration, radiologic knee osteoarthritis (OA), and clinical hand OA in untreated postmenopausal women. Women were categorized according to the presence or absence of lumbar spine disc space narrowing (DSN) and/or radiologic knee OA and/or clinical hand OA. Bars show the mean and SD level of urinary CTX-II in each group.

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of bone, cartilage or other connective tissues, whereas synovial fluid measurements only provide information on the metabolic activity of the specific joint. It is also important to recognize that either measurement only gives us a glimpse of a dynamic process that we may inaccurately interpret as a steady state. Measurements of marker concentrations in joint fluid are attractive, since they provide information about the immediate metabolic activity of the affected joint. Synovial fluid measurements are almost exclusively performed on patients with knee osteoarthritis, since this joint is easily accessible and since knee joint damage often causes a marked fluid effusion. Thus, the knee joint is an excellent model for research, while the potential clinical utility of synovial fluid assessment from other joints to evaluate disease activity may be limited. In this regard, validated and reliable serum or urine markers are desirable. Biochemical markers of bone turnover have so far been tested in larger population-based settings and normative data are available, while such information to a large extent is lacking regarding cartilage markers. This may also be one of the limitations in the reported studies, as the control groups are often relatively small, and this is especially true for studies reporting on synovial fluid concentrations, thus comparisons are often made with rheumatoid arthritis. Given that one or several of the cartilage markers emerges as a clinically useful tool to monitor joint disease, progression or therapeutic interventions, it is necessary to include further studies providing normative information on the patient (age, gender, etc.), time (diurnal) and assay-related variability and this type of information is becoming increasingly available. Clearly, advances have been made and promising development involves markers related to collagen type II synthesis and degradation, while attempting to reach further using specific combinations of multiple markers may also prove useful [129].

XI. CONCLUSIONS Greater understanding of the underlying processes of bone and cartilage turnover in joint diseases has been gained through the assessment of biochemical marker molecules of bone and cartilage metabolism in synovial fluid, serum, and urine. Tissue-specific molecular markers provide a means to simultaneously study normal degradation and repair processes, as well as pathological states that are inaccessible by other methods. Despite the extensive work and the increased knowledge obtained from biochemical markers of skeletal and connective tissue origin, the search for more refined markers with the propensity of a wider clinical use must be continued.

References 1. Lohmander, L. S. (2004). Markers of altered metabolism in osteoarthritis. J. Rheumatol 31, 28–35. 2. Petersson, I. F. (1996). Occurrence of osteoarthritis of the peripheral joints in European populations. Ann. Rheum. Dis. 55, 659–661. 3. Danielsson, L. G. (1964). Incidence and prognosis of coxarthrosis. Acta. Orthop. Scand. 42, (SUPPL 66), 61–114. 4. Spector, T. D., Dacre, J. E., Harris, P. A., and Huskisson, E. C. (1992). Radiological progression of osteoarthritis: an 11 year follow up study of the knee. Ann. Rheum. Dis. 51, 1107–1110. 5. Westacott, C. I., Webb, G. R., Warnock, M. G., Sims, J. V., and Elson, C. J. (1997). Alteration of cartilage metabolism by cells from osteoarthritic bone. Arthritis Rheum. 40, 1282–1291. 6. Cooper, C., Campbell, L., Byng, P., Croft, P., and Coggon, D. (1996). Occupational activity and the risk of hip osteoarthritis. Ann. Rheum. Dis. 55, 680–682. 7. Vingard, E. (1996). Osteoarthrosis of the knee and physical load from occupation. Ann. Rheum. Dis. 55, 677–679. 8. Roos, H., Lindberg, H., Gardsell, P., Lohmander, L. S., and Wingstrand, H. (1994). The prevalence of gonarthrosis and its relation to meniscectomy in former soccer players. Am. J. Sports Med. 22, 219–222. 9. Vikkula, M., Palotie, A., Ritvaniemi, P., Ott, J., Ala-Kokko, L., Sievers, U., Aho, K., and Peltonen, L. (1993). Early-onset osteoarthritis linked to the type II procollagen gene. Detailed clinical phenotype and further analyses of the gene. Arthritis Rheum. 36, 401–409. 10. Kjellgren, J. H., Jeffrey, M., and Ball, J. (1963). Atlas of standard radiographs. The Epidemiology of Chronic Rheumatism. In: Scientific OB (ed). 11. Ahlback, S. (1968). Osteoarthrosis of the knee. A radiographic investigation. Acta. Radiol. Diagn. (Stockh):Suppl 277, 7–72. 12. Dieppe, P., Cushnaghan, J., Young, P., and Kirwan, J. (1993). Prediction of the progression of joint space narrowing in osteoarthritis of the knee by bone scintigraphy. Ann. Rheum. Dis. 52, 557–563. 13. McAlindon, T. E., Watt, I., McCrae, F., Goddard, P., and Dieppe, P. A. (1991). Magnetic resonance imaging in osteoarthritis of the knee: correlation with radiographic and scintigraphic findings. Ann. Rheum. Dis. 50, 14–19. 14. Blackburn, W. D., Jr., Chivers, S., and Bernreuter, W. (1996). Cartilage imaging in osteoarthritis. Semin. Arthritis. Rheum. 25, 273–281. 15. Recht, M. P., and Resnick, D. (1998). Magnetic resonance imaging of articular cartilage: an overview. Top Magn. Reson. Imaging. 9, 328–336. 16. Dieppe, P. A., Cushnaghan, J., and Shepstone, L. (1997). The Bristol ‘OA500’ study: progression of osteoarthritis (OA) over 3 years and the relationship between clinical and radiographic changes at the knee joint. Osteoarthritis Cartilage 5, 87–97. 17. Burstein, D., Velyvis, J., Scott, K. T., Stock, K. W., Kim, Y. J., Jaramillo, D., Boutin, R. D., and Gray, M. L. (2001). Protocol issues for delayed Gd(DTPA)(2-)-enhanced MRI (dGEMRIC) for clinical evaluation of articular cartilage. Magn. Reson. Med. 45, 36–41. 18. Tiderius, C. J., Olsson, L. E., Leander, P., Ekberg, O., and Dahlberg, L. (2003). Delayed gadolinium-enhanced MRI of cartilage (dGEMRIC) in early knee osteoarthritis. Magn. Reson. Med. 49, 488–492. 19. Tiderius, C. J., Svensson, J., Leander, P., Ola, T., and Dahlberg, L. (2004). dGEMRIC (delayed gadolinium-enhanced MRI of cartilage) indicates adaptive capacity of human knee cartilage. Magn. Reson. Med. 51, 286–290. 20. Lane, N. E. (1996). Physical activity at leisure and risk of osteoarthritis. Ann. Rheum. Dis. 55, 682–684. 21. Parfitt, A. M. (1993). The physiological and clinical significance of bone histomorphometric data. In: (R R. ed.), Bone Histomorphometry. Techniques and Interpretations. pp. 143–223. CRC Press, Boca Raton, FL.

Chapter 52 Osteoarthritis and Degenerative Spine Pathologies 22. Rodan, G. A., and Martin, T. J. (1981). Role of osteoblasts in hormonal control of bone resorption – a hypothesis. Calcif. Tissue. Int. 33, 349–351. 23. Price, P. A. (1985). Vitamin K-dependent formation of bone Gla protein (osteocalcin) and its function. Vitam. Horm. 42, 65–108. 24. Eyre, D. R. (1995). The specificity of collagen cross-links as markers of bone and connective tissue degradation. Acta. Orthop. Scand. Suppl. 266, 166–170. 25. Akesson, K. (1995). Biochemical markers of bone turnover. A review. Acta. Orthop. Scand. 66, 376–386. 26. Delmas, P. D. (1995). Biochemical markers of bone turnover. Acta. Orthop. Scand. Suppl. 266, 176–182. 27. Dequeker, J. (1985). The relationship between osteoporosis and osteoarthritis. Clin. Rheum. Dis. 11, 271–296. 28. Gevers, G., Dequeker, J., Geusens, P., Verstraeten, A., and Vanderschueren, D. (1988). Serum bone Gla protein (osteocalcin) and other markers of bone mineral metabolism in postmenopausal osteoporosis and osteoarthritis. J. Orthop. Rheum. 1, 22–27. 29. Geusens, P., Vanderschueren, D., Verstraeten, A., Dequeker, J., Devos, P., and Bouillon, R. (1991). Short-term course of 1,25(OH)2D3 stimulates osteoblasts but not osteoclasts in osteoporosis and osteoarthritis. Calcif. Tissue Int. 49, 168–173. 30. Garnero, P., Piperno, M., Gineyts, E., Christgau, S., Delmas, P. D., and Vignon, E. (2001). Cross sectional evaluation of biochemical markers of bone, cartilage, and synovial tissue metabolism in patients with knee osteoarthritis: relations with disease activity and joint damage. Ann. Rheum. Dis. 60, 619–626. 31. Bruyere, O., Collette, J. H., Ethgen, O., Rovati, L. C., Giacovelli, G., Henrotin, Y. E., Seidel, L., and Reginster, J. Y. (2003). Biochemical markers of bone and cartilage remodeling in prediction of longterm progression of knee osteoarthritis. J. Rheumatol. 30, 1043–1050. 32. Naitou, K., Kushida, K., Takahashi, M., Ohishi, T., and Inoue, T. (2000). Bone mineral density and bone turnover in patients with knee osteoarthritis compared with generalized osteoarthritis. Calcif. Tissue Int. 66, 325–329. 33. Raymaekers, G., Aerssens, J., Van den Eynde, R., Peeters, J., Geusens, P., Devos, P., and Dequeker, J. (1992). Alterations of the mineralization profile and osteocalcin concentrations in osteoarthritic cortical iliac crest bone. Calcif. Tissue Int. 51, 269–275. 34. Campion, G. V., Delmas, P. D., and Dieppe, P. A. (1989). Serum and synovial fluid osteocalcin (bone gla protein) levels in joint disease. Br. J. Rheumatol. 28, 393–398. 35. Poole, A. R., Ionescu, M., Swan, A., Campion, G. V., Rosenberg, L. C., and Dieppe, P. (1991). Analysis of cartilage and bone molecules and hyaluronic acid in synovial fluid and peripheral bloods of patients with arthritis. In: “Transactions of the combined meeting of the Orthopaedic Research Societies”. University of Calgary Printing Service, Banff, Alberta. 36. Mattei, J. P., Ferrera, V., Boutsen, Y., Franceschi, J. P., Botti, D., De Boissezon, C., Kaphan, G., and Roux, H. (1992). Serum and synovial fluid osteocalcin in rheumatic diseases. Int. J. Clin. Pharmacol. Res. 12, 103–107. 37. Sharif, M., George, E., and Dieppe, P. A. (1995). Correlation between synovial fluid markers of cartilage and bone turnover and scintigraphic scan abnormalities in osteoarthritis of the knee. Arthritis Rheum. 38, 78–81. 38. Fairney, A., Patel, K. V., Hollings, N. P., and Seifert, M. H. (1990). Abnormal osteocalcin binding in rheumatoid arthritis. Ann. Rheum. Dis. 49, 229–230. 39. Garnero, P., Sornay-Rendu, E., Chapuy, M. C., and Delmas, P. D. (1996). Increased bone turnover in late postmenopausal women is a major determinant of osteoporosis. J. Bone Miner. Res. 11, 337–349. 40. Peel, N. F., Barrington, N. A., Blumsohn, A., Colwell, A., Hannon, R., and Eastell, R. (1995). Bone mineral density and bone turnover in spinal osteoarthrosis. Ann. Rheum. Dis. 54, 867–871.

885 41. Mansell, J. P., Tarlton, J. F., and Bailey, A. J. (1997). Biochemical evidence for altered subchondral bone collagen metabolism in osteoarthritis of the hip. Br. J. Rheumatol. 36, 16–19. 42. Risteli, J., Niemi, S., Kauppila, S., Melkko, J., and Risteli, L. (1995). Collagen propeptides as indicators of collagen assembly. Acta. Orthop. Scand. Suppl. 266, 183–188. 43. Charles, P., Mosekilde, L., Risteli, L., Risteli, J., and Eriksen, E. F. (1994). Assessment of bone remodeling using biochemical indicators of type I collagen synthesis and degradation: relation to calcium kinetics. Bone Miner. 24, 81–94. 44. Scariano, J. K., Garry, P. J., Montoya, G. D., Duran-Valdez, E., and Baumgartner, R. N. (2002). Diagnostic efficacy of serum cross-linked N-telopeptide (NTx) and aminoterminal procollagen extension propeptide (PINP) measurements for identifying elderly women with decreased bone mineral density. Scand. J. Clin. Lab. Invest. 62, 237–243. 45. Tahtela, R., Seppanen, J., Laitinen, K., Katajamaki, A., Risteli, J., and Valimaki, M. J. (2005). Serum tartrate-resistant acid phosphatase 5b in monitoring bisphosphonate treatment with clodronate: a comparison with urinary N-terminal telopeptide of type I collagen and serum type I procollagen amino-terminal propeptide. Osteoporos. Int. 16, 1109–1116. 46. Schmidt-Rohlfing, B., Thomsen, M., Niedhart, C., Wirtz, D. C., and Schneider, U. (2002). Correlation of bone and cartilage markers in the synovial fluid with the degree of osteoarthritis. Rheumatol. Int. 21, 193–199. 47. Wilkinson, J. M., Stockley, I., Hamer, A. J., Barrington, N. A., and Eastell, R. (2003). Biochemical markers of bone turnover and development of heterotopic ossification after total hip arthroplasty. J. Orthop. Res. 21, 529–534. 48. Eyre, D. R. (1991). The collagens of articular cartilage. Semin. Arthritis Rheum. 21, 2–11. 49. Delmas, P. D., Eastell, R., Garnero, P., Seibel, M. J., and Stepan, J. (2000). The use of biochemical markers of bone turnover in osteoporosis. Committee of Scientific Advisors of the International Osteoporosis Foundation. Osteoporos. Int. 11, (Suppl 6) S2–17. 50. Garnero, P., Mulleman, D., Munoz, F., Sornay-Rendu, E., and Delmas, P. D. (2003). Long-term variability of markers of bone turnover in postmenopausal women and implications for their clinical use: the OFELY study. J. Bone Miner. Res. 18, 1789–1794. 51. Gerdhem, P., Ivaska, K. K., Alatalo, S. L., Halleen, J. M., Hellman, J., Isaksson, A., Pettersson, K., Vaananen, H. K., Akesson, K., and Obrant, K. J. (2004). Biochemical markers of bone metabolism and prediction of fracture in elderly women. J. Bone Miner. Res. 19, 386–393. 52. Garnero, P., Landewe, R., Boers, M., Verhoeven, A., Van Der Linden, S., Christgau, S., Van Der Heijde, D., Boonen, A., and Geusens, P. (2002). Association of baseline levels of markers of bone and cartilage degradation with long-term progression of joint damage in patients with early rheumatoid arthritis: the COBRA study. Arthritis Rheum. 46, 2847–2856. 53. Hermann, J., Mueller, T., Fahrleitner, A., and Dimai, H. P. (2003). Early onset and effective inhibition of bone resorption in patients with rheumatoid arthritis treated with the tumour necrosis factor alpha antibody infliximab. Clin. Exp. Rheumatol. 21, 473–476. 54. Takahashi, M., Kushida, K., Hoshino, H., Suzuki, M., Sano, M., Miyamoto, S., and Inoue, T. (1996). Concentrations of pyridinoline and deoxypyridinoline in joint tissues from patients with osteoarthritis or rheumatoid arthritis. Ann. Rheum. Dis. 55, 324–327. 55. Robins, S. P., Stewart, P., Astbury, C., and Bird, H. A. (1986). Measurement of the cross linking compound, pyridinoline, in urine as an index of collagen degradation in joint disease. Ann. Rheum. Dis. 45, 969–973. 56. Black, D., Duncan, A., and Robins, S. P. (1988). Quantitative analysis of the pyridinium crosslinks of collagen in urine using ion-paired reversed-phase high-performance liquid chromatography. Anal. Biochem. 169, 197–203.

886 57. Seibel, M. J., Duncan, A., and Robins, S. P. (1989). Urinary hydroxypyridinium crosslinks provide indices of cartilage and bone involvement in arthritic diseases. J. Rheumatol. 16, 964–970. 58. Thompson, P. W., Spector, T. D., James, I. T., Henderson, E., and Hart, D. J. (1992). Urinary collagen crosslinks reflect the radiographic severity of knee osteoarthritis. Br. J. Rheumatol. 31, 759–761. 59. Astbury, C., Bird, H. A., McLaren, A. M., and Robins, S. P. (1994). Urinary excretion of pyridinium crosslinks of collagen correlated with joint damage in arthritis. Br. J. Rheumatol. 33, 11–15. 60. Graverand, M. P., Tron, A. M., Ichou, M., Dallard, M. C., Richard, M., Uebelhart, D., and Vignon, E. (1996). Assessment of urinary hydroxypyridinium cross-links measurement in osteoarthritis. Br. J. Rheumatol. 35, 1091–1095. 61. MacDonald, A. G., McHenry, P., Robins, S. P., and Reid, D. M. (1994). Relationship of urinary pyridinium crosslinks to disease extent and activity in osteoarthritis. Br. J. Rheumatol. 33, 16–19. 62. Garnero, P., Rousseau, J. C., and Delmas, P. D. (2000). Molecular basis and clinical use of biochemical markers of bone, cartilage, and synovium in joint diseases. Arthritis Rheum. 43, 953–968. 63. Bettica, P., Cline, G., Hart, D. J., Meyer, J., and Spector, T. D. (2002). Evidence for increased bone resorption in patients with progressive knee osteoarthritis: longitudinal results from the Chingford study. Arthritis Rheum. 46, 3178–3184. 64. Sinigaglia, L., Varenna, M., Binelli, L., Bartucci, F., Arrigoni, M., Ferrara, R., and Abbiati, G. (1995). Urinary and synovial pyridinium crosslink concentrations in patients with rheumatoid arthritis and osteoarthritis. Ann. Rheum. Dis. 54, 144–147. 65. Yaziji, H., Janckila, A. J., Lear, S. C., Martin, A. W., and Yam, L. T. (1995). Immunohistochemical detection of tartrate-resistant acid phosphatase in non-hematopoietic human tissues. Am. J. Clin. Pathol. 104, 397–402. 66. Oddie, G. W., Schenk, G., Angel, N. Z., Walsh, N., Guddat, L. W., de Jersey, J., Cassady, A. I., Hamilton, S. E., and Hume, D. A. (2000). Structure, function, and regulation of tartrate-resistant acid phosphatase. Bone 27, 575–584. 67. Halleen, J. M., Alatalo, S. L., Suominen, H., Cheng, S., Janckila, A. J., and Vaananen, H. K. (2000). Tartrate-resistant acid phosphatase 5b: a novel serum marker of bone resorption. J. Bone Miner. Res. 15, 1337–1345. 68. Halleen, J. M., Ylipahkala, H., Alatalo, S. L., Janckila, A. J., Heikkinen, J. E., Suominen, H., Cheng, S., and Vaananen, H. K. (2002). Serum tartrate-resistant acid phosphatase 5b, but not 5a, correlates with other markers of bone turnover and bone mineral density. Calcif. Tissue Int. 71, 20–25. 69. Halleen, J. M. (2003). Tartrate-resistant acid phosphatase 5B is a specific and sensitive marker of bone resorption. Anticancer Res. 23, 1027–1029. 70. Janckila, A. J., Neustadt, D. H., Nakasato, Y. R., Halleen, J. M., Hentunen, T., and Yam, L. T. (2002). Serum tartrate-resistant acid phosphatase isoforms in rheumatoid arthritis. Clin. Chim. Acta. 320, 49–58. 71. Oldberg, A., Franzen, A., and Heinegard, D. (1988). The primary structure of a cell-binding bone sialoprotein. J. Biol. Chem. 263, 19430–19432. 72. Seibel, M. J., Woitge, H. W., Pecherstorfer, M., Karmatschek, M., Horn, E., Ludwig, H., Armbruster, F. P., and Ziegler, R. (1996). Serum immunoreactive bone sialoprotein as a new marker of bone turnover in metabolic and malignant bone disease. J. Clin. Endocrinol. Metab. 81, 3289–3294. 73. Conrozier, T., Shan, C., Saxen, T., Mathieu, P., Pipemo, M., and Vignon, E. (1997). Serum level of bone sialoprotein correlates with subchondral bone changes in hip osteoarthritis. Osteoarthritis Cartilage 5 (Suppl), 49. 74. Petersson, I. F., Boegard, T., Dahlstrom, J., Svensson, B., Heinegard, D., and Saxne, T. (1998). Bone scan and serum markers of bone and

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75.

76. 77.

78.

79.

80.

81.

82.

83.

84.

85.

86.

87.

88.

89.

90.

91.

92.

cartilage in patients with knee pain and osteoarthritis. Osteoarthritis Cartilage 6, 33–39. Petersson, I. F., Boegard, T., Svensson, B., Heinegard, D., and Saxne, T. (1998). Changes in cartilage and bone metabolism identified by serum markers in early osteoarthritis of the knee joint. Br. J. Rheumatol. 37, 46–50. Poole, A. R., and Dieppe, P. (1994). Biological markers in rheumatoid arthritis. Semin. Arthritis. Rheum. 23, 17–31. Poole, A. R., Ionescu, M., Swan, A., and Dieppe, P. A. (1994). Changes in cartilage metabolism in arthritis are reflected by altered serum and synovial fluid levels of the cartilage proteoglycan aggrecan. Implications for pathogenesis. J. Clin. Invest. 94, 25–33. Lohmander, L. S., Dahlberg, L., Ryd, L., and Heinegard, D. (1989). Increased levels of proteoglycan fragments in knee joint fluid after injury. Arthritis Rheum. 32, 1434–1442. Lohmander, L. S., Saxne, T., and Heinegard, D. K. (1994). Release of cartilage oligomeric matrix protein (COMP) into joint fluid after knee injury and in osteoarthritis. Ann. Rheum. Dis. 53, 8–13. Dahlberg, L., Ryd, L., Heinegard, D., and Lohmander, L. S. (1992). Proteoglycan fragments in joint fluid. Influence of arthrosis and inflammation. Acta. Orthop. Scand. 63, 417–423. Thonar, E. J., Lenz, M. E., Klintworth, G. K., Caterson, B., Pachman, L. M., Glickman, P., Katz, R., Huff, J., and Kuettner, K. E. (1985). Quantification of keratan sulfate in blood as a marker of cartilage catabolism. Arthritis Rheum. 28, 1367–1376. Campion, G. V., McCrae, F., Schnitzer, T. J., Lenz, M. E., Dieppe, P. A., and Thonar, E. J. (1991). Levels of keratan sulfate in the serum and synovial fluid of patients with osteoarthritis of the knee. Arthritis Rheum. 34, 1254–1259. Bleasel, J. F., Poole, A. R., Heinegard, D., Saxne, T., Holderbaum, D., Ionescu, M., Jones, P., and Moskowitz, R. W. (1999). Changes in serum cartilage marker levels indicate altered cartilage metabolism in families with the osteoarthritis-related type II collagen gene COL2A1 mutation. Arthritis Rheum. 42, 39–45. Ratcliffe, A., Flatow, E. L., Roth, N., Saed-Nejad, F., and Bigliani, L. U. (1996). Biochemical markers in synovial fluid identify early osteoarthritis of the glenohumeral joint. Clin. Orthop. 330, 45–53. Sharif, M., Osborne, D. J., Meadows, K., Woodhouse, S. M., Colvin, E. M., Shepstone, L., and Dieppe, P. A. (1996). The relevance of chondroitin and keratan sulphate markers in normal and arthritic synovial fluid. Br. J. Rheumatol. 35, 951–957. Spector, T. D., Woodward, L., Hall, G. M., Hammond, A., Williams, A., Butler, M. G., James, I. T., Hart, D. J., Thompson, P. W., and Scott, D. L. (1992). Keratan sulphate in rheumatoid arthritis, osteoarthritis, and inflammatory diseases. Ann. Rheum. Dis. 51, 1134–1137. Sharif, M., George, E., Shepstone, L., Knudson, W., Thonar, E. J., Cushnaghan, J., and Dieppe, P. (1995). Serum hyaluronic acid level as a predictor of disease progression in osteoarthritis of the knee. Arthritis Rheum. 38, 760–767. Lohmander, L. S., Ionescu, M., Jugessur, H., Poole, A. R. (1999). Changes in joint cartilage aggrecan after knee injury and in osteoarthritis. Arthritis Rheum. 42, 534–544. Slater, R. R., Jr., Bayliss, M. T., Lachiewicz, P. F., Visco, D. M., and Caterson, B. (1995). Monoclonal antibodies that detect biochemical markers of arthritis in humans. Arthritis Rheum. 38, 655–659. Ratcliffe, A., Grelsamer, R. P., Kiernan, H., Saed-Nejad, F., and Visco, D. (1995). High levels of aggrecan aggregate components are present in synovial fluids from human knee joints with chronic injury or osteoarthrosis. Acta. Orthop. Scand. Suppl. 266, 111–115. Lewis, S., Crossman, M., Flannelly, J., Belcher, C., Doherty, M., Bayliss, M. T., and Mason, R. M. (1999). Chondroitin sulphation patterns in synovial fluid in osteoarthritis subsets. Ann. Rheum. Dis. 58, 441–445. Ishiguro, N., Ito, T., Ito, H., Iwata, H., Jugessur, H., Ionescu, M., and Poole, A. R. (1999). Relationship of matrix metalloproteinases and

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Chapter 52 Osteoarthritis and Degenerative Spine Pathologies

93.

94.

95.

96.

97.

98.

99.

100.

101.

102.

103.

104.

105.

106.

107.

their inhibitors to cartilage proteoglycan and collagen turnover: analyses of synovial fluid from patients with osteoarthritis. Arthritis Rheum. 42, 129–136. Hedbom, E., Antonsson, P., Hjerpe, A., Aeschlimann, D., Paulsson, M., Rosa-Pimentel, E., Sommarin, Y., Wendel, M., Oldberg, A., and Heinegard, D. (1992). Cartilage matrix proteins. An acidic oligomeric protein (COMP) detected only in cartilage. J. Biol. Chem. 267, 6132–6136. Saxne, T., and Heinegard, D. (1992). Cartilage oligomeric matrix protein: a novel marker of cartilage turnover detectable in synovial fluid and blood. Br. J. Rheumatol. 31, 583–591. Petersson, I. F., Sandqvist, L., Svensson, B., and Saxne, T. (1997). Cartilage markers in synovial fluid in symptomatic knee osteoarthritis. Ann. Rheum. Dis. 56, 64–67. Vilim, V., Vytasek, R., Olejarova, M., Machacek, S., Gatterova, J., Prochazka, B., Kraus, V. B., and Pavelka, K. (2001). Serum cartilage oligomeric matrix protein reflects the presence of clinically diagnosed synovitis in patients with knee osteoarthritis. Osteoarthritis Cartilage 9, 612–618. Sharif, M., Saxne, T., Shepstone, L., Kirwan, J. R., Elson, C. J., Heinegard, D., and Dieppe, P. A. (1995). Relationship between serum cartilage oligomeric matrix protein levels and disease progression in osteoarthritis of the knee joint. Br. J. Rheumatol. 34, 306–310. Conrozier, T., Saxne, T., Fan, C. S., Mathieu, P., Tron, A. M., Heinegard, D., and Vignon, E. (1998). Serum concentrations of cartilage oligomeric matrix protein and bone sialoprotein in hip osteoarthritis: a one year prospective study. Ann. Rheum. Dis. 57, 527–532. Hakala, B. E., White, C., and Recklies, A. D. (1993). Human cartilage gp-39, a major secretory product of articular chondrocytes and synovial cells, is a mammalian member of a chitinase protein family. J. Biol. Chem. 268, 25803–25810. Johansen, J. S., Jensen, H. S., and Price, P. A. (1993). A new biochemical marker for joint injury. Analysis of YKL-40 in serum and synovial fluid. Br. J. Rheumatol. 32, 949–955. Johansen, J. S., Hvolris, J., Hansen, M., Backer, V., Lorenzen, I., and Price, P. A. (1996). Serum YKL-40 levels in healthy children and adults. Comparison with serum and synovial fluid levels of YKL-40 in patients with osteoarthritis or trauma of the knee joint. Br. J. Rheumatol. 35, 553–559. Volck, B., Johansen, J. S., Stoltenberg, M., Garbarsch, C., Price, P. A., Ostergaard, M., Ostergaard, K., Lovgreen-Nielsen, P., Sonne-Holm, S., and Lorenzen, I. (2001). Studies on YKL-40 in knee joints of patients with rheumatoid arthritis and osteoarthritis. Involvement of YKL-40 in the joint pathology. Osteoarthritis Cartilage 9, 203–214. Shinmei, M., Kobayashi, T., Yoshihara, Y., and Samura, A. (1995). Significance of the levels of carboxy terminal type II procollagen peptide, chondroitin sulfate isomers, tissue inhibitor of metalloproteinases, and metalloproteinases in osteoarthritis joint fluid. J. Rheumatol. Suppl. 43, 78–81. Lohmander, L. S., Yoshihara, Y., Roos, H., Kobayashi, T., Yamada, H., and Shinmei, M. (1996). Procollagen II C-propeptide in joint fluid: changes in concentration with age, time after knee injury, and osteoarthritis. J. Rheumatol. 23, 1765–1769. Kobayashi, T., Yoshihara, Y., Samura, A., Yamada, H., Shinmei, M., Roos, H., and Lohmander, L. S. (1997). Synovial fluid concentrations of the C-propeptide of type II collagen correlate with body mass index in primary knee osteoarthritis. Ann. Rheum. Dis. 56, 500–503. Kobayashi, T., Yoshihara, Y., Yamada, H., and Fujikawa, K. (2000). Procollagen IIC-peptide as a marker for assessing mechanical risk factors of knee osteoarthritis: effect of obesity and varus alignment. Ann. Rheum. Dis. 59, 982–984. Sugiyama, S., Itokazu, M., Suzuki, Y., and Shimizu, K. (2003). Procollagen II C propeptide level in the synovial fluid as a predictor

108.

109.

110.

111.

112.

113. 114.

115.

116.

117.

118. 119. 120.

121.

122.

123.

124.

of radiographic progression in early knee osteoarthritis. Ann. Rheum. Dis. 62, 27–32. Nelson, F., Dahlberg, L., Laverty, S., Reiner, A., Pidoux, I., Ionescu, M., Fraser, G. L., Brooks, E., Tanzer, M., Rosenberg, L. C., Dieppe, P., and Poole, A. (1998). Evidence for altered synthesis of type II collagen in patients with osteoarthritis. J. Clin. Invest. 102, 2115–2125. Garnero, P., Ayral, X., Rousseau, J. C., Christgau, S., Sandell, L. J., Dougados, M., and Delmas, P. D. (2002). Uncoupling of type II collagen synthesis and degradation predicts progression of joint damage in patients with knee osteoarthritis. Arthritis Rheum. 46, 2613–2624. Christgau, S., Garnero, P., Fledelius, C., Moniz, C., Ensig, M., Gineyts, E., Rosenquist, C., and Qvist, P. (2001). Collagen type II C-telopeptide fragments as an index of cartilage degradation. Bone 29, 209–215. Lohmander, L. S., Atley, L. M., Pietka, T. A., and Eyre, D. R. (2003). The release of crosslinked peptides from type II collagen into human synovial fluid is increased soon after joint injury and in osteoarthritis. Arthritis Rheum. 48, 3130–3139. Matyas, J. R., Atley, L., Ionescu, M., Eyre, D. R., and Poole, A. R. (2004). Analysis of cartilage biomarkers in the early phases of canine experimental osteoarthritis. Arthritis Rheum. 50, 543–552. Murphy, G. (1995). Matrix metalloproteinases and their inhibitors. Acta. Orthop. Scand. Suppl. 266, 55–60. Lohmander, L. S., Hoerrner, L. A., and Lark, M. W. (1993). Metalloproteinases, tissue inhibitor, and proteoglycan fragments in knee synovial fluid in human osteoarthritis. Arthritis Rheum. 36, 181–189. Chevalier, X., Conrozier, T., Gehrmann, M., Claudepierre, P., Mathieu, P., Unger, S., and Vignon, E. (2001). Tissue inhibitor of metalloprotease-1 (TIMP-1) serum level may predict progression of hip osteoarthritis. Osteoarthritis Cartilage 9, 300–307. Dahlberg, L., Friden, T., Roos, H., Lark, M.W., and Lohmander, L. S. (1994). A longitudinal study of cartilage matrix metabolism in patients with cruciate ligament rupture–synovial fluid concentrations of aggrecan fragments, stromelysin-1 and tissue inhibitor of metalloproteinase-1. Br. J. Rheumatol. 33, 1107–1111. Thonar, E. J., and Glant, T. (1992). Serum keratan sulfate – a marker of predisposition to polyarticular osteoarthritis. Clin. Biochem. 25, 175–180. Buckwalter, J. A. (1995). Aging and degeneration of the human intervertebral disc. Spine 20, 1307–1314. Pokharna, H. K., and Phillips, F. M. (1998). Collagen crosslinks in human lumbar intervertebral disc aging. Spine 23, 1645–1648. Kang, J. D., Stefanovic-Racic, M., McIntyre, L. A., Georgescu, H. I., and Evans, C. H. (1997). Toward a biochemical understanding of human intervertebral disc degeneration and herniation. Contributions of nitric oxide, interleukins, prostaglandin E2, and matrix metalloproteinases. Spine 22, 1065–1073. Duance, V. C., Crean, J. K., Sims, T. J., Avery, N., Smith, S., Menage, J., Eisenstein, S. M., and Roberts, S. (1998). Changes in collagen cross-linking in degenerative disc disease and scoliosis. Spine 23, 2545–2551. Sztrolovics, R., Alini, M., Mort, J. S., and Roughley, P. J. (1999). Age-related changes in fibromodulin and lumican in human intervertebral discs. Spine 24, 1765–1771. Roberts, S., Caterson, B., Menage, J., Evans, E. H., Jaffray, D. C., and Eisenstein, S. M. (2000). Matrix metalloproteinases and aggrecanase: their role in disorders of the human intervertebral disc. Spine 25, 3005–3013. Kuiper, J. I., Verbeek, J. H., Frings-Dresen, M. H., and Ikkink, A. J. (1998). Keratan sulfate as a potential biomarker of loading of the intervertebral disc. Spine 23, 657–663.

888 125. El Miedany, Y. M., Mehanna, A. N., and El Baddini, M. A. (2000). Altered bone mineral metabolism in patients with osteoarthritis. Joint Bone Spine 67, 521–527. 126. Garnero, P., Sornay-Rendu, E., Arlot, M., Christiansen, C., and Delmas, P. D. (2004). Association between spine disc degeneration and type II collagen degradation in postmenopausal women: the OFELY study. Arthritis Rheum. 50, 3137–3144. 127. Iwamoto, J., and Takeda, T. (2002). Effect of surgical treatment on physical activity and bone resorption in patients with neurogenic intermittent claudication. J. Orthop. Sci. 7, 84–90.

Kristina Åkesson 128. Pluijm, S. M., van Essen, H. W., Bravenboer, N., Uitterlinden, A. G., Smit, J. H., Pols, H. A., and Lips, P. (2004). Collagen type I alpha1 Sp1 polymorphism, osteoporosis, and intervertebral disc degeneration in older men and women. Ann. Rheum. Dis. 63, 71–77. 129. Otterness, I. G., Swindell, A. C., Zimmerer, R. O., Poole, A. R., Ionescu, M., and Weiner, E. (2000). An analysis of 14 molecular markers for monitoring osteoarthritis: segregation of the markers into clusters and distinguishing osteoarthritis at baseline. Osteoarthritis Cartilage 8, 180–185.

Index

A

Advanced glycation end-product (AGE), 49 AED, see Antiepileptic drugs (AED) AER, see Apical ectodermal ridge (AER) African-American women hip and vertebral fractures, 286 AGE, see Advanced glycation end-product (AGE) Age bone turnover biochemical markers, 570 Age-related osteoporosis bone remodeling, 672–673 bone turnover skeletal markers, 671–683 epidemiology, 672–679 calcium, 673–674 estrogen, 677–678 falls, 672 pathogenesis, 672–673 Aggrecan, 72, 88–93, 409, 429, 854–855 ADAMTS, 432–433 cartilage, 92 cartilage extracellular matrix, 72 cartilage structure and function, 91–92 catabolism, 430 composition, 90 electron micrograph, 91 extracellular assembly, 92 G1 domain, 90 G3 domain, 93 G1-G2 region, 92 metabolic products, 429–434 monoclonal antibodies, 433–434 osteoarthritis, 878 polyanionic properties, 90

AAS, see Atomic absorption spectrometry (AAS) Acid phosphatases, 165–176 biochemical and physiological significance, 166 mammalian, 166 protein structure, 168–169 ACP, see Aldol condensation product (ACP) Activated leukocyte cell-adhesion molecule (ALCAM), 231 Activating protein (AP), 155–157 structure, 156 Activating protein-1 (AP)-1 osteoblast differentiation, 234 Activin receptor-like kinase-3, 102 ADAM, see A disintegrin and metalloproteinase (ADAM) family Adenomatous polyposis coli (APC), 105 Adhesion molecules activated leukocyte cell, 231 chondroblast/chondrocyte, 130, 132 chondrocytes, 130, 142–144 functional role, 131–132 molecular structure, 130–134 osteoblast, 130, 131, 134–139 osteoclast, 130, 131, 139–142 rheumatoid arthritis cartilage metabolism, 857–858 A disintegrin and metalloproteinase (ADAM) family, 186, 853 A disintegrin and metalloproteinase thrombospondin-1 (ADAMTS), 186, 853 aggrecan, 432–433 Adolescents oral bisphosphonates osteogenesis imperfecta, 836 zoledronate osteogenesis imperfecta, 836

889

890 (Continued) protein fragments osteoarthritis, 878 structure, 88–90, 89 turnover, 92 Aggrecan aggregate electron micrograph, 91 Aggrecan family, 89 Aggregates molecular organization, 92 Aggregation molecular interactions, 90–91 stoichiometry, 92 Albers-Schonberg disease, 819 Albumin arthritis, 454 kinetics, 452 osteoarthritis, 453 ALCAM, see Activated leukocyte cell-adhesion molecule (ALCAM) Aldimine formation, 46 Aldimine lysinonorleucine, 46 Aldol condensation product (ACP), 46 Alendronate alkaline phosphatase, 660 BALP, 657 bone loss in long-term transplant recipients, 711 bone mineral density, 657 bone turnover, 653 preventing transplantation bone loss, 710 primary hyperparathyroidism, 772 PTH, 639 steroid-induced osteoporosis, 695 teriparatide, 638 Alfacalcidiol therapeutic use, 318 ALK-3 (activin receptor-like kinase-3), 102 Alkaline phosphatase (ALP), see also Bone alkaline phosphatase (BALP) alendronate, 660 bone formation, 584, 585 bone isoenzyme alendronate, 657 bone metastases, 796, 799 TIH, 798 tumor bone disease, 803 bone-specific with endogenous hyperthyroidism, 719 kidney transplantation, 704 cleidocranial dysplasia, 817 fibrodysplasia (myositis) ossificans progressiva, 824 fibrogenesis imperfecta ossium, 823 fibrous dysplasia, 826 genes and enzymes, 154 germ-cell, 153 hepatitis C-associated osteosclerosis, 822 intestinal, 153 juvenile dermatomyositis, 824

Index kidney transplantation, 703 osteoarthritis, 875–876 osteoblast secretion rate, 586 osteogenesis imperfecta, 815 placental, 153 renal osteodystrophy, 759 serum kidney transplantation, 703 renal osteodystrophy, 759 total bone metastases, 796 glucocorticoid, 692–693 in primary hyperparathyroidism, 770 Allele definition, 473 ALP, see Alkaline phosphatase (ALP) Alpha2-macroglobulin, 191 Aluminum serum renal osteodystrophy, 759 Amenorrhea athletes, 726–727 exercise-induced, 726–727 4-aminophenymercuric acetate (AMPA), 189 Amyloid A serum rheumatoid arthritis, 604 Anabolic treatment monitoring, 629–642 Anabolic window, 634 Anchorin CII rheumatoid arthritis cartilage metabolism, 857–858 Androgen(s) effect on bone cells and bone turnover, 333–335 effects on bone metabolism in men vs. women, 335–336 formation, 328 steroid osteoporosis, 691 Androgen receptors in bone cells, 331 Ankylosing spondylitis, 850 genetic markers, 595–597 link protein, 436 Ankylosis, 160 Annexin 5A rheumatoid arthritis cartilage metabolism, 857 Annexin V rheumatoid arthritis cartilage metabolism, 857 Anorexia nervosa, 727–729 Antiepileptic drugs (AED) causing osteoporosis, 730 Antiosteoporosis therapy bone marker monitoring, 682–683 bone markers, 682–683 Antiremodeling bone mass, 384 Antiresorptive therapy

891

Index bone alkaline phosphatase, 534 bone markers, 591–592 markers impact on bone turnover, 651–660 monitoring, 649–664 osteocalcin, 536 type 1 procollagen propeptides, 532 AP, see Activating protein (AP) APC, see Adenomatous polyposis coli (APC) Apert syndrome, 104 Apical ectodermal ridge (AER), 225 Apoptosis chondrocytes, 298 regulating bone balance, 381–383 rheumatoid arthritis, 604 Arachidonate, 115 Arterial calcification, 62, 160 Arthritis, see also Rheumatoid arthritis albumin, 454 reactive, 850 genetic markers, 597–598 Articular cartilage, 77, 852 structure correlated with biomechanical role, 415–416 Ascorbic acid collagen biosynthesis, 25 Aspartate residues in proteins, 398 Aspartic proteinases, 181 Asporin, 74, 75, 94 Athletes with amenorrhea, 726–727 bone mineral density, 727 Atomic absorption spectrometry (AAS) calcium, 488–489 Autosomal dominant benign osteopetrosis, 819 Axial osteomalacia, 750

B Bacterial artificial chromosomes (BAC), 465 BALP, see Bone alkaline phosphatase (BALP) Basic calcium phosphate calcium (BCP), 437 Basic multicellular units (BMU), 260 balance, 217 BCP, see Basic calcium phosphate calcium (BCP) Beta-adrenergic agonisms cancellous bone mass, 366 Beta-amino-proprionitrile (beta-APN), 49 Beta2-microglobulin renal osteodystrophy, 760–761 Biglycan, 41, 74, 93, 94–95 Biochemical markers bone formation, 530, 584–585 measurement, 529–538 response, 530 bone histomorphometry, 588–592 bone resorption, 586–587, 587 measurement, 541–557 bone turnover, 530, 569, 583–592, 614, 618–624 age, 570

analytic and biologic coefficient variation, 573 Bayesian approach, 576–577 measurement, 565–578 preanalytical variability, 566–571, 567 regression toward the mean, 574–575, 577 statistical considerations variability, 571–577 true change, 576–577 true value, 575 fracture risk, 618–624, 682 fractures, 572 glucocorticoids bone turnover, 693 monitoring in bone loss, 680–681 Paget’s disease bone assessment, 785 bone metabolism, 783–784 Turner’s syndrome bone remodeling, 725–726 validation by calcium kinetics, 584–588 Bisphosphonates adolescents osteogenesis imperfecta, 836 bone loss in long-term transplant recipients, 711 bone mass, 653–654 hepatitis C-associated osteosclerosis, 822 osteogenesis imperfecta, 835–836 age administration, 837 oral vs. intravenous, 836 treatment length, 837–838 preventing transplantation bone loss, 710–711 primary hyperparathyroidism, 772–773 steroid-induced osteoporosis, 695 Blomstrand’s lethal chondrodysplasia, 299–300 BMD, see Bone mineral density (BMD) BMP, see Bone morphogenetic proteins (BMP) BMU, see Basic multicellular units (BMU) Bone, see also Cancellous bone balance apoptosis, 381–383 biophysical characteristics, 41 building levers, 215–216 building springs, 216 carbonate content, 202 collagen metabolism, 4 compact, 201 composition, 202–204 cortical, 201 basic structural units, 380 microradiographs, 385 dense, 201 erosion focal subchondral and marginal, 846–848 extracellular matrix proteins, 205 functional properties, 79 Gla proteins, 56 leucine-rich proteoglycans, 93–96 material composition and structural design, 214–216 mineral, 202 mineral ions, 201

892 (Continued) molecular remodeling constituents, 77 periosteal formation during aging, 218 phenotypes, 207 signaling in, 259–266 spongy, 201 strength maintenance, 217–219 structure and function, 201–204 structure and strength, 213–219 subchondral, 852 trabecular, 201 Bone alkaline phosphatase (BALP) alendronate, 657 antiresorptive therapy, 534 bone formation, 533–535, 585 assays, 534 bone remodeling, 534–535 change, 534 specificity and clearance, 533–534 variability, 534 and bone formation, 585 bone metastases, 796, 799 TIH, 798 tumor bone disease, 803 Bone antiosteogenic pathways leptin and Y2-regulated bone interaction, 371–373 Bone apatite, 202 Bone biology developments, 71 Bone cells, 221–242 androgen receptors, 331 bone turnover androgens, 333–335 Bone collagen, 7–9 metabolism products, 391–403 regulation, 21–23 growth factors, 21–23 synthesis biochemical basis, 392–393 products, 392–397 Bone densitometry primary hyperparathyroidism, 769 Bone diseases rare, 811–826 transgenic models, 459–467 future directions, 467 Bone extracellular matrix, 77–79 Bone formation alkaline phosphatase, 584, 585 biochemical markers, 530, 584–585 measurement, 529–537 response, 530 bone alkaline phosphatase, 533–535, 585 assays, 534 bone remodeling, 534–535 change, 534

Index specificity and clearance, 533–534 variability, 534 calcium, 585 calcium kinetics, 585 cell biology, 204 coupling to resorption, 264–265 autocrine/paracrine regulation, 266 osteoclastic source, 266–267 pathways, 267 developmental signals, 222–227 hormones regulating, 460–461 markers, 619 in primary hyperparathyroidism, 770 osteocalcin, 535–537 assays, 535–536 bone remodeling, 536 change, 536 specificity and clearance, 535 variability, 536 total alkaline phosphatase, 533–535 assays, 533 bone remodeling, 533 specificity and clearance, 533 variability, 533 Bone histomorphometry biochemical markers, 588–592 primary hyperparathyroidism, 769–770 Bone isoenzyme alkaline phosphatase alendronate, 657 bone metastases, 796, 799 TIH, 798 tumor bone disease, 803 Bone loss, see also Postmenopausal bone loss biochemical marker monitoring, 680–681 glucocorticoids, 689–690 in long-term transplant recipients alendronate, 711 bisphosphonates, 711 calcitirol, 711 post-transplantation prevention, 709–710 trabecular, 217 Bone markers during antiresorptive and anabolic therapies, 591–592 calciopenic rickets/osteomalacia, 743–747 kidney transplantation, 703 longitudinal primary hyperparathyroidism, 771 monitoring antiosteoporosis therapy, 682–683 osteomalacia, 751 patient compliance, 659–660 phosphopenic disorders, 749–750 steroid-treated patients, 694 Bone marrow stromal cells osteogenesis imperfecta, 839 transplants bone mass, 367

893

Index osteogenesis imperfecta, 839 osteoporosis, 707 Bone mass bisphosphonates, 653–654 genistein, 652–653 high disorders causing, 818 Bone matrix proteins post-translational modification, 620–621 Bone metabolism estrogen men vs. women, 335–336 in men vs. women androgens, 335–336 Bone metastases bone isoenzyme alkaline phosphatase, 796, 799 bone resorption markers, 800–801 bone turnover markers, 796–797 hydroxyproline, 800 osteocalcin, 537 prediction, 805 type 1 procollagen propeptides, 533 Bone mineral density (BMD), 530 alendronate, 657 amenorrheic and eumenorrheic athletes, 727 antiresorptive treatment, 655–660 dual X-ray absorptiometry, 620 heart transplantation, 705 juvenile rheumatoid arthritis, 852 lung transplantation, 707 rheumatoid arthritis, 848 Bone mineralization, 204–209 electron micrograph, 205 infrared imaging, 208 Bone modeling, 209 construction during growth and decay with advancing age, 216–217 Bone morphogenetic proteins (BMP), 21, 24, 100, 102–103 fibrodysplasia (myositis) ossificans progressiva, 824 MGP, 62 osteoblasts, 265 receptor-mediated regulation, 26 receptors, 102 in skeletal development, 223–224 spondylarthropathies, 850 Bone morphogenetic protein-2 (BMP)-2 prostaglandins, 116 Bone remodeling, 209 age-related osteoporosis, 672–673 beta-adrenergic control, 367–368 biochemical markers Turner’s syndrome, 725–726 bone fragility, 385 cathepsin K, 378 central control, 361–373 cycle, 78, 377–380 decreasing benefits, 385–386 definition, 209

endogenous subclinical hyperthyroidism, 721 with endogenous subclinical hyperthyroidism, 721 fibroblast growth factor, 379 fracture protection, 383–386 fracture risk, 383–386 functions, 380–381 leptin, 362–365 sympathetic control, 365–367 neuropeptide Y system ligands, 369 new concepts in, 377–386 NPY in bone tissue, 369–370 osteoblasts, 379 osteocalcin, 379 pancreatic polypeptide and Y4 receptor, 371 regulating hormone, 209 sympathetic nervous system, 365–369 Y2 associated anabolic activity, 373 Y2 receptor deletion effects, 370–371 Y receptors, 369 Y2Y4 receptor double deletion, 371 Bone remodeling unit (BRU), 377–386 cross-sectional diagram, 378 scanning electron micrograph, 378 Bone resorption biochemical markers, 586–587 measurement, 541–557 bone formation, 264–265 osteoclastic source, 266–267 pathways, 267 calcium, 588 calcium kinetics, 588 circadian rhythm collagen related markers, 552 collagen cross-link peptides, 586–588 collagen related markers, 543–553 preanalytical issues, 552–553 cyclosporine A, 709 cytokines, 461–462 cytokines regulating, 461–462 eating disorders, 728 hydroxyproline, 543–544, 586 hypercalcemia, 351 markers, 619–620 bone metastases, 800–801 Cushing’s syndrome, 693 new vertebral fractures, 659 in primary hyperparathyroidism, 770–771 noncollagenous proteins bone matrix, 553–554 osteoarthritis, 874 osteoclast enzymes measurement, 555–557 Bone sialoprotein (BSP), 79 measurement, 543, 553–554, 554 osteoarthritis, 877–878 Paget’s disease bone, 787 Bone-specific alkaline phosphatase (BSAP) with endogenous hyperthyroidism, 719 kidney transplantation, 704

894 Bone turnover age biochemical markers, 570 age-related osteoporosis, 676–679 alendronate, 653 anorexia nervosa, 728 antiresorptive treatment, 655–660, 656–657 biochemical markers, 530, 569, 583–592, 584, 616, 618–624 age, 570 analytic and biologic coefficient variation, 573 Bayesian approach, 576–577 measurement, 565–578 preanalytical variability, 566–571 regression toward the mean, 574–575, 577 statistical considerations variability, 571–577 true change, 576–577 true value, 575 bone cells androgens, 333–335 calcitonin, 654 diet biochemical markers, 568 diurnal variation biochemical markers, 568–569 elderly women, 681 estrogen, 331–333 exercise biochemical markers, 568 fracture risk biochemical markers, 571 gender biochemical markers, 570 glucocorticoids biochemical markers, 693 histological indices, 691–692 markers, 692–694 growth biochemical markers, 570 hormone replacement therapy, 651–652 hypercalcemic states, 591 markers age-related osteoporosis, 680 clinical implications, 679–683 antiresorptive agents effect on, 651–655 antiresorptive therapy, 651–660 Bayesian approach, 662–663 bone metastases, 796–797 changes in, 661–664 distribution-based approach, 661 metastatic bone disease, 795–796 practical implications, 663–664 menstrual cycle biochemical markers, 570 molecular markers bone, cartilage, synovium, 845 normal individuals, 589 osteoporosis, 589–590 pretreatment effect on therapeutic outcomes, 650–651 primary hyperparathyroidism, 591

Index renal osteodystrophy, 590 skeletal markers age-related osteoporosis, 671–683 thyrotoxicosis, 591 Bovine tendon, 50 Brain isoenzyme creatine kinase, 820 BRU, see Bone remodeling unit (BRU) Bruck syndrome, 834 BSAP, see Bone-specific alkaline phosphatase (BSAP) BSP, see Bone sialoprotein (BSP) Bulimia, 727–729

C Cadherin(s), 130, 131, 132–133 chondrocytes, 143 osteoblasts, 137–138 osteoclast, 141 Cadherin-4 osteoblast expression, 231 Cadherin-11 osteoblast expression, 231 Calcidiol preventing transplantation bone loss, 710 Calciferol, see Vitamin D (calciferol) Calcification extraskeletal associated disorders, 823 pathological, 60 popcorn, 814 Calcimimetic agents primary hyperparathyroidism, 774 Calciopenic rickets/osteomalacia, 743–747 bone markers, 743–747 Calciotropic hormone liver transplantation, 706 Calcitirol bone loss in long-term transplant recipients, 711 Calcitonin bone turnover, 654 hepatitis C-associated osteosclerosis, 822 osteoclastogenesis, 241 preventing transplantation bone loss, 710 Calcitriol renal osteodystrophy, 759, 760 Calcium, see also Urinary calcium age-related osteoporosis, 673–674 atomic absorption spectrometry, 488–489 biochemical markers, 584–588 body distribution, 346 bone flux, 347–348 bone formation, 585 bone growth, 352–353 bone resorption, 588 dietary pycnodystosis, 820 extracellular concentration, 346–349 homeostatic response to decrease, 352

Index homeostasis, 65, 347 calcium fluxes, 349 intestinal flux, 347 measurement, 487–495 technology, 488–491 millimolar, 63 physiology, 345–357 preventing transplantation bone loss, 710 reference range, 487–488 renal fluxes, 348–349 renal handling, 691 resorption from bone, 495 serum, 487–493 circadian rhythm, 491, 492 preanalytical and biological variability, 491–493 seasonal variation, 491 venous occlusions, 491 soft tissues, 348 total vs. ionized, 487, 488 urinary excretion, 494 Calcium phosphate mineral deposition bone, 205 Calcium pyrophosphate dehydrate (CPPD), 437 Calcium-sensing receptor, 276 Campomelic dysplasia, 24 Cancellous bone, 201 basic structural units, 380 beta-adrenergic agonisms, 366 scanning electron micrographs, 384 Candidate genes genetic association studies pitfalls, 477 risk alleles, 473 Carbamazepine causing osteoporosis, 730–731 Carbonate bone, 202 Carbonic anhydrase II (CA II) deficiency osteopetrosis, 819 CART (cocaine amphetamine-regulated transcript), 368–369 Cartilage aggrecan, 91–92 aggregation function, 91–92 structure, 91–92 articular, 77, 852 biophysical characteristics, 41 collagen II, 72 collagen metabolism, 4 COMP, 75 elastic, 9 matrix composition, 9 formation developmental signals, 222–227 Gas6, 64 hyaline collagenous components, 410 type II collagen, 10 leucine-rich proteoglycans, 93–96

895 metabolic products, 421–439 MGP, 61 perlecan, 96 physical properties, 88 proteins, 76–77 proteoglycans, 87–88 tissue extract investigations, 96 Cartilage biology developments, 71 Cartilage collagen, 9–14 regulation, 24–25 synthesis cytokines, 25 growth factors, 25 Cartilage-derived morphogenetic proteins, 100 Cartilage-derived proteins, 855–856 Cartilage-derived retinoic-acid-sensitive protein (CD-RAP), 856 Cartilage extracellular matrix, 72–77 aggrecan, 72 components list, 72 LRR-rich proteins, 73 Cartilage fibrils, 409 collagen crosslinking in, 412 diameter, 410 mechanical disruption tissue, 411–412 model, 417 structure, 416–417 x-ray diffraction, 411 Cartilage Gla proteins, 56 Cartilage-inducing factors (CIP), 101 Cartilage link protein function, 91 Cartilage matrix biochemistry, 409–411 extrafibrillar, 409 future perspectives, 417–418 light and electron micrograph, 408–409 supramolecular structure, 407–418 turnover in knee joint, 453 Cartilage matrix protein (CMP), 75 Cartilage metabolism rheumatoid arthritis adhesion molecules, 857–858 Cartilage oligomeric matrix protein (COMP), 75, 438, 853 osteoarthritis, 879–880 proteolytic degradation products, 851 serum changes, 880 Cartilage-pannus junction, 850–851 Cartilage remodeling joint inflammation, 850–856 Casein kinase-1 (CK-1), 105 Cathepsin B, 182 Cathepsin D, 181 Cathepsin K, 184 bone remodeling, 378 measurement, 541, 554–555 pycnodystosis, 820 rheumatoid arthritis, 853

896 Cathepsin L, 182 measurement, 543 Cathepsin S, 182 Caucasian women hip and vertebral fractures, 286 CC chemokine receptors rheumatoid arthritis, 603 CCD, see Cleidocranial dysplasia (CCD) CCP, see Cyclic citrullinated peptide (CCP) CD44, 134, 139 chondrocytes, 143–144 osteoclast, 141–142 CD-RAP, see Cartilage-derived retinoic-acid-sensitive protein (CD-RAP) C/EBP osteoblast differentiation, 236 Cell-adhesion molecule activated leukocyte, 231 Cell-binding proteins, 204 Cell-cell adhesion proteins osteoblast maturation, 231 Cell-matrix interactions, 85 Cerebral calcification osteopetrosis, 819 CFU, see Colony-forming unit (CFU) Chemiluminescent assay 25-hydroxyvitamin D, 516–517 Chemokines rheumatoid arthritis cartilage metabolism, 857 CHF, see Congestive heart failure (CHF) Child abuse vs. osteogenesis imperfecta, 834 Children osteogenesis imperfecta oral bisphosphonates, 836 zoledronate, 836 total magnesium reference range, 501 urinary phosphate, 499 zoledronate osteogenesis imperfecta, 836 Cholecalciferol HPLC assay, 520–521 liquid chromatography-tandem mass spectroscopy assay, 522–523 measurement, 513 Chondroadherin, 74, 94 Chondroblasts adhesion molecules, 130, 132 Chondrocalcin, see C-propeptide pro [alpha(II)] chain (CPII) Chondrocytes, 852 adhesion molecules, 130, 142–144 apoptosis, 298 cadherins, 143 extracellular matrix, 422–424 immunoglobulin, 143 nutrition, 422 proliferation and differentiation, 296, 298

Index Chondrodysplasia Blomstrand’s lethal, 299–300 Jansen’s metaphyseal, 281, 299–300 Chondrogenesis control, 223 Chondroitin sulfate (CS), 86–87, 429, 854–855 dermatan sulfate chain structures, 86 osteoarthritis, 879 Chromosomes bacterial artificial, 465 Chronic renal failure bone disease with, 318–319 TRACP, 172 Ciliary neurotrophic factor (CNTF), 121 Cinacalcet primary hyperparathyroidism, 774 CIP, see Cartilage-inducing factors (CIP) Circadian rhythm bone resorption collagen related markers, 552 phosphate, 498–499 serum calcium, 491, 492 Circulating procollagen I carboxy-terminal propeptide clearances, 394–395 Citrate photometry calcium, 489 CK-1, see Casein kinase-1 (CK-1) Class I HLA-DM rheumatoid arthritis, 599–600 Class I HLA-DQ rheumatoid arthritis, 599 Class II HLA-DRB1 shared epitope rheumatoid arthritis, 598–599 Cleidocranial dysplasia (CCD), 817 Club-shaping metaphyses osteopetrosis, 819 CMP, see Cartilage matrix protein (CMP) CNTF, see Ciliary neurotrophic factor (CNTF) Cocaine amphetamine-regulated transcript (CART), 368–369 COL2A1 osteoarthritis, 872 Cole-Carpenter syndrome, 834 Collagen, see also Bone collagen; Type I collagen; Type II collagen; Type V collagen apoptosis, 3 biomechanical properties, 25 bone, 3–26 cartilage, 3–26 regulation, 24–25 synthesis, 25 cartilage synthesis cytokines, 25 growth factors, 25 cytoskeletal organization, 3 FACIT, 44 collagenous domains, 12 noncollagenous domains, 12

897

Index fibril-associated, 11–13 structural domains, 11 fibril-forming, 5 collagens, 6–7 function, 6–7 procollagens, 6 types, 6 fibrillar, 10, 43–44 gene coding, 18–20 regulatory domains, 18 transcriptional regulation, 18 genetic tools, 26 growth factors, 22 heparan-sulfate-containing, 14 3-hydroxy-pyridinium crosslinks HPLC, 546–547 measurement, 545–548 intracellular modifications, 50 measurement, 542 microfibrillar, 11, 13 network forming, 13–14 post-translational modifications, 50 pressure resistance, 3 proliferation, 3 soluble aggregate reconstitution, 412–414 tension, 3 torsion, 3 type III genes, 19–20 type IX cartilage extracellular matrix, 73 degradation, 428 type VI, 11, 13 genes, 20 type X, 13–14 genes, 20–21 type XI, 10–11 genes, 20 type XII, 12 type XIV, 12 type XIX, 12 type XVI, 12 type XX, 13 type XXI, 13 type XXII, 13 Collagenase, 231 Collagen associated molecules, 73–75 Collagen biosynthesis, 14 ascorbic acid, 25 crosslinking, 17 cytokines, 22, 23 glycosylation, 14–15 growth factors, 22 hydroxylation, 14–15 oncogenes, 23 post-translational regulation, 17 regulating factors, 21–25 steps, 14–17

steroid hormones, 23 transcription factors, 22, 23 translational regulation, 17 Collagen cross-link peptides bone resorption, 586–588 Colla2 genes, 19 Collagen families, 5–7 Collagen fibers formation, crosslinking and aging, 397–399 subpopulations, 73 Collagen fibrils cartilage, 73 crosslinking, 44, 45–48, 48 analysis, 49 maturation kinetics, 48 molecular packing, 48–49 tissue specificity, 45 decorin effects, 94 difunctional crosslinks, 45–48 interfibrillar crosslinks, 49 intermediate crosslinks, 45–48 intermolecular crosslinks biological consequences, 49 intrafibrillar crosslinks, 49 matrix constituents, 44 mechanical properties mechanical properties, 50 mineralization, 48 pyridinium crosslinks, 47 pyrrole crosslinks, 50 pyrrolic crosslinks, 47–48 Collagen genes humans, 7 mice, 7 regulation, 17 transcriptional regulation, 17–21 Collagen heterofibril, 7 Collagen I alpha I helicoidal peptide measurement, 538 Collagen metabolism bone, 4 Collagen network, 72–73 cartilage extracellular matrix, 72–73 Collagen peptides measurement, 548–552 Collagen pyridinoline crosslinks proteolytic degradation products, 853 Collagen related markers bone resorption, 543–553 preanalytical issues, 552–553 Collagen triple helix, 5–6 Collagen VI network, 76 Colony-forming unit (CFU) formation, 238 Colony-stimulating factor (CSF) bone remodeling, 378 COMP, see Cartilage oligomeric matrix protein (COMP) Competitive protein binding assay (CPBA) 25-hydroxyvitamin D, 516, 517

898 Computed tomography (CT) metastatic bone disease, 795 Congestive heart failure (CHF), 705 Contraceptives eating disorders, 728 Cornea epithelium, 10 Cortical bone, 201 basic structural units, 380 microradiographs, 385 Cortisol urinary eating disorders, 728 COX-1, see Cyclooxygenase (COX-1) CPBA, see Competitive protein binding assay (CPBA) CPII, see C-propeptide pro [alpha(II)] chain (CPII) CPPD, see Calcium pyrophosphate dehydrate (CPPD) C-propeptide pro [alpha(II)] chain (CPII), 6, 425 synovial fluid levels, 425 Cranio facial mesenchyme, 10 Craniosynostosis syndrome, 104 C-reactive protein (CRP) rheumatoid arthritis, 848 CrossLaps (CTx) antigens size distribution, 401 Crouzon syndrome, 104 CRP, see C-reactive protein (CRP) CS, see Chondroitin sulfate (CS) CsA, see Cyclosporine A (CsA) CSF, see Colony-stimulating factor (CSF) CT, see Computed tomography (CT) C-terminal crosslinked telopeptide (CTP) antigens size distribution, 401 assays, 877 bone metastases, 796–797, 801 bone resorption, 587 bone turnover, 575 Paget’s disease bone, 787 renal osteodystrophy, 760 tumor bone disease, 803 CTLA4, see Cytotoxic T-lymphocyte antigen-4 (CTLA4) CTP, see C-terminal crosslinked telopeptide (CTP) CTX, see also CrossLaps (CTx) antigens osteoporotic fractures, 621 CTX-1 osteoarthritis, 876–877 CTX-II, 881, 882 urinary levels, 883 Cushing’s syndrome, 690–691, 692 resorption markers, 693 Cyclic adenosine monophosphate second-messenger system parathyroid hormone, 281–282 Cyclic citrullinated peptide (CCP), 855 Cyclooxygenase (COX-1) prostaglandins, 116 Cyclooxygenase (COX-2) prostaglandins, 116 rheumatoid arthritis cartilage metabolism, 856–857

Index Cyclosporine A (CsA) bone resorption, 709 Cysteine proteinases, 182–184 Cystic fibrosis, 707 Cytokine-induced markers rheumatoid arthritis cartilage metabolism, 856–857 Cytokines collagen biosynthesis, 22 osteoclast formation and function, 117–118 regulating bone resorption, 463–464 renal osteodystrophy, 760 rheumatoid arthritis cartilage metabolism, 856–857 Cytotoxic T-lymphocyte antigen-4 (CTLA4) rheumatoid arthritis, 602

D 1,25(OH)2D, see Hydroxyvitamin D 25(OH)D, see Hydroxyvitamin D Decorin, 41, 73–75, 93, 94–95, 409, 410–411, 434 properties, 95 Deferoxamine (DFO) renal osteodystrophy, 759 Dehydroepiandrosterone (DHEA), 328–329, 334 Dense (compact) bone, 201 Densitometry bone primary hyperparathyroidism, 769 Dentin matrix protein 1 (DMP1), 206, 458 Dentinogenesis imperfecta translucent teeth, 813 Dentin sialophosphoprotein (DSPP), 206 Deoxypyridinoline (DPD), 854 bone metastases, 796 lysyl, 854 measurement, 542 osteoarthritis, 876–877 primary hyperparathyroidism, 797 rheumatoid arthritis, 849 tumor bone disease, 802 urinary bone loss, 694 DEQAS radioimmunoassay, 523 Dermatan sulfate, 86–87, 87 chondroitin sulfate chain structures, 86 proteoglycans in cartilage fibrils, 410 Dermatomyositis, see Juvenile dermatomyositis DFO, see Deferoxamine (DFO) DHEA, see Dehydroepiandrosterone (DHEA) DHLNL, see Dihydroxy-lysinonorleucine (DHLNL) Dickkopf (DKK), 105 Diet bone turnover biochemical markers, 568

899

Index Dietary calcium pycnodystosis, 820 Dihydroxy-lysinonorleucine (DHLNL), 46 Diphenylhydantoin causing osteoporosis, 730 Disintegrin, see A disintegrin and metalloproteinase (ADAM) family Diurnal variation bone turnover biochemical markers, 568–569 DKK, see Dickkopf (DKK) DMP1, see Dentin matrix protein 1 (DMP1) DNA polymorphisms physiological processes, 478 Doxercalciferol (Hecterol), 319 DPD, see Deoxypyridinoline (DPD) dpp, see Drosophila decapentaplegic gene product (dpp) Drosophila decapentaplegic gene product (dpp), 100 Drosophila tolloid, 16 D6S273 rheumatoid arthritis, 600 DSPP, see Dentin sialophosphoprotein (DSPP)

E Eating disorders, 727–729 bone resorption, 728 estrogen, 728 EBF, see Endochondral bone formation (EBF) ECM, see Extracellular matrix (ECM) ECMV, see Extracellular matrix vesicles (ECMV) Ectopic calcification, 823–825 Elastic cartilage, 9 matrix composition, 9 Elderly women bone turnover, 681 Embryonic stem cells (ESC), 227 Endochondral bone formation (EBF), 222 control, 223 regulation, 225–227 Endochondral ossification, 204, 209 Endogenous retrovirus rheumatoid arthritis, 604–605 Endogenous subclinical hyperthyroidism bone remodeling, 721 Endosteal hyperostosis, 822 Enzymatic measurement calcium, 490 inorganic phosphate, 498 Enzymes alkaline phosphatase, 154 leukocyte, 184 Epidermal growth factor, 105–106 Epiphycan, 94 Epithelial growth factor, 61 ePPi, see Extracellular inorganic pyrophosphate (ePPi) Ergocalciferol HPLC assay, 520–521

liquid chromatography-tandem mass spectroscopy assay, 522–523 measurement, 513 ERK, see Extracellular signal-regulated kinase (ERK) Erlenmeyer flask deformity osteopetrosis, 819 Erythrocyte sedimentation rate (ESR) rheumatoid arthritis, 849 ESC, see Embryonic stem cells (ESC) ESH, see Expansile skeletal hyperplasia (ESH) ESR, see Erythrocyte sedimentation rate (ESR) Estradiol steroid osteoporosis, 691 Estrogen age-related osteoporosis, 677–678 bone alkaline phosphatase, 534 bone metabolism men vs. women, 335–336 bone turnover, 331–333 eating disorders, 728 formation, 328 mechanisms action, 329 osteoblasts, 231, 331 osteocalcin, 536 osteoclasts, 331 osteocytes, 331 primary hyperparathyroidism, 773 type 1 procollagen propeptides, 532 Estrogen receptors in bone cells, 331 Ethnicity hip and vertebral fractures, 286 Etidronate preventing transplantation bone loss, 710 steroid-induced osteoporosis, 695 Exercise bone turnover biochemical markers, 568 preventing transplantation bone loss, 709–710 Exercise-induced amenorrhea, 726–727 Expansile skeletal hyperplasia (ESH), 780, 816 External osteolysis premature adult teeth loss, 817 Extracellular calcium-sensing receptor, 349–352 Extracellular inorganic pyrophosphate (ePPi), 437 Extracellular matrix (ECM), 77–79, 85, 422 bone, 77–79 cartilage, 72–77 aggrecan, 72 components list, 72 LRR-rich proteins, 73 chondrocytes, 422–424 proteins bone, 205 proteolysis, 182, 183 Extracellular matrix vesicles (ECMV), 205–206 electron microscopy, 206 Extracellular signal-regulated kinase (ERK), 423

900

Index

Extraskeletal calcification associated disorders, 823 Extraskeletal ossification associated disorders, 823

F FACIT, see Fibril Associated with Interrupted Triple-helix (FACIT) FAK, see Focal adhesion kinase (FAK) Falecalcitriol (Hornel), 319 Falls age-related osteoporosis, 672 Familial expansile osteolysis (FEO), 780, 816 Familial hypocalciuric hypercalcemia (FHH), 768 Fasting plasma reference range, 496 FD, see Fibrous dysplasia (FD) Femur rhizomelic shortening, 833 FEO, see Familial expansile osteolysis (FEO) FGF, see Fibroblast growth factor (FGF) FGFR, see Fibroblast growth factor receptor (FGFR) FHH, see Familial hypocalciuric hypercalcemia (FHH) Fibril assembly mechanisms, 42 Fibril-associated collagen, 11–13 structural domains, 11 Fibril Associated with Interrupted Triple-helix (FACIT), 41 Fibril Associated with Interrupted Triple-helix (FACIT) collagen, 44 collagenous domains, 12 noncollagenous domains, 12 Fibril formation, 42 collagen I, 42 factors affecting, 43 Fibril-forming collagen, 5 function, 6–7 functional diversity, 6–7 procollagens, 6 structural diversity, 6–7 types, 6 Fibrillar collagen, 10, 43–44 gene coding, 18–20 regulatory domains, 18 transcriptional regulation, 18 Fibrillogenesis collagens, 41–51 Fibril structure, 42 Fibroblast growth factor (FGF), 103–105 bone cell replication, 231 bone remodeling, 379 periosteal osteoprogenitors, 228 skeletal development, 223 Fibroblast growth factor receptor (FGFR), 104 Fibroblast-like cell type, 852–853 Fibrodysplasia (myositis) ossificans progressiva (FOP), 823–824 alkaline phosphatase, 824 bone morphogenetic proteins, 824

Fibrogenesis imperfecta ossium (FIO), 750, 822–823 alkaline phosphatase, 823 Fibromodulin, 41, 73–75, 94, 95, 434 Fibronectin, 79, 204 bone extracellular matrix, 79 Fibrous dysplasia (FD) alkaline phosphatase, 826 McCune-Albright syndrome, 825–826 monostotic, 825 polyostotic, 825 Fibulin, 77 FIO, see Fibrogenesis imperfecta ossium (FIO) Fluorometric titration calcium, 490–491 Focal adhesion kinase (FAK), 132 FOP, see Fibrodysplasia (myositis) ossificans progressiva (FOP) Fourier transform infrared imaging (FTIRI), 49 Fourier transform infrared microspectroscopy (FTIRM), 202 Fracture(s) biochemical markers, 572 fragility men vs. women, 218–219 hip African-American women, 286 osteopenia, 619 osteoporosis, 619 kidney transplantation, 703 osteoporotic CTX, 621 vertebral African-American women, 286 bone resorption markers, 659 Fracture risk age-related osteoporosis, 672 antiresorptive treatment, 656–657 biochemical markers, 618–624, 682 bone remodeling, 383–386 bone turnover biochemical markers, 571 combined assessment, 623–624 heart transplantation, 705 heritability estimates, 472 homocysteine, 622 primary hyperparathyroidism, 770 Fragility fractures men vs. women, 218–219 FTIRI, see Fourier transform infrared imaging (FTIRI) FTIRM, see Fourier transform infrared microspectroscopy (FTIRM)

G Gadolinium interference photometry calcium, 489 GAG, see Glycosaminoglycan (GAG) Gamma-carboxyglutamic acid (Gla), 56 matrix protein, 59–63 biosynthesis, 59–60

901

Index Gas6, 63–64 Protein S, 63 SHB domain, 64 GCAP, see Germ-cell alkaline phosphatase (GCAP) G3 domain, 93 Gender bone turnover biochemical markers, 570 Gene knockout models, 459 Genes alkaline phosphatase, 154 Genetic(s) terminology, 473 Genetically altered matrix proteins mutant animals, 207 Genetic markers ankylosing spondylitis, 595–597 reactive arthritis, 597–598 Genetic Markers Osteoporosis, 481 Genetic matrix diseases transgenic mice, 414–415 Genistein bone mass, 652–653 GENOMOS (Genetic Markers Osteoporosis), 481 Genotoxic stress rheumatoid arthritis, 603–604 Germ-cell alkaline phosphatase (GCAP), 153 GFP, 465 GH, see Growth hormone (GH) GIO, see Glucocorticoid-induced osteoporosis (GIO) Gla, see Gamma-carboxyglutamic acid (Gla) Glial cell line-derived neurotrophic factor, 100 Glucocorticoid-induced osteoporosis (GIO) teriparatide with antiresorptive agents, 639–640 Glucocorticoids bone loss, 689–690 bone turnover biochemical markers, 693 histological indices, 691–692 markers, 692–694 osteoblasts, 231 type 1 procollagen propeptides, 532 Glycogen synthase kinase-3 beta (GSK-3 beta), 105 Glycoproteins noncollagenous proteins, 71–80 Glycosaminoglycan (GAG), 86–88, 93, 429 chains, 96 components, 89 disaccharide sequences, 87 protein links, 86 Glycosylated hydroxyproline bone resorption, 544–545 Glycosylation, 14–15 Glycosyl-phosphatidylinositol (GPI), 157 GP-39, 76 GPI, see Glycosyl-phosphatidylinositol (GPI) Granzymes, 185 Graves’ disease, 719 hyperthyroidism, 718

Gravity bone stiffness, flexibility, lightness and speed, 213–214 Growth bone cortex thickness, 215 bone turnover biochemical markers, 570 Growth and differential factors skeletal development, 223–224 Growth factors, 99–106 collagen biosynthesis, 22 Growth hormone (GH), 99 age-related osteoporosis, 678–679 eating disorders, 728 osteogenesis imperfecta, 838–839 Turner’s syndrome, 726 Growth plate VDR inactivation, 317 GSK-3 beta, see Glycogen synthase kinase-3 beta (GSK-3 beta)

H HA, see Hyaluronan (HA) HABr, see Hyaluronic acid-binding region (HABr) Hallux valgus fibrodysplasia (myositis) ossificans progressiva, 823 Haplotype definition, 473 HA synthase genes, 436 Haversian systems, 201 HB-GAM, see Heparin-binding growth-associated molecule (HB-GAM) HCAO, see Hepatitis C-associated osteosclerosis (HCAO) HC-gp39, see Human cartilage glycoprotein-39 (HC-gp39) Heart transplantation bone mineral density, 705 fracture risk, 705 osteoporosis, 703, 705–706 Heat shock proteins (HSP) rheumatoid arthritis, 600 Hecterol, 319 Helicoidal peptide (HELP) measurement, 542 Helix-loop-helix (HLH) proteins osteoblast differentiation, 234 HELP, see Helicoidal peptide (HELP) Hematopoietic stem cells (HSC), 227 Hemoglobin photometry calcium, 489 Heparan sulfate, 85 Heparan-sulfate-containing collagen, 14 Heparin-binding growth-associated molecule (HB-GAM), 139 Hepatitis C-associated osteosclerosis (HCAO), 822 alkaline phosphatase, 822 bisphosphonates, 822 calcitonin, 822 Hereditary hypocalcemic vitamin D-resistant rickets (HVDRR), 315 HHMD, see Histidino-hydroxymerodesmosine (HHMD) High Mobility Group (HMG), 19

902 High performance liquid chromatography (HPLC), 522, 523 25-hydroxyvitamin D, 518–520 Hip fractures African-American women, 286 osteopenia, 619 osteoporosis, 619 Histidino-hydroxymerodesmosine (HHMD), 46 HLA-B27 joint diseases, 595–596, 596 reactive arthritis, 597–598 HLA class III rheumatoid arthritis, 600–601 HLH, see Helix-loop-helix (HLH) proteins HLKNL, see Hydroxylysino-5-keto-norleucine (HLKNL) HLNL, see Hydroxylysinonorleucine (HLNL) HMG, see High Mobility Group (HMG) Homocysteine fracture risk, 622 Hormone replacement therapy (HRT) bone turnover, 651–652 Hormones age-related osteoporosis, 676–679 regulating bone formation, 462–463 sex postmenopausal bone loss, 612–613 replacement in steroid-induced osteoporosis, 695 rheumatoid arthritis, 604 steroid osteoporosis, 691 Hornel, 319 HPAP, see Human prostatic acid phosphatase (hPAP) HPLC, see High performance liquid chromatography (HPLC) HRE-binding protein (HRE-BP), 170 HRT, see Hormone replacement therapy (HRT) HSC, see Hematopoietic stem cells (HSC) HSP, see Heat shock proteins (HSP) Human cartilage glycoprotein-39 (HC-gp39), 855–856, 880 Human Genome Project, 472–473 Human prostatic acid phosphatase (hPAP) androgen regulation, 176 function, 175 gene structure, 174 prostate expression, 175–176 protein structure, 174–175 Human vitamin D receptor (hVDR) gene and protein structure, 310 Humerus rhizomelic shortening, 833 Humoral hypercalcemia malignancy TRACP, 172 HVDR, see Human vitamin D receptor (hVDR) HVDRR, see Hereditary hypocalcemic vitamin D-resistant rickets (HVDRR) Hyaline cartilage collagenous components, 410 type II collagen, 10 Hyaluronan (HA), 90, 409, 422, 453 extracellular matrix, 92 metabolic products, 435–436 synthesis, 90

Index Hyaluronan-binding proteoglycans, 89 Hyaluronate receptor, 134 Hyaluronic acid, 882 Hyaluronic acid-binding region (HABr), 854–855 24-hydroxylase catabolism, 309 Hydroxylysine-glycosides measurement, 542 Hydroxylysino-5-keto-norleucine (HLKNL), 46 pyridinium crosslinks, 47 pyrrolic crosslinks, 47 Hydroxylysinonorleucine (HLNL), 46 intermolecular cross-link formation, 46 intramolecular cross-link formation, 46 Hydroxylysyl pyridinoline, 854 Hydroxyproline, 400 bone metastases, 800 bone resorption, 543–544, 586 HPLC, 544 measurement, 542 osteogenesis imperfecta, 815 3-hydroxy-pyridinium crosslinks collagen HPLC, 546–547 measurement, 545–548 Hydroxyvitamin D, 518 assay variability, 516–518 chemiluminescent assay, 516–517 competitive protein binding assay, 516, 517 contemporary normal range, 524 current assays, 524 HPLC, 518–520 measurement, 513–524, 533 contemporary approach, 520–524 normal circulating concentration, 514–516 osteoclastogenesis, 240–241 radioimmunoassay, 523–524 reference range, 514 renal osteodystrophy, 759 serum, 518 contemporary normal range, 524 sun-exposed, 515 Hypercalcemia, 351 bone resorption, 351 causes, 493 familial hypocalciuric, 768 net bone resorption, 350 Hypercalciuria osteogenesis imperfecta, 814 Hypermagnesemia causes, 502 Hyperostosis, 817–823 endosteal, 822 progressive diaphyseal dysplasia, 821 Hyperostosis corticalis generalisata, 822 Hyperparathyroidism calcium, 492 primary bisphosphonates, 772–773 renal osteodystrophy, 510, 758

903

Index Hyperphosphatemia causes, 497 kidney transplantation, 702–703 paraproteins, 498 renal osteodystrophy, 757–758 Hyperprolactinemia eating disorders, 729–730 Hyperthyroidism endogenous, 718–720 endogenous subclinical, 720–721 bone remodeling, 721 exogenous, 721 subclinical, 721–723 Graves’ disease, 718 L-T4 suppressive therapy, 721–723 osteoporosis, 717–723, 724–730 Hypocalcemia causes, 493 effectors, 352 homeostasis responses, 349–352 Hypomagnesemia causes, 502 Hypophosphatasia, 750–751 infantile, 158 Hypophosphatemia causes, 497 X-linked, 748, 749–750 Hypothyroidism serum osteocalcin, 723

I IAP, see Intestinal alkaline phosphatase (IAP) Ibandronate steroid-induced osteoporosis, 695 ICAM, see Intracellular adhesion molecule (ICAM) Idiopathic juvenile osteoporosis, 815–816 Ig, see Immunoglobulin (Ig) IGFBP, see Insulin-like growth factor binding proteins (IGFBP) IGF-I, see Insulin-like growth factor I (IGF-I) IL-1, see Interleukin-1 (IL-1) IL-4, see Interleukin-4 (IL-4) IL-6, see Interleukin-6 (IL-6) IL-10, see Interleukin-10 (IL-10) IL-11, see Interleukin-11 (IL-11) Immobilization TRACP, 172 Immunoglobulin (Ig), 133, 138 chondrocytes, 143 osteoclast, 141 Immunosuppressive therapy bone loss, 702–703 Inducible nitric oxide synthase (iNOS) chondrocytes, 437 Inducible nitric oxide synthetase (iNOS) rheumatoid arthritis cartilage metabolism, 856–857 Infantile hypophosphatasia, 158

Infectious agents joint diseases, 596–597 rheumatoid arthritis, 601–602 Inflammation markers, 849 Inorganic pyrophosphate (PPi) osteogenesis imperfecta, 815 iNOS, see Inducible nitric oxide synthase (iNOS) Insulin-like growth factor (IGF), 99–100 Insulin-like growth factor binding proteins (IGFBP), 99–100 postmenopausal bone loss, 615 Insulin-like growth factor I (IGF-I), 99–101, 120 bone and mineral homeostasis, 355 eating disorders, 728 endogenous subclinical hyperthyroidism, 721 hepatitis C-associated osteosclerosis, 822 osteoblasts, 265 postmenopausal bone loss, 615 regulating bone formation, 460–461 Turner’s syndrome, 726 Insulin-like growth factor-II (IGF-II), 99–100 Insulin receptor substrate-1 (IRS-1), 100 Integrins, 129–144, 130, 131, 132 bone extracellular matrix, 78 chondrocytes, 142–143 molecular structure, 130–132 osteoblasts, 134–137 osteoclast, 139–141 osteocytes, 134–137 Interleukin-1 (IL-1) bone, 118–119 bone remodeling, 378 cartilage, 119 renal osteodystrophy, 760 rheumatoid arthritis, 602 cartilage metabolism, 856–857, 857 Interleukin-4 (IL-4) rheumatoid arthritis, 603 Interleukin-6 (IL-6) bone, 120–121 bone remodeling, 378 cartilage, 120 renal osteodystrophy, 760 Interleukin-10 (IL-10) rheumatoid arthritis, 603 Interleukin-11 (IL-11) bone, 120–121 Interleukin-1 receptor antagonist (IL-1RA) rheumatoid arthritis, 602–603 Intestinal alkaline phosphatase (IAP), 153 Intracellular adhesion molecule (ICAM) rheumatoid arthritis cartilage metabolism, 857–858 Intracellular mediators rheumatoid arthritis, 858 Intramembranous ossification, 204 Intravenous pamidronate osteogenesis imperfecta, 835

904

Index

Ion selective electrodes (ISE) ionized calcium, 490 reference electrode measuring cell, 490 IRS-1, see Insulin receptor substrate-1 (IRS-1) ISE, see Ion selective electrodes (ISE) Isodesmosine, 545

J Jackson-Weiss syndrome, 104 JAK, see Janus kinases (JAK) Jansen’s metaphyseal chondrodysplasia, 281, 299–300 Janus kinases (JAK), 240 Joint destruction markers, 849 Joint diseases genetic markers, 595–606 infectious agents, 596–597 markers, 451–456 interpretation, 451–453 Joint immobilization PG, 423 JPD, see Juvenile Paget’s disease (JPD) JRA, see Juvenile rheumatoid arthritis (JRA) Juvenile dermatomyositis, 824–825 alkaline phosphatase, 824 dystrophic calcification, 824 ectopic calcification, 824 types, 824 Juvenile-onset spondyloarthropathy, 850 Juvenile Paget’s disease (JPD), 817, 834 Juvenile rheumatoid arthritis (JRA), 851–852 bone mineral density, 852 genetic markers, 605–606 link protein, 436

K kbp definition, 473 Keratan sulfate (KS), 87, 429, 854–855 osteoarthritis, 878–879 proteoglycans in cartilage fibrils, 410 proteolytic degradation products, 853 Keratocan, 94 Kidneys calcium homeostasis control, 348 Kidney transplantation bone markers, 703 bone-specific alkaline phosphatase, 704 fractures, 703 hyperphosphatemia, 702–703 osteoporosis, 702–703, 703 Knee albumin, 452 volume and clearance rate, 455 KS, see Keratan sulfate (KS)

L Laminin, 204 LAP, see Lysosomal AcP (LAP) Lathyrus oderatus, 49 LEF, see Lymphoid enhancer-binding factor (LEF) Leptin bone remodeling, 362–365 sympathetic control, 365–367 cancellous bone mass, 362, 363 deficiency and bone in humans, 364 deficiency and bone in mice, 362–363 energy homeostasis, 362 excess and bone in mice, 363–364 human bone, 364–365 mutation, 372 postmenopausal bone loss, 615 Leucine-rich family, 93–94 Leucine-rich proteoglycans, 93–94, 94 bone, 93–96 cartilage and bone, 93–96 glycosaminoglycan chains, 96 Leukemia inhibitory factor (LIF), 120, 121 bone, 120–121 rheumatoid arthritis cartilage metabolism, 856–857 Leukocyte enzymes, 184–185 LIF, see Leukemia inhibitory factor (LIF) Linkage definition, 473 Linkage disequilibrium definition, 473 Link protein (LP), 409, 422 cartilage, 91 function, 91 metabolic products, 435–436 turnover, 436 Lipopolysaccharides (LPS) chondrocytes, 437 Lipoprotein receptor-related protein 5 (LRP5) gain-of-function mutation, 822 Lipoxygenase, 115 Liquid chromatography/tandem mass spectrometry assay 25-hydroxyvitamin D, 520 Liver transplantation osteoporosis, 703, 705 LKNL, see Lysino-5-keto-norleucine (LKNL) LMP, see Low-molecular-weight proteasome (LMP) LNL, see Lysinonorleucine (LNL) Loads cortical bone vs. trabecular bones, 216 Locus definition, 473 Longitudinal bone markers primary hyperparathyroidism, 771 Long-term transplant recipients bone loss, 711 Low-molecular-weight proteasome (LMP) joint diseases, 597

905

Index LOX, see Lysyl oxidase (LOX) LP, see Link protein (LP) LPS, see Lipopolysaccharides (LPS) LRP5, see Lipoprotein receptor-related protein 5 (LRP5) LRR proteins, 74 LRR-rich proteins illustrations, 73 L-T4 suppressive therapy subclinical hyperthyroidism, 721–723 Lubricin, 76 Lumican, 73–75, 94, 95–96, 435 Lung transplantation, 705–706 bone mineral density, 707 osteoporosis, 703 Lymphoid enhancer-binding factor (LEF), 105 Lysine hydroxylation, 45 Lysino-5-keto-norleucine (LKNL), 46 Lysinonorleucine (LNL), 46 Lysosomal AcP (LAP), 167 Lysosomal cysteine protease pycnodystosis, 820 Lysosomal hydrolytic enzyme activity deficiency, 819 Lysyl deoxypyridinoline (DPD), 854 Lysyl oxidase (LOX), 44 procollagen, 44

M Macrophage colony-stimulating factor (M-CSF), 116, 846 in osteoclast formation and function, 118 Mad, see Mothers against dpp (Mad) Magnesium functions, 500 homeostasis, 500 measurement, 499–502 technology, 500–501 PTH, 500 serum measurement, 499–501 venous occlusions, 491 urinary, 501–502 Magnetic resonance imaging (MRI) metastatic bone disease, 795 MAK, see Mitogen-activated kinase (MAK) Mammalian acid phosphatases, 166 Mammalian tartrate-resistant acid phosphatase, 167 MAPC, see Multipotential adult progenitor cells (MAPC) Marble bone disease, 818 Markers, see also Biochemical markers; Bone markers; Molecular markers bone formation, 619 primary hyperparathyroidism, 770 bone resorption, 619–620 bone turnover age-related osteoporosis, 680 clinical implications, 679–683 antiresorptive agents effect on, 651–655

antiresorptive therapy, 651–660 Bayesian approach, 662–663 bone metastases, 796–797 changes in, 661–664 distribution-based approach, 661 metastatic bone disease, 795–796 practical implications, 663–664 cytokine-induced rheumatoid arthritis, 856–857 genetic ankylosing spondylitis, 595–597 reactive arthritis, 597–598 inflammation, 849 joint destruction, 849 joint diseases, 451–456 interpretation, 451–453 plasma pooling, 453 resorption glucocorticoids, 693–694 Mast cell proteinases, 185 Mastocytosis, 826 Matrilins, 75–76 Matrix extracellular phosphoglycoprotein (MEPE), 206 Matrix Gla Protein (MGP), 55, 56 bone, 56 bone morphogenetic proteins, 62 function, 62–63 hydroxyapatite binding, 63 interaction diagram, 59 mRNA In situ hybridization, 61 phenotypic change, 62 structure, 59–60 studies, 62 synthesis, 61 VSMC, 60 Matrix metalloproteinases (MMP), 186, 187–189, 191–192, 881–882, see also A disintegrin and metalloproteinase (ADAM) family; Tissue metalloproteinase (TIMP) activation, 190 aggrecan, 431–432 cartilage collagen degradation, 852–853 cartilage metabolism, 426–428, 427 gene expression, 188 inhibitors, 182 tissue inhibitors, 190–191 Matrix proteinases, 181–192 Maxacalcitol (Oxarol), 319 Mbp definition, 473 McCune-Albright syndrome fibrous dysplasia, 825–826 MCP-1, see Monocyte chemoattractant protein-1 (MCP-1) M-CSF, see Macrophage colony-stimulating factor (M-CSF) Mechanical stresses matrix, 423 Melanoma inhibitory activity, 856 Menopause TRACP, 172

906 Menstrual cycle bone turnover biochemical markers, 570 MEPE, see Matrix extracellular phosphoglycoprotein (MEPE) Mesenchymal stem cells, 227–230 Mesenchymal stromal cells (MSC), 227 Metal ion-dependent adhesion site (MIDAS), 130 Metalloproteinases, see Matrix metalloproteinases (MMP) Metastatic bone disease, 793–806 bone formation markers, 798–800 clinical aspects, 794 computed tomography, 795 magnetic resonance imaging, 795 monitoring, 795–796 Methylimidazoleacetic acid systemic mastocytosis, 826 MGP, see Matrix Gla Protein (MGP) Microcracks, 382 resorption space, 381 Microfibrillar collagen, 11, 13 Microphalmia-associated transcription factor (MITF), 239, 242 Microsatellite definition, 473 Microsomal PGE synthase-1 (mPGES-1) rheumatoid arthritis cartilage metabolism, 856–857 MIDAS, see Metal ion-dependent adhesion site (MIDAS) Millimolar calcium, 63 Minerals bone, 205, 215 Minisatellite definition, 473 MITF, see Microphalmia-associated transcription factor (MITF) Mitogen-activated kinase (MAK), 423 MMP, see Matrix metalloproteinases (MMP) Molecular markers bone turnover bone, cartilage, synovium, 845 Molybdate method inorganic phosphate determination, 497–498 Monoclonal antibodies aggrecan, 433–434 Monocyte chemoattractant protein-1 (MCP-1) rheumatoid arthritis cartilage metabolism, 856–857 Morphogenetic proteins, see also Bone morphogenetic proteins (BMP) cartilage-derived, 100 Mothers against dpp (Mad), 101 mPGES-1, see Microsomal PGE synthase-1 (mPGES-1) MRI, see Magnetic resonance imaging (MRI) MSC, see Mesenchymal stromal cells (MSC) Multiple myeloma, 795 osteocalcin, 799–800 TRACP, 172 Multiplexins, 14 Multipotential adult progenitor cells (MAPC), 227 Mutation definition, 473

Index Myeloma bone alkaline phosphatase, 534–535 Myositis ossificans progressiva, 823–824

N N-cadherin osteoblast expression, 231 nCaRE, see Negative calcium regulatory element (nCaRE) Negative calcium regulatory element (nCaRE), 276 Neo-osseous osteoporosis idiopathic juvenile osteoporosis, 815, 816 Neridronate osteogenesis imperfecta, 836 Network forming collagen, 13–14 Neuropeptide Y system ligands bone remodeling, 369 NFAT, see Nuclear factor activated T cells (NFAT) NHERF, see Sodium-hydrogen exchanger regulatory factor (NHERF) Nitric oxide (NO) chondrocytes, 437 Nitric oxide synthase-3 (NOS-3) chondrocytes, 437 4-nitrophenylphosphate (4-NPP), 172 3-nitrotyrosine osteoarthritis, 437 NO, see Nitric oxide (NO) Noncollagenous domains, 6 Noncollagenous proteins, 71–80 measurement, 541 NOS-3, see Nitric oxide synthase-3 (NOS-3) Nosology Silence, 812 4-NPP, see 4-nitrophenylphosphate (4-NPP) NPY in bone tissue bone remodeling, 369–370 N-terminal crosslinked telopeptide assays, 877 N-terminal propeptide, 854 NTPPPH, see Nucleoside triphosphate pyrophosphohydrolases (NTPPPH) NTX-1, 882 bone metastases, 801 bone resorption, 587 osteoarthritis, 876–877 tumor bone disease, 803 Nuclear factor activated T cells (NFAT), 462 Nuclear receptors transgenic approaches to, 462–463 Nuclear vitamin D receptor, 307, 309–313 A/B domain, 310 C-domain, 310 characteristics, 309 cross talk, 312 dimerization, 310–311 DNA-domain, 310 E-domain, 310–311 gene transcription, 311 ligand-binding-domain, 310–311

907

Index ligand-binding-function, 310 modulation, 312 phosphorylation, 312 receptor activity, 312 regulation abundance, 312 structure-function analysis, 309–310 transcriptional activation, 311–312 Nucleation, 205 Nucleoside triphosphate pyrophosphohydrolases (NTPPPH), 438 Nutrition chondrocytes, 422

O OA, see Osteoarthritis (OA) OB-cadherin osteoblast expression, 231 ODF, see Osteoclast differentiation factor (ODF) OI, see Osteogenesis imperfecta (OI) Olpadronate osteogenesis imperfecta, 836 Oncogenes collagen biosynthesis, 23 Oncostatin M (OSM), 120 effects on bone, 120–121 rheumatoid arthritis cartilage metabolism, 856–857 OPG, see Osteoprotegrin (OPG) OPN serum PP1, 159 OPPG, see Osteoporosis pseudoglioma (OPPG) syndrome Opticin, 94 Oral bisphosphonates adolescents osteogenesis imperfecta, 836 Oral contraceptives eating disorders, 728 Oral olpadronate osteogenesis imperfecta, 836 OSM, see Oncostatin M (OSM) Osmolarity regulating matrix turnover, 423 Ossification extraskeletal associated disorders, 823 Osteoadherin, 74, 79, 94 bone extracellular matrix, 79 Osteoarthritis (OA), 871–884 aggrecan, 878 protein fragments, 878 albumin, 453 biochemical aspects, 873–874 biochemical markers, 875 bone resorption, 874 bone sialoprotein, 877–878 bone turnover markers, 874–875 cartilage metabolism, 878–879 characteristics, 872 chondroitin sulfate, 879

CPII, 425 CTX-1, 876–877 deoxypyridinoline, 876–877 diagnosis, 873 etiology, 872–873 link protein, 436 treatment, 873 TSG-6, 437 Osteoblastic phenotype molecular mechanisms regulating allocation to, 229–230 Osteoblast maturation cell-cell adhesion proteins, 231 Osteoblastogenesis regulation, 229 Osteoblasts, 230–231 adhesion molecules, 130, 131, 134–139 apoptosis, 383 bone extracellular matrix, 78 bone formation, 266 bone morphogenetic proteins, 265 bone remodeling, 379 cadherin-4, 231 cadherin-11, 231 cadherins, 137–138 cell-to-cell communication, 284–285 differentiation transcriptional control, 233–236 estrogen, 331 formation, 160 glucocorticoid effects on, 690–691 glucocorticoids, 231 helix-loop-helix proteins, 234 integrins, 134–137 PTH1 receptors, 283 RANKL, 262 secretion rate alkaline phosphatase, 586 secretion rates bone marker formation, 585–586 signals, 261 Osteocalcin, 56–59, 57 antiresorptive therapy, 536 biosynthesis, 56–57 bone formation, 530, 535–537, 584, 585 assays, 535–536 bone remodeling, 536 change, 536 specificity and clearance, 535 variability, 536 bone loss, 694 bone metastases, 537 bone remodeling, 379 calcium-supplemented children, 353 under carboxylated, 535 catabolism, 58 chemical investigations, 56 endogenous hyperthyroidism, 719 fragments in urine measurement, 554–555 function, 57–58

908 (Continued) gene structure, 58 glucocorticoid, 692 heart transplantation, 705 helical regions, 57 immunochemical investigations, 56 immunoreactive forms, 535 intact and large N-terminal mid-fragment, 535 kidney transplantation, 703 liver transplantation, 705 measurement, 543 MGP, 56 m RNA, 58 osteoarthritis, 874–875, 876 osteoblast secretion rate, 586 osteomalacia, 746–747 postmenopausal osteoporosis, 622 primary hyperparathyroidism, 770, 797 promoters gene targeting, 458 regulation, 58 renal osteodystrophy, 759–760 serum hypothyroidism, 723 idiopathic juvenile osteoporosis, 816 juvenile rheumatoid arthritis, 852 spectral investigations, 56 structure, 56–57 transplantation, 708 tumor bone disease, 802 Osteoclast differentiation factor (ODF), 262 Osteoclast enzymes bone resorption measurement, 555–557 measurement, 543 Osteoclastic bone breakdown, 77 Osteoclastogenesis calcitonin, 241 inhibitor, 121 regulation, 239 Osteoclast-osteoblast cross talk hepatitis C-associated osteosclerosis, 822 Osteoclasts, 260–261, 283 adhesion molecules, 130, 131, 139–142 adhesion to bone surface, 237–238 apoptosis, 382 bone mineral resorption, 236–242 bone resorption, 874 cellular features, 237–238 bone signaling, 261 cadherins, 141 cathepsin, 182 cell-to-cell communication, 284–285 contact-dependent regulation, 261–262 cytokines, 117–118, 613 differentiation, 240 estrogen, 331 glucocorticoids, 691 hormonal regulation and transcriptional control, 240–242

Index immunoglobulin, 141 integrins, 139–141 molecular mechanisms mediating ontogeny, 238–240 physiological signaling, 262–263 secretion rates bone resorption markers, 588 TRACP, 172 Osteocytes apoptosis, 383 differentiation, 232–233 estrogen, 331 integrins, 134–137 Osteodystrophy, see Renal osteodystrophy Osteogenesis imperfecta (OI), 8, 812–815, 831–839 alkaline phosphatase, 815 biochemical defects, 812 bisphosphonates, 835–836 age administration, 837 treatment length, 837–838 bone histology, 814 vs. child abuse, 834 children oral bisphosphonates, 836 zoledronate, 836 classification, 832–833 clinical features, 812 diagnosis, 833–834 differential diagnosis, 834 expanded silence classification, 832 growth hormone, 838–839 hypercalciuria, 814 inheritance, 812 inorganic pyrophosphate, 815 intravenous pamidronate, 835 mild tarda, 814 neridronate, 836 ocular form, 834 oral vs. intravenous bisphosphonates, 836 pathogenesis, 834–835 silence types, 814 type 1, 813, 832 iliac bone samples, 838 type 3, 814 type II, 832–833 type III, 833 type IV, 833 wrist radiograph, 836 type 1 procollagen propeptides, 532 types, 812 type V, 833 type VI, 833 type VII, 833 Osteogenic lineage cells, 227–236 Osteoglycin, 94 Osteoids formation, 204 mineralization, 206

909

Index Osteolysis external premature adult teeth loss, 817 familial expansile, 780, 816 Osteomalacia, 739–751, 740, see also Calciopenic rickets/osteomalacia axial, 750 bone density, 746 bone markers, 751 calciopenic, 743–747 clinical manifestations, 742–743 etiology, 741–742 incidence, 742–743 metabolic abnormalities in, 744–745 normal mineral, 750–751 phosphopenic, 747–750 TRACP, 172 Osteonal bone mineral property, 203 Osteonectin, 79 bone extracellular matrix, 79 Osteons, 201 Osteopenia, 812–817 amenorrhea, 727–729 hip fractures, 619 idiopathic juvenile osteoporosis, 816 PTHrP haploinsufficiency, 300 Turner’s syndrome, 725–726 Osteopetrosis, 818–819 autosomal dominant benign, 819 carbonic anhydrase II deficiency, 819 cerebral calcification, 819 club-shaping metaphyses, 819 Erlenmeyer flask deformity, 819 types, 818–819 Osteopontin, 58, 204, 207 Osteoporosis, see also Postmenopausal osteoporosis; Steroid-induced osteoporosis age-related bone remodeling, 672–673 bone turnover skeletal markers, 671–683 epidemiology, 672–679 pathogenesis, 672–673 amenorrhea, 727–729 anticonvulsant drugs, 730–732 risk factors, 732 antiepileptic drugs causing, 730 bone density measurement in, 530 candidate gene polymorphisms association analysis, 476–478 candidate genes collagen type alpha1, 482–483 VDR, 482–483 candidate genes polymorphism analysis, 476, 479 carbamazepine, 730–731 complex genetic trait, 472 genes finding, 473–476 genetic variation in, 471–484

gene-wide genotype combinations, 481 genome-wide association analysis, 475–476 genome-wide linkage analysis, 474–475 haplotypes, 478–480 heart transplantation, 703, 705–706 heritability estimates, 472 hip fractures, 619 hyperthyroidism, 717–723, 724–730 kidney transplantation, 702–703, 703 meta-analyses, 480–482 neo-osseous idiopathic juvenile osteoporosis, 815, 816 secondary, 717–732 TRACP, 172 transplantation biochemical correlates pathogenesis and treatment, 701–711 pretransplantation measures, 709 prevention and management, 709–711 treatment, 711 Osteoporosis pseudoglioma (OPPG) syndrome, 105, 225, 834 Osteoporotic fractures CTX, 621 Osteoprotegrin (OPG), 239 bone modeling, 209 bone resorption, 461 discovery, 262–263 glucocorticoids, 691 osteoblasts, 690 osteoclasts, 654 postmenopausal bone loss, 615–616 Osteosclerosis, 817–823 hepatitis C-associated alkaline phosphatase, 822 Oxarol, 319 Oxygen tension in cartilage, 422

P Pachydermoperiostosis, 823 Paget’s disease, 237 biochemical markers, 532 juvenile, 817, 834 TRACP, 172 Paget’s disease bone, 779–788, 816 biochemical markers assessment, 785 bone metabolism, 783–784 bone sialoprotein, 787 cellular and molecular mediators, 783 clinical decision making in treatment monitoring, 787 clinical features, 783 diagnosis, 783–784 etiology, 779–784 genetic factors in, 779–782 pathogenesis, 779–784 predicting duration remission, 787–788 radiological changes, 783 RANK-NFKB signaling pathway, 781

910 (Continued) susceptibility loci, 780 treatment, 784–788, 785–786 viral infection in, 782–783 Pamidronate bone loss in long-term transplant recipients, 711 intravenous osteogenesis imperfecta, 835 observational trials, 836–837 preventing transplantation bone loss, 710 steroid-induced osteoporosis, 695 Pannocyte, 852–853 Panostotic fibrous dysplasia, 834 PAP, see Prostatic acid phosphatase (PAP) Paraproteins interference in photometry calcium, 489 Parathyroidectomy bone turnover markers following, 772 Parathyroid hormone (PTH), 273–287, 508 alendronate, 639 anabolic actions, 283–294 biosynthesis, 278 bone density, 632–633, 641 bone geometry, 635–637 bone metabolism, 260 bone resorption age-related and nocturnal increases, 286 idiopathic juvenile osteoporosis, 815, 816 cartilaginous growth and trabecular bone, 300 cellular and regulatory mechanisms, 630–631 effect on bone markers, 286–287 future considerations, 642 heart transplantation, 705 immunometric assays, 509–510 improving bone quality, 632–633 lung transplantation, 707 measurement, 507–511 microarchitecture, 635 mineralization, 637 nonclassical actions, 284–285 osteoblastic collagenase production, 264–265 osteoclastogenesis, 240–241 osteoclasts, 283 osteogenesis imperfecta, 839 osteoporotic vs. postmenopausal women, 286 and osteosarcoma, 641–642 postmenopausal bone loss, 614 preferential actions at selected skeletal sites, 285 animal models, 285 preferential actions at selected skeletal sties, 285–287 renal osteodystrophy, 760 schematic presentation, 509 secretion, 507–508 skeletal response to, 285 steroid osteoporosis, 691 Parathyroid hormone (PTH) gene chromosome location, 274–275 1,25 dihydroxyvitamin D, 276–277 estrogen, 278

Index expression, 275–278 extracellular calcium, 275–276 phosphate, 277–278 protein kinase A, 278 protein kinase C, 278 structure, 273–274 Parathyroid hormone (PTH)/parathyroid hormone-related protein (PTHrP) receptor interactions, 279 structure, 279–281 Parathyroid hormone (PTH) receptor catabolic actions at cellular level, 282–283 second, 282–284 Parathyroid hormone-related peptide (PTHrP), 25, 103 bone metastases, 801 bone remodeling, 300–301 cartilage regulatory factors interaction, 298–299 cartilaginous growth and trabecular bone, 300 chondrocyte proliferation, 296 eating disorders, 729 endochondral bone formation, 225 endocrine, autocrine/paracrine, and intracrine action, 297–298 genes, 294–295 haploinsufficiency, 300 interaction with PTHR-1, 296 osteoblast biology, 300–301 osteoblast growth, 228 osteoporosis, 301 as polyhormone, 295 post-translational processing, 295–296 production, 295–296 PTH-1 receptor, 299–300 receptors and signal transduction pathways, 296–297 secretion, 296 signaling and chondrocyte biology, 298–300 skeletal actions, 298–301 skeleton interaction, 293–302 tumor-induced hypercalcemia, 797 Paricalcitol (Zemplar), 319 PARP, see Proline/arginine rich domain (PARP) Pathological calcification, 60 PBM, see Peripheral blood monocytes (PBM) PDD, see Progressive diaphyseal dysplasia (PDD) PDGF, see Platelet-derived growth factor (PDGF) Periosteal bone formation during aging, 218 Periosteal osteoprogenitors fibroblast growth factor, 228 Peripheral blood monocytes (PBM), 119 Perlecan cartilage, 96 molecular-weight, 96 Pfeiffer syndrome, 104 PG, see Proteoglycans (PG) Phenobarbital causing osteoporosis, 730 Phenotypic analysis, 459–460

911

Index Phosphate bone fluxes, 354 circadian rhythm, 498–499 extracellular concentration, 353–356 homeostases, 345–357, 354, 355, 356–357, 357 intestinal fluxes, 353–354 measurement, 495–499 physiology, 346 primary hyperparathyroidism, 772 renal fluxes, 354–355 renal handling, 691 renal tubular reabsorption, 355 serum effects on calcium, 496 measurement, 495–499 technology, 497–499 reference range, 496–497 soft tissues, 356 threshold, 499 urinary, 499 children, 499 in vivo functions, 495–496 Phosphatonin, 355, 356 Phosphopenic disorders bone markers, 749–750 Phosphopenic osteomalacia, 747–750 Phosphopenic rickets, 747–750 Phosphoprotein dishevelled, 105 Phosphoprotein phosphorylation blocking, 208 Phosphorus body distribution, 353 serum venous occlusions, 491 Photometry calcium, 489 PHP, see Pseudohypoparathyroidism (PHP) PICP, see Procollagen I carboxy-terminal propeptide (PICP) PIIINP, 856 PINNP, 880 PINP, see Propeptide type I procollagen (PINP) PKA, see Protein kinase-A (PKA) PKC, see Protein kinase-C (PKC) Placental alkaline phosphatase (PLAP), 153, 155 PLAP, see Placental alkaline phosphatase (PLAP) Plasma fasting reference range, 498 Plasmin, 184–185 Plasminogen activators, 184–185 Platelet-derived growth factor (PDGF), 105–106 bone remodeling, 379 Pleiotrophin, 139 P53 mutation rheumatoid arthritis, 603–604 POA, see Primary knee osteoarthritis (POA) Poly-aspartate sequence, 207

Polymorphism definition, 473 Polyostotic fibrous dysplasia, 834 Popcorn calcifications, 814 Postmenopausal bone loss, 612–616 cytokines, 614–616 estimation rate, 617–618 growth factors, 614–616 pattern, 612–614 Postmenopausal osteoporosis bone mineral density, 616–617 bone turnover measurement biochemical markers, 617 laboratory assessment, 611–624 management, 616–624 Postmenopausal women receiving estrogen replacement therapy, 286 Post-transplantation bone loss prevention, 709–710 PPi, see Inorganic pyrophosphate (PPi) Prednisolone type 1 procollagen propeptides, 532 PRELP, see Proline Arginine-Rich End Leucine-Rich Repeat Protein (PRELP) Preproparathyroid hormone, 278 metabolism, 278–279 Primary hyperparathyroidism, 510, 767–775 alendronate, 772 biochemical profile, 768 bone densitometry, 769 bone density, 630, 774 bone formation markers, 770 bone histomorphometry, 769–770 bone markers in, 770–772 bone resorption markers, 770–771 bone turnover, 591 calcimimetic agents, 773–774 cinacalcet, 773–774 clinical presentation, 768–770 cytokines in, 772 deoxypyridinoline, 797 estrogen, 773 etiology, 767–768 extraskeletal manifestations, 768–769 fracture risk, 770 medical management, 773 skeleton in, 769–770 surgery, 772–773 treatment, 772–775 Primary hypertrophic osteoarthropathy, 823 Primary knee osteoarthritis (POA), 881 Procollagen, 5 biosynthesis, 15 chaperones, 16 processing, 16–17 Procollagen I carboxy-terminal propeptide (PICP), 393 and bone formation, 584–585 circulating clearances, 394–395

912 clearances, 394–395 distribution, 396 glucocorticoids, 693 inherited circulating concentrations, 395 kidney transplantation, 704 measurement, 542 osteoblast secretion rate, 586 osteomalacia, 746, 751 phosphopenic disorders, 749–750 vs. PINP in physiological and clinical situations, 395–396 renal osteodystrophy, 760 rheumatoid arthritis, 849 specificity and clearance, 530–531 Procollagen molecule, 393 Procollagen type II carboxy-terminal propeptide (PIICP), 854, 880 Progesterone effect on bone cells and bone turnover, 333 Progestins formation, 328 Progressive diaphyseal dysplasia hyperostosis, 821 Progressive diaphyseal dysplasia (PDD), 820–821, 821 Proinflammatory cytokines bone metabolism, 118–121 cartilage metabolism, 118–121 juvenile rheumatoid arthritis, 852 Proline/arginine rich domain (PARP), 9 Proline Arginine-Rich End Leucine-Rich Repeat Protein (PRELP), 74, 93, 94, 435 Propeptides kinetics, 43 Propeptide type I procollagen (PINP) amino-terminal, 394 bone formation, 584–585 measurement, 542 bone formation, 530 bone turnover, 575 circulating clearances, 394–395 distribution, 396 glucocorticoids, 693 osteomalacia, 746, 751 phosphopenic disorders, 749–750 rheumatoid arthritis, 848–849 specificity and clearance, 530–531 Prostaglandins, 115–117 bone formation, 117 bone resorption, 116–117 cartilage, 117 clinical implications, 117 cyclooxygenase, 116 proinflammatory cytokines, 115–120 rheumatoid arthritis cartilage metabolism, 856–857 skeletal metabolism, 115 synthesis, 116

Index synthesis regulation, 116 Prostatic acid phosphatase (PAP), 173–176 Protein(s) aspartate residues, 398 cartilage-derived, 855–856 serum proteomic analysis, 395 venous occlusions, 491 Proteinases aspartic, 181 Protein calorie malnutrition age-related osteoporosis, 672–673 Protein kinase-A (PKA), 116 prostaglandins, 116 Protein kinase-C (PKC) prostaglandins, 116 Proteoglycans (PG), 41, 76–77, 429 cartilage, 87–88 in cartilage, 87–88 dermatan sulfate in cartilage fibrils, 410 glycosaminoglycans, 85–96 hyaluronan-binding, 89 joint fluid distribution, 879 rheumatoid arthritis, 852–853 Proteoglycan tandem repeat (PTR), 90 Proteolytic degradation products matrix components, 853 Pseudohypoparathyroidism (PHP), 510–511 PTH, see Parathyroid hormone (PTH) PTR, see Proteoglycan tandem repeat (PTR) Pycnodystosis, 820 cathepsin K, 820 dietary calcium, 820 PYD, see Pyridinoline (PYD) Pyridinium crosslinks immunoassay, 547–548 ELISA, 548 Pyridinoline (PYD), 854 bone metastases, 796 crosslinks, 545, 547 HPLC, 548 hydroxylysyl, 854 measurement, 542 osteoarthritis, 876–877 renal osteodystrophy, 760 rheumatoid arthritis, 849 tumor bone disease, 802 urinary excretion, 877 Pyridinoline crosslinks osteoarthritis, 876–877

Q QTL definition, 473

913

Index

R RA, see Retinoic acid; Rheumatoid arthritis Radioimmunoassays (RIA) 25-hydroxyvitamin D, 516–517, 519 Radionuclide bone scan metastatic bone disease, 795 Raloxifene bone mass, 653 new vertebral fracture risk, 658 teriparatide, 638 RANKL, see Receptor activator nuclear factor-KB ligand (RANKL) RANTES rheumatoid arthritis cartilage metabolism, 857 Rare bone diseases, 811–826 Reactive arthritis, 850 genetic markers, 597–598 Reactive oxygen species (ROS) chondrocytes, 439 generator, 171 REC, see Rough endoplasmic reticulum (REC) Receptor activator nuclear factor-KB ligand (RANKL), 100, 118, 847 bone modeling, 209 bone remodeling, 378 bone resorption, 461 glucocorticoids, 691 osteoblasts, 690 in osteoclast formation and function, 118 osteoclasts, 654 signaling, 263–264 TNF, 120 Receptor activator nuclear factor-KB ligand (RANKL)/OPG/RANK/NF-KB signaling heritable disorders, 816–817 RECK, 191 Recombinant IGF-I (rhIGF-1) eating disorders, 726 Reiter’s syndrome, 850 Renal disease osteocalcin, 536 type 1 procollagen propeptides, 532 Renal failure chronic bone disease with, 318–319 TRACP, 172 Renal osteodystrophy, 755–762 assessment, 757 beta2-microglobulin, 760–761 biochemical assessment, 757–761 bone density measurement, 761–762 bone scintiscans, 761 bone turnover, 590 calcitriol, 759, 760 calcium measurements in, 757–758 clinical features, 756

cytokines, 760 deferoxamine, 759 with ERSD, 756 25-hydroxyvitamin D, 759 hyperparathyroidism, 510, 758 hyperphosphatemia, 757–758 PET scans, 761 phosphorus measurements in, 757–758 PTH measurements in, 758 skeletal imaging in, 761–762 vitamin D metabolites measurements in, 758–759 x-rays, 761 Renal tubular acidosis (RTA) osteopetrosis, 819 RER, 16 Resorption bone formation autocrine/paracrine regulation, 266 Resorption markers glucocorticoids, 693–694 Resorption space microcracks, 381 Retina pigment epithelium, 10 Retinoic acid, 24–25 MGP, 61 Retrovirus endogenous rheumatoid arthritis, 604–605 RF, see Rheumatoid factor (RF) RFLP definition, 473 RGD-integrin-binding peptides, 78 Rheumatoid arthritis, 843–854, see also Juvenile rheumatoid arthritis (JRA) apoptosis, 604 associated bone disease, 844–850 bone erosion features, 845 bone mineral density, 848 cartilage loss, 852–853 cartilage metabolism adhesion molecules, 857–858 anchorin CII, 857–858 annexin 5A, 857 annexin V, 857 chemokines, 857 cyclooxygenase, 856–857 cytokines, 856–857 local markers, 856–858 systemic markers, 853–856 cathepsin K, 853 class I HLA-DQ, 599 class II HLA-DRB1 shared epitope, 598–599 CPII in, 425 deoxypyridinoline, 849 D6S273, 600 endogenous retrovirus, 604–605 erythrocyte sedimentation rate, 849 generalized bone loss, 848–849

914

Index

(Continued) genetic markers, 598–605 genotoxic stress, 603–604 heat shock proteins, 600 HLA class III, 600–601 infectious agents, 601–602 link protein, 436 periarticular bone loss, 848 synovial tissues characterization, 846 cytokine analysis, 847 osteoclast-inducing and activating factors, 847 TSG-6, 437 Rheumatoid factor (RF), 849 rhIGF-1, see Recombinant IGF-I (rhIGF-1) RIA, see Radioimmunoassays (RIA) Ribbing disease, 821 Rickets, 315, 739–751, 740 calciopenic, 743–747 clinical manifestations, 742–743 etiology, 741–742 incidence, 742–743 normal mineral, 750–751 phosphopenic, 747–750 Risedronate primary hyperparathyroidism, 772–773 steroid-induced osteoporosis, 695 ROS, see Reactive oxygen species (ROS) Rough endoplasmic reticulum (REC), 14 RTA, see Renal tubular acidosis (RTA) RUNX factors osteoblast differentiation, 235–236

S Sclerosteosis, 822 SDF-1, see Stromal cell-derived growth factor (SDF-1) Seasonal variation bone turnover biochemical markers, 570, 571 serum calcium, 491 Secondary osteoporosis, 717–732, 718 Selectins, 134, 139 chondrocytes, 144 osteoclast, 142 Selective estrogen receptor modulator (SERM) therapeutic use, 531 Serine proteinases, 184–185 SERM, see Selective estrogen receptor modulator (SERM) Seronegative spondylarthropathies, 850–851 Serum alkaline phosphatase kidney transplantation, 703 osteoarthritis, 875–876 renal osteodystrophy, 759 Serum aluminum renal osteodystrophy, 759 Serum amyloid A rheumatoid arthritis, 604

Serum calcium, 487–493 circadian rhythm, 491, 492 preanalytical and biological variability, 491–493 seasonal variation, 491 venous occlusions, 491 Serum cartilage oligomeric matrix protein levels changes, 880 Serum 25-hydroxyvitamin D, 518 contemporary normal range, 524 Serum magnesium measurement, 499–501 venous occlusions, 491 Serum osteocalcin hypothyroidism, 723 idiopathic juvenile osteoporosis, 816 juvenile rheumatoid arthritis, 852 Serum phosphate effects on calcium, 496 measurement, 495–499 technology, 497–499 reference range, 496–497 Serum phosphorus venous occlusions, 491 Serum protein proteomic analysis, 395 venous occlusions, 491 Serum vitamin D eating disorders, 728 Sex hormones postmenopausal bone loss, 612–613 replacement in steroid-induced osteoporosis, 695 rheumatoid arthritis, 604 steroid osteoporosis, 691 Sex steroid-binding domain (SHB), 63 Sex steroids bone cells and bone turnover, 331–335 bone metabolism, 327–338 extraskeletal calcium homeostasis, 336–337 mechanisms action, 329–330 structure and synthesis, 328–329 transcriptional coregulator function, 330–331 SHB, see Sex steroid-binding domain (SHB) Sialoprotein, 204, see also Bone sialoprotein (BSP) SIBLING protein, 207 Signal transducers and activators transcription (STAT), 240 Skeletal development bone morphogenetic proteins, 223–224 fibroblast growth factor, 223 inductive events in, 222–225 transcriptional regulators, 226 Skeletal markers bone turnover age-related osteoporosis, 671–683 Skeletal remodeling joint inflammation, 844–852 Skeleton expansile hyperplasia, 780 SLRP, see Small Leucine-Rich Proteoglycans (SLRP) SM, see Systemic mastocytosis (SM)

915

Index Small body size (sma) genes C. elegans, 101 Small interfering RNAs, 465 Small Leucine-Rich Proteoglycans (SLRP) metabolic products, 434–435 SNP definition, 473 Sodium-hydrogen exchanger regulatory factor (NHERF) parathyroid hormone-related peptide, 296 Sodium valproate causing osteoporosis, 730 SOFA, see Stromal osteoclast-forming activity (SOFA) Solid organ transplantation osteoporosis after, 703 Soluble collagen aggregate reconstitution, 412–414, 413 Soluble phospholipase A2 (sPLA2) rheumatoid arthritis cartilage metabolism, 856–857 Somatomedin C, see Insulin-like growth factor I (IGF-I) SOX9, 103 chondrogenic potency, 24 Soy isoflavones bone mass, 652–653 SPARC, see Osteonectin Spectrometry atomic absorption calcium, 488–489 Spine degeneration markers, 882–883 sPLA2, see Soluble phospholipase A2 (sPLA2) Spondylitis, 850 Spondyloarthropathy bone morphogenetic proteins, 850 juvenile-onset, 850 Spongy (trabecular) bone, 201 Sry-type high-mobility group box, 102–103 STAT, see Signal transducers and activators transcription (STAT) Stem cells embryonic, 227 hematopoietic, 227 mesenchymal, 227–230 to osteoprogenitor, 228 populations and properties, 227–228 Steroid hormones collagen biosynthesis, 23 Steroid-induced osteoporosis, 689–695 alendronate, 695 androgens, 691 bisphosphonates, 695 epidemiology, 689–690 etidronate, 695 ibandronate, 695 pathogenesis, 690–691 treatment and follow-up, 694–695 Steroid response elements osteoblast differentiation, 236

Steroid-treated patients bone markers, 694 evaluation, 694 Stress genotoxic rheumatoid arthritis, 603–604 Stromal cell-derived growth factor (SDF-1), 240 rheumatoid arthritis cartilage metabolism, 857 Stromal cells bone marrow osteogenesis imperfecta, 839 Stromal osteoclast-forming activity (SOFA), 262 Stromelysin, 881 Subchondral bone, 852 Sympathetic nervous system bone remodeling, 365–369 Syndecans, 130, 131, 132, 133–134, 138–139 chondrocytes, 143 osteoclast, 141 Synovial fluid C-telopeptide fragments, 881 Systemic mastocytosis (SM), 826

T Tacrolimus bone resorption, 709 TALP, see Total alkaline phosphatase (TALP) TAP, see Transporter associated with antigen processing (TAP) genes Tartrate-resistant acid phosphatase (TRAP), 166–173, 167, 846 bone resorption, 588 bone turnover biochemical markers, 567 expression, 167 mammalian, 167 measurement, 543, 555–556 osteoarthritis, 877 osteogenesis imperfecta, 815 Paget’s disease bone, 787 in primary hyperparathyroidism, 770–771 serum 5b bone resorption, 173 clinical significance, 173 subcellular localization, 167–168 T-cell factor/lymphoid enhancer-binding factor (TCF/LEF), 105 T cell receptors (TCR) rheumatoid arthritis, 601 TCF/LEF, see T-cell factor/lymphoid enhancer-binding factor (TCF/LEF) TCR, see T cell receptors (TCR) Teeth premature adult loss external osteolysis, 817 translucent dentinogenesis imperfecta, 813 Telopeptides, 43

916 Teriparatide alendronate and raloxifene, 638 antiresorptive agents in glucocorticoid-induced osteoporosis, 639–640 bone microarchitecture, 636 bone mineral density, 632 bone turnover, 633–635 pharmacokinetics, 631 postmenopausal osteoporosis, 287 sequential and combination therapy with antiresorptive agent, 637–639 Testosterone steroid osteoporosis, 691 TGF, see Transforming growth factor alpha (TGF alpha) Thrombospondin-1, see A disintegrin and metalloproteinase thrombospondin-1 (ADAMTS) Thyrotoxicosis bone turnover, 591 TIH, see Tumor-induced hypercalcemia (TIH) TIMP, see Tissue metalloproteinase (TIMP) Tissue metalloproteinase (TIMP), 191–192, 881 gene expression, 188 rheumatoid arthritis, 853 Tissue metalloproteinase (TIMP-1), 120 Tissue minerals bone stiffness, 214 Tissue-nonspecific AP (TNAP), 153 clinical use, 160–162 function, 157–160 gene structure and regulation, 153–155 regulating OPN, 159 TNAP, see Tissue-nonspecific AP (TNAP) TNF, see Tumor necrosis factor (TNF) Total alkaline phosphatase (TALP) bone formation, 533–535 assays, 533 bone remodeling, 533 specificity and clearance, 533 variability, 533 bone metastases, 796 glucocorticoid, 692–693 in primary hyperparathyroidism, 770 Total magnesium children reference range, 501 determination, 501 measurement, 500–501 reference range, 501 TPA-responsive element (TRE), 188 Trabecular bone, 201 Trabecular bone loss, 217 TRACP as clinical tool, 172–173 functions, 170–171 gene, 169–170 isoforms, 169 structure, 168 TRACP-deficient mice, 171–173

Index TRAF, see Tumor necrosis factor receptor-associated factor (TRAF) family adapter proteins Transcription factors transgenic models, 463–465 Transforming growth factor alpha (TGF alpha), 105–106 activating element, 19 Transforming growth factor beta (TGF-beta), 100–102 Transforming growth factor beta 1 (TGF-beta 1) osteoblasts, 265 periosteal osteoprogenitors, 228 progressive diaphyseal dysplasia, 821 Transforming growth factor beta 2 (TGF-beta 2), 101 Transforming growth factor beta 3 (TGF-beta 3), 101 Transforming growth factor beta 4 (TGF-beta 4), 101 Transforming growth factor beta 5 (TGF-beta 5), 101 Transforming growth factor beta receptors (T beta R), 101 Transgenic mouse models, 171–173, 457–459 in bone biology, 460–465 Translucent teeth dentinogenesis imperfecta, 813 Transplantation bone loss, 707–709, 708 alendronate, 710 bisphosphonates, 710–711 calcitonin, 710 calcium, 710 etidronate, 710 exercise, 709–710 bone marrow bone mass, 369 osteogenesis imperfecta, 839 osteoporosis, 707 heart bone mineral density, 705 fracture risk, 705 osteoporosis, 703 liver osteoporosis after, 703, 705 lung, 705–706 bone mineral density, 707 osteoporosis, 703 solid organ osteoporosis after, 703 Transplantation osteoporosis biochemical correlates pathogenesis and treatment, 701–711 pretransplantation measures, 709 prevention and management, 709–711 treatment, 711 Transporter associated with antigen processing (TAP) genes joint diseases, 597 TRAP, see Tartrate-resistant acid phosphatase (TRAP) TRE, see TPA-responsive element (TRE) Trifunctional pyridinium, 48 Tumor bone disease bisphosphonate therapy monitoring, 804–805 bone isoenzyme alkaline phosphatase, 803 bone turnover markers, 802–805 Tumor-induced hypercalcemia (TIH), 797–798 bone isoenzyme alkaline phosphatase (BALP), 798

917

Index Tumor-induced osteolysis pathophysiology, 794–795 Tumor necrosis factor (TNF), 118 effects on bone, 119–120 effects on cartilage, 120 osteoclast formation, 119 receptor bone modeling, 209 Tumor necrosis factor-alpha (TNF-alpha) renal osteodystrophy, 760 rheumatoid arthritis, 600–601 Tumor necrosis factor receptor-associated factor (TRAF) family adapter proteins, 462 Tumor necrosis factor receptor II (TNFRII) rheumatoid arthritis, 602 Tumor necrosis factor stimulated gene-6 (TSG-6), 437 Turner’s syndrome, 724–726 biochemical markers bone remodeling, 725–726 growth hormone, 726 Type I collagen, 7–8 amino-terminal telopeptide antigenic epitope, 399 products derived from, 398–399 carboxyl- and amino-propeptides osteoarthritis, 876 carboxyl-terminal telopeptide antigenic epitopes in, 401 products derived from, 400–401 crosslinked N-telopeptide osteogenesis imperfecta, 815 crosslinked telopeptides carboxy-terminal, 550–552 crosslinking, 546 c-telopeptide fragments assay, 550–551 CTX epitope, 549 degradation pathways, 398 degradation products, 397–402 physiological, 401–402 genes, 18 helical domain products derived from, 399–400 helical peptide in urine assay, 551–552 molecular structure, 7–8 N-telopeptide fragments assay, 551 osteoarthritis, 874 physiological functions, 8 post-translational modification, 621 structure, 392–393 telopeptides distribution, 402 tissue distribution, 7–8 Type II collagen, 10, 854 carboxy- and amino-propeptide, 880 carboxy- and amino-telopeptide, 881–882 cleavage, 426

degradation lumbar and thoracic disk degradation, 883 fibrils, 424 genes, 19 localization, 10 proteolytic degradation, 425–426 proteolytic degradation products, 853 stability, 10 structure, 10 Type III collagen genes, 19–20 Type I procollagen carboxy-terminal vs. amino-terminal propeptides, 393–394 propeptides, 530–533 antiresorptive therapy, 532 assays, 531 bone formation changes, 532 bone metastases, 533 bone remodeling, 532–533 preanalytical variability, 531 specificity and clearance, 530–531 variability, 531–532 Type II procollagen osteoarthritis, 872 Type V collagen, 8–9 corneal stroma, 8 genes, 20 immunohistochemical identification, 9 lung, 8 placenta, 8 regulation, 23–24 skeletal muscle, 8 Type VI collagen, 11 genes, 20 Type IX collagen cartilage extracellular matrix, 73 degradation, 428 Type X collagen, 13–14 genes, 20–21 Type XI collagen, 10–11 genes, 20 Type XII collagen, 12 Type XIV collagen, 12 Type XIX collagen, 12 Type XVI collagen, 12 Type XX collagen, 13 Type XXI collagen, 13 Type XXII collagen, 13 Tyrosine kinase receptors, 63

U ucOC, see Undercarboxylated osteocalcin (ucOC) Undercarboxylated osteocalcin (ucOC), 64 Urinary calcium bone loss, 694 collection and storage, 494 excretion diurnal and seasonal variation, 494–495

918

Index

(Continued) 24-hour reference range, 493–494 measurement, 493–495 resorption, 693 Urinary cortisol eating disorders, 728 Urinary deoxypyridinoline bone loss, 694 Urinary hydroxyproline bone loss, 694 Turner’s syndrome, 725–726 Urinary magnesium, 501–502 Urinary phosphate, 499 children, 499 Urinary pyridinium bone metastases, 798 Urinary pyridinoline with endogenous hyperthyroidism, 719 Urine collection bone turnover biochemical markers, 567–568

V Van Buchem disease, 822 Vascular cell adhesion molecule (VCAM)-1 rheumatoid arthritis cartilage metabolism, 857–858 Vascular endothelial growth factor (VEGF), 105–106 Vascular permeability factor, 105–106 VCAM-1, see Vascular cell adhesion molecule (VCAM)-1 VDR, see Vitamin D receptor (VDR) VDRE, see Vitamin D responsive elements (VDRE) VEGF, see Vascular endothelial growth factor (VEGF) Vertebral fractures African-American women, 286 bone resorption markers, 659 Vgr gene products, 100 Vitamin D2 (ergocalciferol) HPLC assay, 520–521 liquid chromatography-tandem mass spectroscopy assay, 522–523 measurement, 513 Vitamin D3 (cholecalciferol) HPLC assay, 520–521 liquid chromatography-tandem mass spectroscopy assay, 522–523 measurement, 513 Vitamin D (calciferol) age-related osteoporosis, 674–676 assessment, 513–524 bioactivation, 308 and bone cells, 312–314 cell cycle genes, 313 chondrocytes, 314 deficiency, 315 extracellular matrix proteins, 313

gene expression osteoblast characteristics, 313 gene targets, 316 genomic vs. nongenomic action, 312 growth factors, 313 hereditary defects, 315–316 metabolism, 208–209 osteoblast growth, differentiation and apoptosis, 313 osteoclastogenesis and osteoclast activity, 314 pathology, 314–315 postmenopausal bone loss, 614 preventing transplantation bone loss, 710 serum eating disorders, 728 source, 308 steroid-induced osteoporosis, 695 steroid osteoporosis, 691 synthesis, 308 therapeutic use, 318–319 transcription factors in osteoblast differentiation, 313 transport, 308 Vitamin D-dependent rickets type I 1alpha-hydroxylase-null mice, 318 Vitamin D-dependent rickets type II VDR-null mice, 316–317 Vitamin D hormone, 307–319 Vitamin D metabolites bone loss in long-term transplant recipients, 711 preventing transplantation bone loss, 710 Vitamin D receptor (VDR), see also Nuclear vitamin D receptor bone pathology VDR-null mice, 317 transgenic approaches to, 464 Vitamin D-related calcium/bone diseases pathology, 315 Vitamin D responsive elements (VDRE), 307 Vitamin K, 64–65 age-related osteoporosis, 676 blood coagulation, 55 carboxylated proteins, 55 cellular functions, 55 deficiency, 65 MGP, 59 steroid-induced osteoporosis, 695 studies, 65 Vitamin K dependent proteins bone, 55–65 cartilage, 55–65 VNTR definition, 473 von Willebrand Factor A domains (vWFA), 75 vWFA, see von Willebrand Factor A domains (vWFA)

W Warfarin, 64–65 Warfarin administration, 65 WARP, 77

919

Index Williams syndrome, 311–312 Wnt in skeletal development, 224–225 Wnts, 104–105 Women elderly bone turnover, 681 postmenopausal receiving estrogen replacement therapy, 286

X X-linked hypophosphatemia, 748, 749–750

Y Y2 associated anabolic activity bone remodeling, 372 YKL-40, 76, 855, 856, 880

Y2 receptor deletion effects bone remodeling, 370–371 Y receptors bone remodeling, 369 Y2Y4 receptor double deletion bone remodeling, 371

Z Zemplar, 319 Zoledronate adolescents osteogenesis imperfecta, 836 osteogenesis imperfecta, 836 Zoledronic acid preventing transplantation bone loss, 710

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Color Plate 1

This 14-year-old girl has remarkably blue sclera characteristic of type I osteogenesis imperfecta (photographed digitally with

a built-in flash).

Color Plate 2 osteogenesis imperfecta.

The translucent teeth of dentinogenesis imperfecta encountered in a child with

Color Plate 3 Three-dimensional structure of mammalian APs. Top panel: Modeled structure of human TNAP dimer [35] based on the 3D crystallographic coordinates of PLAP [34]. One subunit is colored magenta and the other white, both in backbone representation. The location of the top “crown domain” is indicated. The site of GP1 anchor attachment, the amino terminal arm and the active site are only indicated for one subunit but are present in both. The active site metals and phosphate (CPK colors) are indicated. Bottom left panel: Active site environment of PLAP in the vicinity of the Zn1 and Zn2 metal sites and their ligands. Water molecules are shown as red spheres. Green dotted lines denote metal–ligand interactions and hydrogen bonds. Bottom right panel: Top view of the entrance to the PLAP active site, showing the position of the Y367 residue from one subunit (wireframe representation) in the immediate vicinity of the E429 residue of the other subunit (space filling representation). The lower left and right panels were initially published in [39] and are reproduced here with permission from the Journal of Biological Chemistry.

Color Plate 4 The three-dimensional structure of rat bone TRACP determined at 2.2 Å resolution (PDB code 1QHW). The core of the enzyme was formed by two seven-stranded β-sheets which packed together, making up a β-sandwich. The β-sandwich was surrounded by three solvent-exposed α-helices on both sides. The di-iron center was located on the other edge of the β-sandwich. Two N-acetylglucoseamine (NAG) molecules were linked to the side chain of Asn118. One zinc and one sulfate ion were found in the active site and one further zinc and sulfate ion were involved in crystal contact. The protease sensitive loop, seen at the upper left corner of the structure, is located close to the active site [29].

Color Plate 5 Central leptin administration reduces cancellous bone mass. Histological sections from vertebrae of 4-month-old ob/ob (A) and wild-type (B) mice following central infusion into the third ventricle with leptin or PBS for 28 days. In both leptin-deficient and wild-type mice, central leptin administration reduced trabecular bone mass and volume. Underlined numbers indicate a statistically significant difference between experimental and control groups of mice (P < 0.05). (Adapted from Ducy et al. Cell 100: 197–207, 2000.)

Color Plate 6

Peripheral leptin elevation reduces cancellous bone mass. Vertebrae of 6-month-old models in which transgenic leptin overproduction in liver led to modest (A) and extreme (B) elevation of serum leptin (ng/ml). In addition to severe lipodystrophy (not shown here), cancellous bone volume (BV/TV) was significantly decreased in both models, associated with a reduction in osteoblast activity (BFR/BS) (*P < 0.05). (Adapted from Elefteriou et al. Proc Nat Acad Sci USA 101: 3258–3263, 2004.)

Color Plate 7 Pharmacological β-adrenergic agonism reduces cancellous bone mass. Proximal tibia and vertebrae from 1-month-old ob/ob (A) and wild-type (C) mice following 6 week treatment with the β-adrenergic agonist isoproterenol (3 mg/kg/day for ob/ob and 30 mg/kg/day for wild-type). Cancellous bone volume was significantly reduced in both groups (B, D) following β-agonism associated with a reduction in osteoblast activity (BFR/BS) and number (N.Ob./B.Pm.) (*P < 0.01). (Adapted from Takeda et al. Cell 111: 305–317, 2002.)

Color Plate 8 Pharmacological β-adrenergic antagonism increases cancellous bone mass. Tibia and vertebrae from 1-month-old wild-type mice following 5-week treatment with the β-adrenergic antagonist propranolol (orally, 0.4 mg/day). Cancellous bone volume (A, BV/TV) was significantly increased following β-antagonism associated with (B) an increase in osteoblast activity (BFR/BS) and number (N.Ob./B.Pm.) (*P < 0.01). (Adapted from Takeda et al. Cell 111: 305–317, 2002.)

Color Plate 9 Reciprocal bone marrow transplants confirm sympathetic control of bone mass. Vertebrae from 6-month-old recipient mice, collected 4 months after lethal irradiation followed by transplantation with nonadherent marrow cells from donor mice. Transplantation of bone marrow cells from wild-type mice into Adrb2-/- mice, which lack β2 adrenergic receptor, reduced bone volume (BV/TV) and osteoblast activity (BFR) and surface (Ob.S/BS) to normal levels and increased urinary elimination of deoxypryidinoline (Dpd/creat.), an indicator of osteoclast function. Conversely, transplantation of Adrb2-/- marrow cells increased bone formation and decreased bone resorption in wild-type recipients (*P < 0.05). (Adapted from Elefteriou et al. Nature 434: 514–520, 2005.)

Color Plate 10

Cart gene deletion associated with decreased cancellous bone mass. Vertebrae from 6-month-old Cart-/- mice, displaying reduced cancellous bone volume (BV/TV) associated with increased osteoclast perimeter (Ob.Nb/BPm and Oc.S/BS) and bone resorption (Dpd/creat.), with no change in bone formation (BFR) (*P < 0.05). (Adapted from Elefteriou et al. Nature 434: 514–520, 2005.)