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.

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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

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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.

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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.

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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.

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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.

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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

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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

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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.

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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.

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Chapter 7 Prostaglandins and Proinflammatory Cytokines

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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.

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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

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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.

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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.

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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

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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 th