Regenerative Medicine and Biomaterials for the Repair of Connective Tissues

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Regenerative medicine and biomaterials for the repair of connective tissues

Related titles: Biomaterials and regenerative medicine in ophthalmology (ISBN 978-1-84569-443-2) With an increasingly aged population, eye diseases are becoming more widespread. Biomaterials have contributed in recent years to numerous medical devices for the restoration of eyesight, improving many patients' quality of life. This book provides readers with a definitive coverage of biomaterials and techniques used for the repair and regeneration of the eye. Initial chapters review applications in the anterior segment of the eye. A second group of chapters discuss applications in the posterior segment of the eye. The final group of chapters cover other pertinent topics such as hydrogel sealants for wound repair in ophthalmic surgery, orbital enucleation implants and polymeric materials for orbital reconstruction. Tissue engineering using ceramics and polymers (ISBN 978-1-84569-176-9) Tissue engineering is rapidly developing as a technique for the repair and regeneration of diseased tissue in the body. This authoritative and wide-ranging book reviews how ceramic and polymeric biomaterials are being used in tissue engineering. The first part of the book reviews the nature of ceramics and polymers as biomaterials together with techniques for using them such as building tissue scaffolds, transplantation techniques, surface modification and ways of combining tissue engineering with drug delivery and biosensor systems. The second part of the book discusses the regeneration of particular types of tissue from bone, cardiac and intervertebral disc tissue to skin, liver, kidney and lung tissue. Natural-based polymers for biomedical applications (ISBN 978-1-84569-264-3) Natural origin biopolymers are attractive for use in biomedical applications, partly due to their biocompatibility and degradation characteristics. Natural-based polymers for biomedical applications comprehensively reviews this important subject in six in-depth sections. Part one discusses the sources, properties, modification and processing of natural-based polymers, whilst in the second section surface aspects are reviewed. Parts three and four concern natural-origin materials for tissue engineering and regenerative medicine, discussing scaffolds, hydrogels and injectable systems. The sustained release of molecules is reviewed in the fifth part of the book, followed by the biocompatibility of natural-based polymers in the final section. Details of these and other Woodhead Publishing materials books can be obtained by: · visiting our web site at www.woodheadpublishing.com · contacting Customer Services (e-mail: [email protected]; fax: +44 (0) 1223 893694; tel.: +44 (0) 1223 891358 ext. 130; address: Woodhead Publishing Limited, Abington Hall, Granta Park, Great Abington, Cambridge CB21 6AH, UK) If you would like to receive information on forthcoming titles, please send your address details to: Francis Dodds (address, tel. and fax as above; email: francis.dodds@woodhead publishing.com). Please confirm which subject areas you are interested in.

Regenerative medicine and biomaterials for the repair of connective tissues Edited by Charles Archer and Jim Ralphs

Published by Woodhead Publishing Limited, Abington Hall, Granta Park, Great Abington, Cambridge CB21 6AH, UK www.woodheadpublishing.com Woodhead Publishing India Private Limited, G-2, Vardaan House, 7/28 Ansari Road, Daryaganj, New Delhi ± 110002, India www.woodheadpublishingindia.com Published in North America by CRC Press LLC, 6000 Broken Sound Parkway, NW, Suite 300, Boca Raton, FL 33487, USA First published 2010, Woodhead Publishing Limited and CRC Press LLC ß 2010, Woodhead Publishing Limited The authors have asserted their moral rights. This book contains information obtained from authentic and highly regarded sources. Reprinted material is quoted with permission, and sources are indicated. Reasonable efforts have been made to publish reliable data and information, but the authors and the publishers cannot assume responsibility for the validity of all materials. Neither the authors nor the publishers, nor anyone else associated with this publication, shall be liable for any loss, damage or liability directly or indirectly caused or alleged to be caused by this book. Neither this book nor any part may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, microfilming and recording, or by any information storage or retrieval system, without permission in writing from Woodhead Publishing Limited. The consent of Woodhead Publishing Limited does not extend to copying for general distribution, for promotion, for creating new works, or for resale. Specific permission must be obtained in writing from Woodhead Publishing Limited for such copying. Trademark notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation, without intent to infringe. British Library Cataloguing in Publication Data A catalogue record for this book is available from the British Library. Library of Congress Cataloging in Publication Data A catalog record for this book is available from the Library of Congress. Woodhead Publishing Limited ISBN 978-1-84569-417-3 (book) Woodhead Publishing Limited ISBN 978-1-84569-779-2 (e-book) CRC Press ISBN 978-1-4398-0110-9 CRC Press order number: N10011 The publishers' policy is to use permanent paper from mills that operate a sustainable forestry policy, and which has been manufactured from pulp which is processed using acid-free and elemental chlorine-free practices. Furthermore, the publishers ensure that the text paper and cover board used have met acceptable environmental accreditation standards. Typeset by Godiva Publishing Services Limited, Coventry, West Midlands, UK Printed by TJ International Limited, Padstow, Cornwall, UK

Contents

Contributor contact details

1

The structure and regenerative capacity of synovial joint tissues

1

Introduction Structure and function of synovial joint Joint tissues and their biomechanical properties Resident mesenchymal progenitor cells in synovial joint tissues Conclusions and future trends Sources of further information and advice References

1 2 4 16 26 28 29

A.-M. S AÈ AÈ M AÈ N E N , University of Turku, Finland, J. P. A. A R O K O S K I , University of Kuopio and Kuopio University Hospital, Finland, J. S. J U R V E L I N , University of Kuopio, Finland and I. K I V I R A N T A , University of Helsinki, Finland 1.1 1.2 1.3 1.4 1.5 1.6 1.7

2

The myofibroblast in connective tissue repair and regeneration B. H I N Z , University of Toronto, Canada

2.1 2.2 2.3 2.4 2.5 2.6 2.7 2.8 2.9

xiii

Introduction Myofibroblasts: humble tissue construction workers Know thy enemy: a quick guide to identify the myofibroblast Origins of the myofibroblast Mesenchymal stem cells (MSC) and the myofibroblast phenotype: regeneration, repair or risk? What drives myofibroblast differentiation? Lessons to be learned from the myofibroblast for the effective use of mesenchymal stem cells (MSC) Conclusions and future trends References

39 39 41 44 46 49 55 60 62 63

vi

Contents

Part I Cartilage repair and regeneration 3 3.1 3.2 3.3 3.4 3.5 3.6

4

The structure of articular cartilage

E. B. H U N Z I K E R , University of Bern, Switzerland Introduction General structure and function of articular cartilage Dual function of immature articular cartilage during postnatal growth Physiological mechanism underlying the evolution of a mature from an immature articular cartilage structure Inter-species differences in articular cartilage structure, and structure±function correlations in humans References

Measuring the biomechanical properties of cartilage cells

D. L. B A D E R and M. M. K N I G H T , Queen Mary University of London, UK 4.1 4.2 4.3 4.4 4.5 4.6 4.7

5

5.1 5.2 5.3 5.4 5.5 5.6

6

83 84 89 95 98 101

106

Introduction Measurement of chondrocyte biomechanics Intracellular biomechanics Biomechanical conditioning of chondrocytes Future trends Acknowledgements References

106 107 116 117 129 130 130

Understanding tissue response to cartilage injury

137

Introduction Clinical in vivo cartilage injury Animal models of cartilage injury In vitro cartilage injury Conclusions References

137 138 143 146 149 149

F. D E L L ' A C C I O , Barts and The London School of Medicine and Dentistry, UK and T . L . V I N C E N T , Kennedy Institute of Rheumatology, UK

Understanding osteoarthritis and other cartilage diseases T. A I G N E R , Medical Center Coburg, Germany, N. S C H M I T Z , University of Leipzig, Germany and S. S OÈ D E R , University of Erlangen-Nurnberg, Germany

6.1

83

Introduction

155

155

Contents 6.2 6.3 6.4 6.5 6.6 6.7 6.8 6.9

7

The normal joint Major cartilage pathology and pathobiology In vivo cartilage repair Grading/scoring systems for cartilage degeneration Grading/scoring of cartilage repair Sources of further information and advice Future trends References

156 157 165 168 170 173 174 174

Using animal models of cartilage repair to screen new clinical techniques

178

C. W. M c I L W R A I T H , Colorado State University, USA

7.1 7.2 7.3 7.4 7.5 7.6

8

Introduction Review of models in non-equine species Early equine models of cartilage repair Current models of cartilage repair in the equine femoropatellar and femorotibial joints Current status of animal models of cartilage repair References and further reading

Cartilage tissue repair: autologous osteochondral mosaicplasty

L. H A N G O D Y , Uzsoki Hospital, Hungary, G. K I S H , Saint George Medical, USA, T. K O R E N Y , PeÂcs Medical School, Hungary, L. R. H A N G O D Y , Semmelweis Medical School, Hungary and L. M OÂ D I S , Debrecen Medical School, Hungary 8.1 8.2 8.3 8.4 8.5 8.6 8.7

9

Introduction The development of the mosaicplasty resurfacing technique: animal and other studies Surgical technique: pre-operative planning Surgical instruments and choice of surgical technique Arthroscopic mosaicplasty Conclusions References

Cartilage tissue repair: autologous chondrocyte implantation M. B R I T T B E R G , University of Gothenburg, Sweden

9.1 9.2 9.3

vii

Introduction Chondrogeneic cell implantation Articular or other types of chondrocytes, allogeneic or autologous chondrocytes?

178 179 181 185 194 196

201

201 201 202 204 204 206 219 220

227 227 228 228

viii

Contents

9.4 9.5 9.6 9.7 9.8 9.9 9.10 9.11 9.12 9.13

Autologous chondrocytes Human clinical use and studies with autologous chondrocyte implantation Other joints besides the knee joint Clinical follow-up results Imaging evaluation of the cartilage repair Randomised controlled studies Chondrocyte implantation and osteoarthritis (OA) Conclusions and future trends Sources of further information and advice References

10

Cell sheet technologies for cartilage repair

10.1 10.2 10.3 10.4 10.5 10.6 10.7

Introduction Overview of present clinical applications Challenge for cartilage repair Properties of chondrocyte sheets Future trends in cartilage repair Regulations regarding regenerative medicine in Japan References

251 252 253 258 262 262 263

11

Cell therapies for articular cartilage repair: chondrocytes and mesenchymal stem cells

266

M. S A T O , Tokai University School of Medicine, Japan

R. ANDRIAMANALIJAONA, University of Caen, France

11.1 11.2 11.3 11.4 11.5 11.6 11.7 11.8 11.9 11.10 11.11 11.12 11.13

Introduction The chondrocyte: a unique cell The macromolecular network and biomechanical properties of cartilage Phenotypic changes Cell therapy for articular cartilage repair: chondrocytes and mesenchymal stem cells (MSCs) The use of chemical compounds to enhance matrix production Strategies to maintain the chondrogenic phenotype: the use of three-dimensional systems Use of exogenous growth factors to promote chondrogenic phenotype Use of gene therapy to deliver chondrogenic factors Control of chondrocyte phenotype and chondrogenesis by hydrostatic pressure Use of low oxygen tension in cartilage repair Conclusions Acknowledgements

230 233 239 239 242 243 244 245 246 246

251

266 267 271 273 273 277 279 281 283 284 285 289 291

Contents 11.14

12 12.1 12.2 12.3 12.4 12.5 12.6 12.7

13

13.1 13.2 13.3 13.4 13.5

References and further reading

ix 292

Scaffolds for musculoskeletal tissue engineering

301

Introduction Cell types utilized for tissue regeneration Scaffolds for engineering musculoskeletal tissue Tissue remodeling Matrix stimulation and cell±cell communications in tissue regeneration Future trends and perspectives References

301 302 306 310

H. L I and J. H. E L I S S E E F F , Johns Hopkins University, USA

Outcome measures of articular cartilage repair

M. E. T R I C E , Johns Hopkins University School of Medicine, USA Introduction Patient-based (subjective) outcome measures Process-centered (objective) outcome measures Conclusions References

314 318 319

330

330 333 338 344 344

Part II Repair of tendons and ligaments 14 14.1 14.2 14.3 14.4 14.5

15

15.1 15.2 15.3 15.4

The structure of tendons and ligaments

M. B E N J A M I N , Cardiff University, UK

Introduction Basic aspects of cell and extracellular matrix (ECM) structure Specialised regions of tendons and ligaments Conclusions References

Tendon biomechanics

M. K J á R , Bispebjerg Hospital and University of Copenhagen, Denmark, S. P. M A G N U S S O N , University of Copenhagen, Denmark and A . M A C K E Y , Bispebjerg Hospital, Denmark Introduction Biochemical adaptation of tendon to loading Biomechanics of human tendon References and further reading

351 351 354 362 368 369

375

375 376 382 388

x

16

16.1 16.2 16.3 16.4 16.5 16.6 16.7 16.8

Contents

Tendon injury and repair mechanisms

N. M A F F U L L I , Barts and The London School of Medicine and Dentistry, UK, and U. G. L O N G O , P. S H A R M A and V. D E N A R O , Campus Biomedico University, Italy

394

Introduction: tendon injury Tendinopathy Genetics Tendon rupture Pain in tendinopathy Tendon healing following acute injuries Conclusions References

394 394 398 400 406 407 410 410

Tissue engineering for ligament and tendon repair

419

17.1 17.2 17.3 17.4 17.5 17.6

Introduction Tissue engineering approaches for ligament and tendon repair Reconstruction of ligaments and tendons Future trends Sources of further information and advice References

419 420 427 428 430 430

18

Cell-based therapies for the repair and regeneration of tendons and ligaments

17

M. L E E and B. M. W U , University of California, Los Angeles, USA

R. K. W. S M I T H , The Royal Veterinary College, UK 18.1 18.2 18.3 18.4 18.5 18.6 18.7 18.8 18.9 18.10 18.11

Introduction The rationale behind the use of cells to treat tendon and ligament injuries Cell choice for tendon and ligament treatment Mixed cell populations Allogenic versus autologous sources Proposed beneficial actions of stem cells on tendon healing Stem cell-induced tenogenesis in vitro Stem cell-induced tenogenesis in vivo Conclusions Sources of further information and advice References

436 436 437 438 441 442 442 442 443 447 447 447

Contents

19

Scaffolds for tendon and ligament tissue engineering

J. C. H. G O H and S. S A H O O , National University of Singapore, Singapore 19.1 19.2 19.3 19.4 19.5 19.6

xi

452

Criteria and requirements for tendon/ligament tissue engineering scaffolds Biomaterials for tendon and ligament tissue engineering Scaffold architecture Functional scaffolds Future trends References

452 453 455 460 462 463

Index

469


(* = main contact) Editors Professor Charles Archer and Dr Jim Ralphs School of Biosciences Cardiff University Cardiff CF10 3AX UK E-mail: [email protected]; [email protected] Chapter 1 Anna-Marja SaÈaÈmaÈnen* University of Turku Department of Medical Biochemistry and Genetics Institute of Biomedicine Kiinamyllynkatu 10 FIN-20520 Turku Finland E-mail: [email protected]

Jari P. A. Arokoski University of Kuopio and Kuopio University Hospital Institute of Clinical Medicine and Department of Physical and Rehabilitation Medicine PO Box 1627 FIN-70211 Kuopio Finland E-mail: [email protected] Jukka S. Jurvelin Department of Physics University of Kuopio PO Box 1627 FIN-70211 Kuopio Finland E-mail: [email protected] Ilkka Kiviranta University of Helsinki Department of Orthopaedics and Traumatology Topeliuksenkatu 5 B FIN-00260 Helsinki Finland E-mail: [email protected]

xiv


Chapter 2 Dr. Boris Hinz Laboratory of Tissue Repair and Regeneration Matrix Dynamics Group Faculty of Dentistry University of Toronto Room 241, Fitzgerald Building 150 College Street Toronto, ON M5S 3E2 Canada E-mail: [email protected]

Chapter 5 F. Dell'accio Centre of Experimental Medicine and Rheumatology William Harvey Research Institute Barts and The London School of Medicine and Dentistry II floor John Vane Building Charterhouse Square London EC1M 6BQ UK E-mail: f.dell'[email protected]

Chapter 3 E. B. Hunziker Center of Regenerative Medicine for Skeletal Tissues Department of Clinical Research University of Bern Bern Switzerland E-mail: [email protected]

T. L. Vincent* Kennedy Institute of Rheumatology 65 Aspenlea Road London W6 8LH UK E-mail: [email protected]

Chapter 4 D. L. Bader* and M. M. Knight Medical Engineering Division School of Engineering and Materials Science Queen Mary University of London Mile End Road London E1 4NS UK E-mail: [email protected]

Chapter 6 Thomas Aigner* Medical Center Coburg Institute of Pathology Ketschendorferstr. 33 96450 Coburg Germany E-mail: [email protected] Nicole Schmitz Institute of Pathology University of Leipzig Liebigstr. 26 04103 Leipzig Germany E-mail: [email protected]

Contributor contact details Stephan SoÈder Institute of Pathology University of Erlangen-Nurnberg Krankenhausstrasse 810 91054 Erlangen Germany Chapter 7 C. W. McIlwraith Barbara Cox Anthony University Chair Orthopedic Research Laboratory Colorado State University 300 West Drake Ft Collins, CO 80525 USA E-mail: [email protected] Chapter 8 LaÂszloÂ Hangody* Medical and Health Science Center Faculty of Medicine University of Debrecen Debrecen Hungary and Department of Orthopaedics Uzsoki Hospital Budapest Hungary E-mail: [email protected] Gary Kish Wound Care Center Portsmouth Regional Hospital Portsmouth New Hampshire USA and

xv

Medical and Health Science Center Faculty of Medicine Department of Anatomy, Histology and Embryology University of Debrecen Debrecen Hungary E-mail: [email protected] TamaÂs Koreny Faculty of Medicine Institute of Musculoskeletal Surgery Department of Traumatology and Hand Surgery University of PeÂcs PeÂcs Hungary E-mail: [email protected] LaÂszloÂ Rudolof Hangody Faculty of General Medicine Doctoral School Semmelweis University Budapest Hungary E-mail: [email protected] LaÂszloÂ MoÂdis Medical and Health Science Center Faculty of Medicine Department of Anatomy, Histology and Embryology University of Debrecen Debrecen Hungary E-mail: [email protected]

xvi


Chapter 9 M. Brittberg Cartilage Research Unit University of Gothenburg Endoscopium Department of Orthopedics Kungsbacka Hospital S-434 80 Kungsbacka Sweden E-mail: [email protected]

Chapter 13 Michael E. Trice Johns Hopkins University School of Medicine Johns Hopkins Bayview Medical Center 4940 Eastern Avenue Baltimore, MD 21224 USA E-mail: [email protected]

Chapter 10 Associate Professor Masato Sato Department of Orthopaedic Surgery, Surgical Science Tokai University School of Medicine 143 Shimokasuya Isehara Kanagawa 259-1193 Japan E-mail: [email protected]

Chapter 14 Professor Michael Benjamin School of Biosciences Cardiff University Museum Avenue Cardiff CF10 3AX UK E-mail: [email protected]

Chapter 11 R. Andriamanalijaona Normandie Incubation Centre d'Innovation Technologique 17, rue Claude Bloch -BP 55027 14076 Caen cedex 5 France E-mail: [email protected] Chapter 12 Hanwei Li and Jennifer H. Elisseeff* Department of Biomedical Engineering Johns Hopkins University Clark Hall 106 3400 N. Charles Street Baltimore, MD 21218 USA E-mail: [email protected]; [email protected]

Chapter 15 Professor Michael Kjñr* Institute of Sports Medicine and Centre for Healthy Ageing Bispebjerg Hospital Denmark E-mail: [email protected] and Head of Institute of Sports Medicine Faculty of Health Sciences University of Copenhagen Bispebjerg Bakke 23 DK-2400 Copenhagen NV Denmark

Contributor contact details Professor S. P. Magnusson Professor of Musculoskeletal Rehabilitation University of Copenhagen Bispebjerg Bakke 23 DK-2400 Copenhagen NV Denmark E-mail: [email protected] Dr Abigail Mackey Institute of Sports Medicine Bispebjerg Hospital Denmark Chapter 16 N. Maffulli* Centre for Sports and Exercise Medicine Barts and The London School of Medicine and Dentistry Mile End Hospital 275 Bancroft Road London E1 4DG UK E-mail: [email protected] U. G. Longo, P. Sharma and V. Denaro Department of Orthopaedic and Trauma Surgery Campus Biomedico University Via Alvaro del Portilo, 200 00128 Trigoria Rome Italy

xvii

Chapter 17 M. Lee* and B. M. Wu Department of Bioengineering University of California, Los Angeles 5121 Engineering V 420 Westwood Plaza Los Angeles, CA 90095 USA E-mail: [email protected] Chapter 18 R. K. W. Smith Department of Veterinary Clinical Sciences The Royal Veterinary College Hawkshead Lane North Mymms Hatfield AL9 7TA UK E-mail: [email protected] Chapter 19 J. C. H. Goh* and Sambit Sahoo Department of Orthopaedic Surgery Division of Bioengineering National University of Singapore Singapore E-mail: [email protected]

1

The structure and regenerative capacity of synovial joint tissues ÈA È MA È N E N , University of Turku, Finland, A.-M. SA J . P . A . A R O K O S K I , University of Kuopio and Kuopio University Hospital, Finland, J . S . J U R V E L I N , University of Kuopio, Finland and I . K I V I R A N T A , University of Helsinki, Finland

Abstract: This chapter provides an introduction to the structure, function, and biomechanical properties of synovial joint and its tissues with special emphasis to articular cartilage. Structural elements are described at the cellular level. Major extracellular matrix components, their organization and relationship with biomechanical properties are described. Also, a short introduction to basic methodology to measure biomechanical parameters is presented. In addition, the studies demonstrating presence of human endogenous multi-potent mesenchymal stem/stromal cells (MSCs) and mesenchymal progenitors in synovial joint and associated tissues are reviewed. Possible implications of endogenous MSCs in tissue repair potential are discussed. Key words: synovial joint, biomechanics, multi-potent mesenchymal stromal cell, tissue regeneration.

1.1

Introduction

The purpose of this chapter is to introduce the reader to the structure and function of synovial joint and associated structures. First, the macroscopic structure of synovial joint compartments, cellular composition, tissue organization, and description of the major extracellular components will be reviewed in articular cartilage, subchondral bone, tendon and ligaments, synovial membrane, and meniscus. Second, the interrelationship of extracellular matrix composition with biomechanical properties of the tissues will be discussed. In addition, the basic methodology used for measuring the biomechanical parameters of joint tissues, with emphasis on articular cartilage, will be introduced. Third, the presence of resident mesenchymal stromal cells (MSCs) and progenitors in synovial joint tissues will be described, and differences in properties of MSCs derived from different intra-articular and extra-articular tissues will be discussed. MSCs represent the intrinsic repair potential in these tissues, but they also have a significant input in regulating tissue homeostasis by secreting several

2

Regenerative medicine and biomaterials in connective tissue repair

growth factors, cytokines and bioactive factors. The progress of stem cell research during the last ten years has increased our understanding of their function in tissue regeneration. However, knowledge of the role of MSCs in the repair processes of many joints tissues is still deficient.

1.2

Structure and function of synovial joint

The synovial joint is a functional unit with mechanically interacting structural components (Fig. 1.1). The development of synovial joints arises from the mesenchymal cells (Archer et al. 2003; Khan et al. 2007). Hyaline cartilage itself forms the cartilaginous model of the developing skeleton. It is replaced by bone in a process known as endochondral ossification (Mackie et al. 2008). Articular cartilage (AC) covers the ends of the bones and synovial fluid lubricates and nourishes the cartilaginous tissue. Ligaments bind the skeletal elements together and a fibrous capsule encapsulates the joint. The synovial joint (e.g. knee joint) may also contain meniscal structures internally. Each joint tissue, including bone, muscle, AC, ligaments, and tendons, has its unique

1.1 Schematic presentation of the anatomy of the knee joint. A sagittal view.


3

structure and functional properties, and changes in any component may lead to anabolic or catabolic responses in another joint component. The knee joint, joining femur and tibia in the lower limb, is the biggest synovial joint in the body. It consists of three articulating bones (femur, tibia and patella) covered by hyaline cartilage, the quadriceps and hamstring muscles, collateral and cruciate ligaments that hold the joint together, patellar tendon and the menisci. In principle, the knee is constructed of two joints, i.e. the patellofemoral joint and the tibio-femoral joint. The knee joint structures enable compression, rolling and sliding between the contacting bones. Also, the joints transmit loads of the upper body, reaching tibio-femoral loads of eight times the body weight (Kuster et al. 1997) and patello-femoral joint loads seven times the body weight (Nisell 1985) during normal daily activities, such as downhill walking and jogging. Based on the experimental analysis during simulated walking cycle, maximum tibio-femoral contact stresses of 14 MPa were recorded (Thambyah et al. 2005). This could be considered potentially dangerous for AC, knowing that there appears to be a critical threshold stress (15±20 MPa) that causes cell death and rupture of collagen network in vitro (D'Lima et al. 2001; Torzilli et al. 2006). It is suggested that the amount of loadinduced cell death is a function of the duration and magnitude of the applied load. In a healthy synovial joint, the friction coefficient between contacting articular surfaces is low; typical values between 0.01 and 0.04 have been estimated in a human hip (Unsworth et al. 1975). Several lubrication theories have been proposed, including hydrodynamic, squeeze film, and weeping and boosted mechanisms for lubrication. Each of them specifically addresses the role of intrinsic fluid and synovial fluid. Based on the operational demand, more than one lubrication mechanism is needed to provide the low friction within the synovial joint. For lubrication, the highly important mechanism is the interstitial fluid pressurization within the cartilage matrix (Ateshian 2009). However, during static loading, the boundary type of lubrication is facilitated by the molecules such as hyaluronan (HA), glycoproteins, and surface active phospholipids found in the synovial fluid (Katta et al. 2008). Functional adaptation is known as conditioning of the structure, composition and functional properties of the joint tissues to mechanical loads they are exposed to (Hyttinen et al. 2001; Tammi et al. 1987). In a healthy joint, this will lead to optimized joint function. However, mechanical conditioning may fail, leading to overloading of joint structures and, subsequently, to harmful changes in the tissues. Further, this will create an imbalance between tissue properties and functional demands, leading potentially to progressive degeneration of the structures in question. In osteoarthritis (OA), the pathological process may be triggered by changes in any joint component and no consensus has been found for the initial pathological mechanism. However, mechanical factors have been considered critical in the initiation and progress of OA. Changes in cartilage,

4


such as early superficial depletion of proteoglycans (Arokoski et al. 2000; Helminen et al. 2000) has been considered as a primary mechanism for the OA process, but alternative theories address the role of initial subchondral changes, including bone stiffening (Radin and Rose 1986; Burr and Radin 2003), in the pathological degeneration of cartilage tissue.

1.3

Joint tissues and their biomechanical properties

1.3.1

Articular cartilage

Articular cartilage is highly specialized connective tissue, and it is aneural, alymphatic, and generally considered to be avascular. Its nourishment depends on the synovial fluid and subchondral bone. The thickness of AC varies from some micrometers to a few millimeters in different cartilage areas within the joint, in different joints, and animal species. Biomechanically, the primary function of the AC is to provide a covering material that protects the subchondral bone and provides a smooth, lubricated surface that facilitates movements with little friction between the articulating surfaces. Cartilage has the ability to reduce the nominal contact pressures and to increase joint congruence. Traditionally, AC is divided into three pseudo-stratified zones that are separated by a tidemark from the deepest calcified zone (Fig. 1.2). This classification is based on collagen fibril orientation and the organization and morphology of chondrocytes at different depths throughout AC. The zones can be characterized as follows: · Zone 1: Superficial (tangential) zone; adjacent to the joint cavity occupying 5±10% of the matrix volume. It is characterized by relatively low proteoglycan (PG) content. The collagen fibrils are closely packed and the orientation of fibrils is predominantly tangential (parallel) to the surface. The cells are discoidal with their long axes parallel to the surface. · Zone II: Middle (transitional or intermediate) zone; occupies up to 45% of the matrix volume. It is characterized by a significant increase in the PG content. The collagen fibrils are randomly oriented. The cells are spheroidal and equally spaced. · Zone III: Deep (radial) zone; occupies up to 45% of the matrix volume and has the highest PG content. The distance between collagen fibrils increases and they are arranged perpendicularly (radially) to the articular surface. Spheroidal cells often form columns aligned with the radial collagen fibrils. · Zone IV: Calcified zone; adjacent to the subchondral bone occupies 5±10% of the matrix volume. There are a few chondrocytes, the matrix is mineralized with crystals of calcium salts and the PG content is low. The collagen fibrils are radially aligned. A borderline, called the `tidemark', separates the calcified zone from the deep zone.


5

1.2 A schematic presentation of the structure of articular cartilage illustrating the different zones and regions of articular cartilage and subchondral bone. Collagen fibrils are oriented tangentially in the superficial zone and radially in the middle and deep zones of the articular cartilage. Chondrocytes in the superficial zone are discoidal and in the deep zone spheroidal and often form columns. Tidemark at the mineralization front separates deep zone and calcified zone. Subchondral bone plate and trabecular bone provide support to the articular cartilage and is vascularized.

This zonal pattem is present in adult AC of most species, although the relative proportion of each zone varies. The extracellular matrix surrounding each chondrocyte can be subdivided into three discrete zones, which are the pericellular, territorial, and interterritorial matrices. Chondrocytes Cell volume, 2±10% of the total tissue volume, in AC is low in comparison with other tissues. Chondrocytes are the only cell type in AC. As being entrapped within this highly organized matrix and isolated in the lacunae that is called as chondron, mature chondrocytes have essentially no migratory potential, although some in vitro motility has been shown (Poole 1997; Morales 2007). Chondrocytes are ultimately responsible for the integrity, organization, and maintenance of the extracellular matrix (ECM) (Bhosale and Richardson 2008).

6


Three subpopulations of chondrocytes with different morphologies have been identified in AC (Kouri et al. 1996). Type 1 cells found in superficial and upper middle zones of AC represent typical chondrocytes. Type 2 cells represent secretory, fibroblast-like cells that are more abundant in the non-fibrillated osteoarthritic samples than in healthy cartilage. Type 3 cells are degenerating chondrocytes found throughout the OA cartilage although more frequently in the fibrillated tissue. Extracellular matrix Articular cartilage consists of two principal phases: a solid organic ECM, which is predominantly composed of collagen fibrils and PGs, and a mobile interstitial fluid phase. Water and inorganic salts make up about 70±80% of the tissue wet weight. The distribution of the chondrocytes and ECM constituents varies throughout the thickness of the cartilage. The structure of cartilage ECM is reviewed in more detail in Chapter 3. Type II collagen is the most abundant protein in the ECM of AC, where it forms a three-dimensional fibrous network of alpha1 (II) collagen homotrimers assembled into copolymeric bundles with type IX and XI collagens (Eyre 2002). Also minor amounts of type III, VI, XII and XIV collagens are found in cartilage. Type X collagen is normally restricted in the hypertrophic zone in AC cartilage and growth plate. The three-dimensional network of collagens is filled with PGs that are macromolecules consisting of a central core protein to which one or more glycosaminoglycan (GAG) chains and oligosaccharides are attached (Knudson and Knudson 2001; Roughley 2006). On mass basis, aggrecan is the most abundant PG in cartilage (Fig. 1.3). The core protein of the aggrecan can be divided into N-terminal HA binding region with a G1 globular domain, small interglobular domain, another N-terminal globular domain G2, keratan sulfate attachment domain KS, two chondroitin sulfate attachment domains CS1 and CS2, and a lectin binding C-terminal globular domain G3 (Fig. 1.3). G1 domain interacts with link protein to stabilize the binding of aggrecan molecules into large aggregates with HA. The GAGs are unbranched carbohydrates made up of repeating disaccharide units with negatively charged sulfate and carboxyl groups responsible for the water binding properties of the aggrecan molecule that regulate the elastic properties of the cartilage (Cowin and Doty 2007a). Chondrocytes synthesize also numerous other non-collagenous ECM components (Roughley 2001, 2006). Small leucine-rich repeat proteoglycan (SLRP) family members decorin, biglycan, fibromodulin, lumican, and others have functions in regulating the collagen fibrillogenesis, activity and distribution of different cytokines and growth factors, as well as in cell signaling (Schaefer and Iozzo 2008). Several other PGs and glycoproteins are found in the extracellular space, e.g., cartilage oligomeric protein (COMP), versican and tenascin, or on


7

1.3 Schematic presentation of the major extracellular matrix components of articular cartilage. (a) Aggrecan molecules attach to hyaluronan with link protein to form large (> 1 106 Da) proteoglycan aggrecates which are entrapped within the network of type II collagen fibrils. (b) Characteristic structural domains of aggrecan. Hyaluronan (HA); two interglobular domains (IGD) G1 and G2; keratan sulfate binding region (KS), chondroitin sulfate binding region 1 (CS1), chondroitin sulfate binding region 2 (CS2), and a Cterminal globular domain (G3) (adapted from Qu 2007).

the chondrocyte surface, e.g., perlecan, syndecans, integrins and growth factor receptors (van der Kraan et al. 2002; Chiquet-Ehrismann and Tucker 2004; Melrose et al. 2008; Shakibaei et al. 2008). These factors and their relationship in tissue function, homeostasis and degeneration have been reviewed by, for example, Gentili and Cancedda (2009), Goldring et al. (2008) and Goldring and Marcu (2009), and are more thoroughly reviewed in the other chapters of this book. The poor intrinsic repair capacity of adult AC results from avascular nature of the tissue. Lacking the blood vessels, the damage in AC produces no inflammatory reaction, which would lead to chemotactic recruitment of the repair cells, typical in the repair process of other tissues. Also, synthesis and turnover of type II collagen in adult AC are exceptionally low, the half-life of the collagen molecules being over one hundred years (Maroudas et al. 1992; Verzijl et al. 2000). Hence, any marked injuries in the collagen network are likely to initiate a cascade leading to progressive degradation of AC (Aigner and StoÈve 2003).

1.3.2

Subchondral bone

Subchondral bone is formed via endochondral ossification of the cartilage template at the secondary ossification centers of the bone epiphyses during joint development (Burr 2004; Mackie et al. 2008). Subchondral bone provides support to AC, and biomechanically it has an essential role on cartilage homeostasis. Subchondral bone consists of the subchondral bone plate (SBP) to which the subchondral trabecular bone (STB) is attached (Fig. 1.2) (Imhof et al. 2000; Burr

8


2004). Both SBP and STB are formed of bone lamellae. Subchondral bone is composed of two types of lamellae: concentric lamellae around the osteons and flat lamellae. SBP, like compact bone, is composed of subunits called osteons consisting of concentric lamellae surrounding the central (Haversian) canal. STB and the inner surface of SBP are covered by osteoblasts and osteoclasts. Physiologically and mechanistically distinct STB is highly vascularized and it provides another route for cartilage nutrition in addition to synovial fluid (Imhof et al. 2000). The subchondral bone structure, i.e., SBP thickness and STB density vary with region in the joint (Oettmeier et al. 1992). Based on Wolff's law, both the bone density and the organization of bone trabeculae correlate with the magnitude and direction of compressive and tensile stresses of loading (Wolff 1892). There are five different bone cell types ± osteocytes, osteoprogenitors, bone lining cells, osteoblasts, and osteoclasts. Osteoblasts and osteoclasts form bone remodeling units that maintain the integrity of the bone and balance between deposited and resorbed bone (Cowin and Doty 2007b; Bartl and Frisch 2009). Bone matrix is composed of organic and inorganic components. Up to 88% of organic matrix is collagen, mainly type I collagen, which forms an organized template for the matrix mineralization by deposition of hydroxyapatite and apatite. In addition, organic matrix contains up to 12% of the dry weight of osteocalcin, osteonectin, several phosphoproteins, lipids and proteoglycans. Bone marrow of the trabecular bone maintains a heterogeneous population of various multi-potent mesenchymal stromal cells that provide progenitors for differentiation of osteochondral and other mesenchymal cell lineages as well as trophic environment for hematopoiesis.

1.3.3

Meniscus

Menisci are semi-lunar discs between tibia and femur in the knee joint protecting AC from excess shocks by distributing loads and stabilizing joints during movement (Fig. 1.4(a)) (Setton et al. 1999a; Sweigart and Athanasiou 2001). Meniscus tissue is composed of outer, dense connective tissue and inner fibrocartilage regions (Fig. 1.4(b)) (Verdonk et al. 2005). The outer region is vascularized dense fibrous connective tissue that connects to the internal knee joint capsule. Its matrix is maintained by fibroblast-like fibrochondrocytes which produce type I collagen fibrils. Elastin fibers as bridge-like connections between collagen fibrils have been suggested as contributing to the recovery after deformation (Ghosh and Taylor 1987). The inner region of the meniscus is avascular, aneural, and alymphatic tissue, which is why, similar to cartilage, its repair capacity is lower than that of the outer region. Cells in the inner region are chondrocyte-like fibrochondrocytes, and its matrix contains many similar components common to cartilage and tendon. Type II collagen forms about 60% and type I collagen about 40% of the total collagen in the inner region. Minor


9

1.4 Schematic presentation of the menisci in the knee joint. (a) Macroscopic view of the proximal tibia with menisci, and insertion sites of ligaments. (b) Vascularized outer region and avascular inner region of the meniscus. (c) Schematic organization of the collagen fibers in different layers of the meniscus. Collagen fibers at the superficial layer on both tibial and femoral sides are thin and intersect at various angles. Below that on both sides are lamellar layers where collagen fibers are arranged into lamella-like bundles that are radially arranged in the external circumference of the anterior and posterior segments of bundles, and intersect at various angles in the other regions. In the central main layer the bundles of collagen fibers are oriented in a circular manner, with a few interwoven radial tie fibers in the internal circumference. At the external circumference, also loose connective tissue from the capsule penetrates the central main layer (arrow) ((b) adapted from Verdonk et al. 2005 and (c) adapted from Petersen and Tillmann 1998).

amounts of type III, V and VI collagens have been found in both regions (Sweigart and Athanasiou 2001). Collagen fibers are radially and circumferentially organized to provide appropriate structure to resist tensile forces. Based on the collagen fiber orientation and thickness, three different layers can be recognized in the inner fibrocartilage region (Fig. 1.4(c)) (Petersen and Tillmann 1998; Sweigart and Athanasiou 2001). Aggrecan is the major PG responsible for the maintenance of viscoelastic properties of the tissue, and its concentration is highest in the inner and middle parts of the meniscus, decreasing towards the periphery. It forms a spatially organized network in contrast to cartilage, where it is more diffusely distributed (Valiyaveettil et al. 2005). Perlecan and SLRPs decorin, biglycan, fibromodulin, and keratocan, and small amounts of adhesion molecules such as fibronectin and thrombospondin are also found in the meniscus (Melrose et al. 2005, 2008). PGs residing particularly at the surface zone are thought to contribute to the smooth

10


frictionless movement of the menisci over the articular surfaces (Melrose et al. 2005, 2008).

1.3.4

Tendon and ligaments

Tendons are specialized dense connective tissue structures connecting bones and muscles, while transmitting forces and allowing joint movements (Kannus 2000). Tenoblasts in the developing and young tendon, and tenocytes, elongated and dispersed fibroblast-like cells in the adult tendon, form about 90±95% of the cells in tendon. The remaining cells are chondrocyte-like cells at regions of pressure and insertion sites, entheses, and synoviocytes on tendon surface, and vascular cells in the endo- and epitenon regions (Benjamin et al. 2006; Cowin and Doty 2007c; Kannus 2000; Riley 2008). The tendon matrix is maintained by tendon cells that are embedded within the long collagen fibrils running parallel to each other and arranged into bundles in a staggered fashion (Fig. 1.5). Type I collagen is the major collagen component of the ECM, but it contains also `minor' collagens and elastin, glycoproteins and adhesive group molecules, e.g., fibronectin, thrombospondin, tenascin C, and undulin (Kannus 2000). Aggrecan and versican are the large PGs providing, together with HA the properties to resist compressive and tensile forces during movements. SLRPs such as fibromodulin, biglycan, and decorin are found to regulate collagen fibrillogenesis, bone morphogenic protein (BMP) activity or stem cell niche organization (Yoon and Halper 2005; Zhang et al. 2005; Bi et al. 2007). Ligaments are dense connective tissue structures connecting articulating bones and giving stability to the joints (Duthon et al. 2006; Cowin and Doty 2007c; Petersen and Zantop 2007). Although they are anatomically distinct from

1.5 Collagen fibril organization in (a) tendon and (b) ligament (adapted from Nordin and Frankel 1989).


11

tendon, they have an overlapping gene expression profile and matrix composition (Rumian et al. 2007). Spindle shaped fibroblasts maintain the ECM that is mainly composed of type I collagen fibers but also contains some other collagens, e.g., type II collagen in the endotendon, type III collagen in the reticular fibers, type IV collagen in the vascular basement membranes, and type VI collagen as a gliding component between functional fibrillar units (Duthon et al. 2006). Parallel organization of collagen fibrils into bundles together with PGs, glyco-conjugates and elastic components results in the formation of a unique, complex elastic network capable of withholding varying multiaxial stresses and tensile strains (Cowin and Doty 2007c). The degree of collagen fibril organization is lower than in tendon (Fig. 1.5) which also has consequences in the biomechanical properties (see below).

1.3.5

Synovial membrane and synovial fluid

Synovial membrane or synovium secretes joint lubricating components into the synovial fluid, nourishes the joints, and removes debris from the synovial space. It is composed of two layers, the intimal layer and the loose connective tissue layer (FitzGerald and Bresnihan 1995; Iwanaga et al. 2000; Sutton et al. 2009). Three basic cell types, type A synoviocytes, type B synoviocytes, and dendritic cells, are found in the intimal layer. Type A synoviocytes, or macrophage-like synoviocytes, are likely to originate from bone marrow and can be considered as resident or tissue macrophages that are mainly phagocytic with large Golgi complex and lysosomes (Iwanaga et al. 2000). Type B synoviocytes, or fibroblast-like synoviocytes, manufacture collagen, fibronectin, HA, and PG 4 (also known as lubricin, megacaryocyte stimulating factor (MSF) and superficial zone protein (SZP)) into synovial fluid to maintain joint lubrication (Rhee et al. 2005; Elsaid et al. 2007; Wann et al. 2009). They differ from other deeper, subintimal fibroblasts in that they contain characteristic lamellar bodies, and produce also surfactant protein A and VCAM-1 (FitzGerald and Bresnihan 1995; Vandenabeele et al. 2003). Dendritic cells form less than 1% of the synovial cells (FitzGerald and Bresnihan 1995). They are potent antigen presenting cells that have a pro-inflammatory role in initiation of the immune responses in rheumatoid arthritis (RA), where they are the effectors of cartilage destruction and a major source for inflammatory cytokine TNF which indirectly induces cartilage collagenolysis (Lutzky et al. 2007). Below the intimal layer there is a loose connective tissue layer that contains fibroblasts, macrophages, adipocytes, mast cells, nerve fibres, vascular endothelial cells, granulocytes, and lymphocytes (FitzGerald and Bresnihan 1995). It is well vascularised, innervated and supplied by lymphatic vessels (Sutton et al. 2009). Synovial fluid is the joint lubricant and shock absorber for AC. Synovial fluid is a blood plasma dialysate, which contains HA and glycoproteins, synthesized

12


by type B synovial lining cells (Fam et al. 2007). HA contributes to the high viscosity and lubricating properties of synovial fluid and is currently used also as a therapy for OA. Recently the synthesis and active secretion of HA were coupled to the movements and use of the joint (Ingram et al. 2008). In addition to substances secreted by the lining cells, synovial fluid contains plasma proteins that originate from the blood vessels vascularizing synovium (Fam et al. 2007). Cellular components are present in small amounts in normal synovial fluid, including different leukocytes: lymphocytes, monocytes, synovial lining cells, and polymorphonuclear cells (Fam et al. 2007). The rate and mechanism of passage of substances going through synovium depend on the size of the molecules. Gases and crystalloids diffuse rapidly in both directions. Larger proteins are taken out of the synovial fluid by way of lymphatics. Macrophages phagocytose cellular debris and particles that are too large to be removed otherwise (Iwanaga et al. 2000). The turnover time for synovial fluid volume is estimated to be about one hour in rabbit and normal human knees, while that for HA is much longer, 17±30 hours in rabbit knee joints (Levick 1987; Ingram et al. 2008). It is now commonly believed that synovial macrophages are responsible for producing the proinflammatory cytokines into the joint space and drive the inflammatory responses with stimulation of cartilage cytokines and matrix degrading proteases under pathological conditions such as OA or RA (Blom et al. 2007; Sutton et al. 2009).

1.3.6

Biomechanical properties of joint tissues

The mechanical properties of tissues can be determined from the load± deformation behavior in compression, tension, bending, or shear geometry. The 3- or 4-point bending tests have actively been used for bone samples and tension tests for tendons and ligaments. For mechanical testing of AC, compression, tension, and shear techniques have traditionally been applied. In compression, unconfined compression, confined compression, and indentation geometries (Fig. 1.6) and stress±relaxation or creep (Fig. 1.7) test protocols are generally in use. To calculate the true material properties of joint tissues, theoretical analysis of the measurements is needed. In most classical models, soft tissues are simplified as homogeneous isotropic linearly elastic materials. The relationship between the stress and strain is described as linear and two independent elastic constants are needed to describe the material, i.e., the elastic (Young's) modulus (E) and Poisson's ratio () (Table 1.1). Consequently, this model is inadequate for characterizing time-dependent mechanical behavior of soft tissues, especially that of AC. However, it has been used actively to calculate the instantaneous (dynamic) or equilibrium (static) modulus for AC (Hayes et al. 1972; Jurvelin et al. 1990).


13

1.6 Schematic presentation of the typical measurement configurations in use for mechanical testing of the articular cartilage. (a) Unconfined compression: the tissue is compressed between two smooth metallic plates allowing fluid flow in the lateral direction. (b) Confined compression: the tissue is placed in a metallic chamber and compressed with a porous filter allowing fluid flow axially through the filter. (c) Indentation: the tissue is compressed with a cylindrical plane-ended or spherical-ended indenter allowing fluid flow in both lateral and axial directions (from Saarakkala 2007).

The joint tissues are all viscoelastic in their mechanical behavior, i.e., the mechanical response, depends significantly on the rate of loading. Depending on the tissue type, the viscoelasticity may originate from the intrinsic property of the solid tissue, or from the interstitial fluid flow within the tissue under load. Under loads with volumetric dilatation, these two viscoelastic mechanisms are difficult to separate. The latter, called poroelasticity, is especially well recognized in AC. A traditional model for AC, taking the interstitial fluid movement into account, is the linear isotropic biphasic model (Mow et al. 1980). In addition to elastic parameters of the solid matrix, knowledge of the tissue permeability is needed for characterizing the time-dependent behavior of the tissue. To extract the model parameters, experimental mechanical measurements are conducted and, subsequently, the theoretical model is fitted to the experimental data. As an extension of the biphasic model, fibril reinforced models have been introduced (Soulhat et al. 1999; Korhonen et al. 2003; Wilson et al. 2004). In these models, the compression±tension nonlinearity is taken into account by inclusion of the collagen fibril network. The material parameters of the fibrilreinforced model are Young's modulus and Poisson's ratio of the drained porous

1.7 (a) In a creep measurement, cartilage tissue deformation (strain) is recorded under a constant load (stress) applied at t0. (b) In a stress-relaxation measurement, the cartilage tissue load (stress) is recorded under a constant deformation (strain) applied at t0 (from Saarakkala 2007).

14


Table 1.1 Basic equations for the determination of isotropic elastic parameters of cartilage Parameter

Equation

Stress ()

dF dA

Strain ()

L0 ÿ L L

Young's modulus (E) (unconfined compression)

E

a a

Poisson's ratio ()

l a

Shear modulus ()

E 21

Aggregate modulus (HA) (confined compression)

HA

Young's modulus (E) (indentation)

E

1 ÿ 2 a 2h

Shear modulus () (indentation)

1 ÿ a 4h

F A L L0 a and a l a h a=h;

1ÿ E 1 1 ÿ 2

reaction force area of the surface in which the force is acting initial thickness thickness after compression/tension axial stress and strain lateral strain indenter radius cartilage thickness theoretical scaling factor due to finite and variable cartilage thickness (Hayes et al.1972; Jurvelin et al.1990)

matrix, permeability, and Young's modulus of the fibril network. Triphasic theory is an extension of the biphasic model incorporating three phases: an incompressible solid, an incompressible fluid, and a monovalent ionic phase (Lai et al. 1991). The model assumes that the total stress of the tissue is composed of the fluid stress, solid stress and chemical potentials. This model can be used to accurately include the effect of cartilage swelling. Owing to tissue heterogeneity and anisotropy, as well as to mimic realistic loading geometries, the model implementations for nonlinear behavior of AC are most often conducted numerically using finite-element analysis. The structure, composition, and properties of all joint components have evolved on the grounds of their biological and mechanical function. In a healthy


15

joint, uncalcified cartilage and meniscus (elastic modulus in compression of 0.1± 1 MPa), calcified cartilage (elastic modulus ~0.3 GPa), and subchondral bone (elastic modulus >1 GPa) establish a structural and functional continuum with optimal mechanical properties (Mente and Lewis 1994; Setton et al. 1999a; Arokoski et al. 2000; Helminen et al. 2000). The tibio-femoral contact stresses may be very high (>10 MPa), compared with typical Young's modulus of 1 MPa for solid matrix of normal cartilage, indicating that hydrostatic pressure within cartilage must serve as a primary mechanism for successful load bearing. Further, intrinsic fluid pressurization contributes significantly to low friction between articulating surfaces (Ateshian 2009). In OA, critical loss of fluid pressurization mechanism of load support takes place. Tendons, with densely packed collagen fibers, show typically very high tensile modulus (>1 GPa) and strength (>100 MPa). Tendons are highly elastic with minor viscoelastic effects and their nonlinear tensile behavior is related to gradual alignment and stretching of the fibers (Ker 2007). Ligaments, owing to lower collagen content and highly woven collagen structure, are less stiff and strong than tendons. Articular cartilage exhibits significant compression±tension nonlinearity. Compressive equilibrium modulus of healthy cartilage in unconfined compression is ~1 MPa; however, under highly dynamic loads hydraulic stiffening produces a modulus that is much higher, and comparable to that of tensile modulus (5±25 MPa, Setton et al. 1999b). Owing to inhomogeneous structure, e.g., depth-dependent increase of PG content, the compressive modulus and permeability increase and decrease, respectively, along cartilage depth (Schinagl et al. 1997; Boschetti et al. 2004). The tensile modulus is highest in the superficial cartilage zone, where the direction of the collagen fibers is parallel to articular surface, and the collagen content is highest. Deeper in the tissue, the more random orientation produces lower tensile stiffness (Kempson et al. 1973). In joint tissues, each structural component and their interactions contribute to overall mechanical characteristics of the tissue. In cartilage, PGs, owing to the swelling stress they produce, and their effect to tissue permeability, are considered important for mechanical characteristics in compression, while collagen is the primary structure resisting tension (Huang et al. 2001). However, joint tissues can be considered to be biological composites, and the structural interactions critically control the mechanical behavior as well. Therefore, these sophisticated structures have remained difficult to replicate using tissue engineering methods, making tissue repair of, for example, AC, challenging. It is well shown that proper collagen cross-linking is essential for a functional matrix in both native and engineered cartilage (Broom 1984; BastiaansenJenniskens et al. 2008). Further, mechanical properties of AC are sensitively modulated by the changes in structural integrity of the tissue (Fig. 1.8).

16


1.8 Minor degenerative changes in matrix, based on the scoring the histological integrity of the cartilage using Mankin score, can lead to inferior mechanical properties (e.g. dynamic modulus) of AC (from Laasanen et al. 2003).

1.4

Resident mesenchymal progenitor cells in synovial joint tissues

Remarkable progress during the last ten years of stem cell research has increased our understanding of how stem cells can be induced and manipulated to form repair tissue. Adult stem cells, especially bone marrow MSCs, which are a highly variable population of multi-potent mesenchymal stem cells and progenitors, have been actively characterized due to their great potential in regenerative medicine (reviewed by, for example, Barry and Murphy 2004; Keating 2006; Caplan 2007; Phinney and Prockop 2007; Abdallah and Kassem 2008, 2009; Chen and Tuan 2008; Jones and McGonagle 2008; NoÈth et al. 2008; Arthur et al. 2009). Currently, MSCs and progenitors have been found residing virtually in all organs and tissues (Sakaguchi et al. 2005; da Silva Meirelles et al. 2006; Chamberlain et al. 2007; El Tamer and Reis 2009). While the aim of this whole book is to gather together current understanding of the normal biology, disease pathogeneses, and different therapeutic approaches of connective tissue disorders, especially those related to joint and associated tissues, it is important also to be aware of the endogenous stem cells and progenitors residing in these tissues. Therefore, inventory of resident stem cells and progenitors in human synovial and associated tissue joints (Table 1.2), and some of their properties will be briefly discussed in this chapter.

1.4.1

Mesenchymal stromal cells

Stem cells have a potential to self-renew, proliferate, and differentiate into multiple cell types. Adult multi-potent stem cells have a more limited differ-


17

entiation potential in comparison to embryonic stem cells. Early embryonic stem cells (derived from the inner layer of blastocyst) are totipotent and can give rise to all germ layers. Pluripotent embryonic stem cells lack differentiation potential to placental cells, but can differentiate to form other tissues. Two major classes of multi-potent stem cells are found in adult bone marrow, hematopoietic stem cells, and nonhematopoietic stromal cells. MSCs are multi-potent nonhematopoietic cells that can differentiate into mesodermal lineages (Fig. 1.9(a)). They represent a small percentage of the total population of nucleated cells in the bone marrow, where the majority of the cells consist of hematopoietic stem cells and hematopoiesis supporting stromal cells (Pittenger et al. 1999). MSCs are defined as highly clonogenic cells having potential for self-renewal and differentiation into multiple mesenchymal tissues (Pittenger et al. 1999). Johnstone was the first to induce chondrogenic differentiation of mesenchymal progenitor cells isolated from rabbit bone marrow (Johnstone et al. 1998). In contrast to the hematopoietic stem cells, no single mesenchymal stem cell specific marker has been found so far. They appear to be a rather heterogeneous population and most of the cells seem to be progenitors rather than true stem cells. To clarify the confusion in the nomenclature and to attempt to standardize the research in this field, the International Society for Cellular Therapy has made two statements (Fig. 1.9(b)). First, they recommend the term `multi-potent mesenchymal stromal cell' instead of the mesenchymal stem cell, the acronym still remaining the same `MSC' for both (Horwitz et al. 2005). Second, multipotent MSCs should fulfill the minimal criteria of expressing certain surface antigens characteristic for mesenchymal cells. In addition, MSCs should not express some hematopoietic and epithelial cell surface antigens, and they should also have a capacity to differentiate under appropriate conditions to chondrogenic, osteogenic, and adipogenic lineages (Dominici et al. 2006). Several controversial issues prevailing in adult stem cell research including nomenclature, and other MSC characteristics, were critically discussed in a recent review by Darwin Prockop (2009).

1.4.2

Role of mesenchymal progenitor cells in joint homeostasis

Being resident in subchondral bone, bone marrow stromal cells may give rise to a spontaneous AC repair that is seen when cartilage lesions extend to the underlying bone, resulting in a formation of cartilage repair tissue. Already in the 1940s, before the era of the modern arthroplasty surgery, abrasion of the osteoarthritic joint surfaces and drilling several holes 6 mm in diameter to the subchondral bone were used as a therapy for OA (Magnuson 1941; Pridie 1959). However, functionally impaired fibrocartilagenous repair tissue did not give satisfactory results. Currently microfracture is a frequently used technique for the repair of AC lesions of the knee. In this `marrow stimulating' technique an

1.9 Definition of adult mesenchymal stem/stromal cell (MSC). (a) Mesenchymal tissues contain MSCs and progenitor cells that under defined conditions have a capacity to differentiate into multiple connective tissue cell types. (b) The minimal criteria for defining the term `multipotent mesenchymal stromal cell' as suggested by the International Society for Cellular Therapy: (1) they must be plastic adherent, (2) express certain cell surface antigens, and (3) have a capacity to differentiate to at least chondrocytic, osteogenic and adipogenic lineages (Horwitz et al. 2005; Dominici et al. 2006) (adapted from SÌÌmÌnen et al. 2008).


19

awl is used to penetrate the subchondral bone to produce small holes in cartilage defects, allowing marrow cells to migrate to the cartilage lesion site (Steadman et al. 2002). Recent studies have shown that microfracture provides effective short-term functional improvement of knee function, but often results in formation of suboptimal fibrocartilage (Knutsen et al. 2007; Mithoefer et al. 2009). The principal role of adult MSCs and progenitors has been considered to maintain physiological balance in the organism by serving a cellular reserve for tissue remodeling and rejuvenation, but they can do more than just respond to stimuli and differentiate. Newly committed progenitor cells have been shown to secrete several growth factors and cytokines (Haynesworth et al. 1996), and immunosuppressive factors, e.g., HLA-G that interfere with the immune recognition system (Selmani et al. 2009; Siegel et al. 2009). Caplan and Dennis (2006) introduced a term `trophic mediator' to MSCs, and defined the trophic effects as `those chemotactic, mitotic, and differentiation-modulating effects, which emanate from cells as bioactive factors that exert their effects primarily on neighboring cells and whose effects never result in differentiation of the producer cell'. Stem cells are maintained in so-called stem cell niches at specific sites in the tissues (Gregory et al. 2005; Jones and Wagers 2008; Walker et al. 2009). Stem cell niches have been characterized in tendon, AC, and zone of Ranvier, where PGs or their GAG sulfation patterns have been suggested as having important roles in maintaining and organizing the niches, and regulating the local BMP activity (Bi et al. 2007; Hayes et al. 2008; Karlsson et al. 2009). Hence, there seems to be a complex and bidirectional regulation system of the stem cell response to stimuli for differentiation and secretion of bioactive factors, thereby influencing tissue homeostasis (Fig. 1.10) (Caplan and Dennis 2006; Caplan 2009).

1.4.3

Articular cartilage

Articular cartilage was earlier thought to lack stem cells or progenitors but several recent studies have demonstrated their existence in the tissue (Table 1.2). A few years ago it was first shown that young bovine AC contains a multi-potent progenitor cell population in the superficial zone with differentiation plasticity into various connective tissues, including bone, tendon, and perimysium (Dowthwaite et al. 2004). Several studies have shown the presence of multipotent progenitors with limited expandability in AC of healthy young individuals (reviewed by Tallheden et al. 2006). In OA cartilage, increased number of cells with MSC phenotype has been found (Alsalameh et al. 2004; Fickert et al. 2004; Hiraoka et al. 2006). Adult human AC contained a small population of cells that coexpressed surface antigens CD105 and CD166 (ALCAM, activated leukocyte adhesion molecule) (Alsalameh et al. 2004). These markers have been proposed to define a population of MSCs in bone

Table 1.2 Studies demonstrating presence of multipotent progenitor cells in human synovial joint and associated tissues Reference

Multipotency1

Articular cartilage Barbero et al. (2003)

ACO

Fickert et al. (2004) Alsalameh et al. (2004)

ACO ACO

Koelling et al. (2009)

ACO

Thornemo et al. (2005)

ACO

Synovial membrane and synovial fluid De Bari et al. (2001) ACO Vandenabeele et al. (2003) ± Sakaguchi et al. (2005)

ACO

Mochizuki et al. (2006)

ACO

Jones et al. (2008)

ACO

Morito et al. (2008)

ACO

Notes2 Plasticity of dedifferentiated clonal chondrocytes was tested. TGF -1, FGF-2 and plateletderived growth factor BB enhanced C and O but reduced A differentiation capacity. (AC) Osteoarthritic cartilage contained 5% of CD9+/CD90+/CD166+ multi-potent MSCs. (AC) 3.49% of CD105+/CD166+ MSCs in normal articular cartilage and frequency is increased in osteoarthritic cartilage. (AC) Migratory chondrogenic progenitors (type 2 cells with fibroblast like morphology) were found in osteoarthritic cartilage. (AC) Osteoarthritic cartilage contained multi-potent progenitors (3.6% of all cells). (AC) Single cell derived multi-potent MSC clones were isolated from knee joint SM. (AD) Morphologically SF MSCs resembled type B synoviocytes why they likely originate from synovial lining. (AD) MSCs derived from several tissues were compared; synovial MSCs were superior in chondrogenesis. (SM AD PE BM MU) Osteogenic and chondrogenic potential was highest in the synovium-derived populations (SM AS AD) SF MSC prevalence increased 7-fold in early OA. MSCs likely originated from synovium. Chondrogenic potential was more consistent in SF than BM MSCs. (SF BM) SF MSC prevalence increased 100-fold after ACL injury and they aligned to the rupture site. (SF SM BM)

Ligaments and tendon Huang et al. (2008)

ACO

Cheng et al. (2009)

ACO

de Mos et al. (2007) Bi et al. (2007)

ACO ACO

Intra-articular fat pads English et al. (2007)

ACO

Khan et al. (2008)

C

Wickham et al. (2003)

ACO

Other De Bari et al. (2006) Williams et al. (1999) Zheng et al. (2007) Segawa et al. (2009)

ACMO ACMO CMO ±

1

ACL and total knee replacement surgery samples. Variation in tripotency and differentiation and proliferation rate between patients. (ACL) MSCs were isolated from cruciate ligaments and BM. Phenotype was similar in all MSC populations. (ACL PCL BM) Hamstring tendon contained cells with intrinsic differentiation potential. (TE) Tendon MSCs are maintained in a fibromodulin and biglycan modulated niche. They differentiated to form entheses-like tissue. (TE) Hoffa's fat pad contained MSCs that maintained chondrogenic phenotype long time. (AC AD BM) Chondrogenic cells were isolated from infrapatellar fat pad, and expansion in FGF-2 enhanced chondrogenic potential. (AD) Infrapatellar fat pad contained multi-potent MSCs. (AD) Periosteum contained multi-potent MSCs. (PE) Skeletal muscle contained multi-potent mesenchymal progenitor cells. (MU) Myoendothelial cells isolated from skeletal muscle were multi-potent. (MU) Gene expression profiles of intra-articular tissue colony forming MSCs (SM, ME, LI) were closer to each other and articular chondrocytes than to extra-articular tissue MSCs (BM, AD, MU). PRELP was a characteristic highly expressed gene among intra-articular tissue MSCs. (SM ME LI MU AD BM)

Tested differentiation capacity to adipogenic (A), chondrogenic (C), osteogenic (O), or myogenic (M) lineages. Cell source(s) used for MSC isolation are in parentheses: articular cartilage (AC), synovial membrane (SM), synovial fluid (SF), adipose synovium (AS), adipose tissue (AD), anterior cruciate ligament (ACL), posterior cruciate ligament (PCL), medial collateral ligament (MCL), tendon (TE), meniscus (ME), skeletal muscle (MU), periosteum (PE).

2

22


1.10 Mesenchymal stem cells are maintained in stem cell niches and function as trophic mediators and reservoir for tissue remodeling and repair. Proteoglycans, particularly glycosaminoglycan sulfation patterns of aggrecan and perlecan in cartilage (Hayes et al. 2008), and small leucine rich repeat proteoglycans (SLRPs) fibromodulin (Fmn) and biglycan (Bgn) in tendon (Bi et al. 2007), have been suggested to have important functions in regulating the organization and/or growth factor presentation in the niches (principle of the graph adapted from Caplan and Dennis 2006).

marrow stroma and have properties similar to mesenchymal progenitor cells (Majumdar et al. 1998). As the presence of CD105+/CD166+ progenitor cells was significantly increased in OA cartilage, they were speculated to have a role in pathogenesis of OA (Alsalameh et al. 2004). As CD105 is endoglin, a TGF receptor III, its expression in MSC population is likely to enhance the chondrogenic potential due to responsiveness to exogenous TGF , which is used for induction of chondrogenesis. Fate and differentiation regulating factor Notch1 has been regarded as a progenitor marker. Surprisingly high numbers of Notch1 positive cells have been found both in healthy and OA cartilage. Over 70% of the cells in primary culture of cells isolated from AC with induction in OA were Notch1 positive (Hiraoka et al. 2006). In normal human AC, taking all cartilage zones together, over 45% of the cells expressed progenitor markers Notch1, a stem cell marker Stro-1 and vascular endothelial molecule VCAM-1, with the highest expression in the superficial zone (Grogan et al. 2009). Most of the cells in chondrocyte clusters in OA cartilage also expressed all these progenitor markers, suggesting responses to OA pathogenesis. In another study with adult bovine knee AC, Notch1 expression did not correlate with multi-potent properties of the progenitors, thus questioning the value of Notch1 as an early progenitor marker, and suggesting that the actual progenitor cell population is much smaller in adult AC (Karlsson et al. 2008). Supporting this, normal and OA cartilage contained 0.14% so-called side-population (SP) cells identified by their negative staining for Hoechst 33342 dye, that differentiated into chondrocytes and osteocytes, but


23

not adipocytes, thus likely representing a more primitive osteoprogenitor population than Notch1, Stro-1, and VCAM-1 expressing cells (Grogan et al. 2009). Recently, migratory chondrogenic progenitor cells from fibrocartilagenous repair tissue were identified during the later stages of human OA (Khan et al. 2009; Koelling et al. 2009). These cells resemble type 2 cells with a secretory phenotype (see `Chondrocytes' in Section 1.3.1), originally described by Kouri et al. (1996) in OA cartilage. Clonogenic cells were isolated after they migrated out from the cartilage explants onto the plastic. Amazingly, these cells also were able to migrate into the deeper zones of the OA cartilage explant from the surface in tissue culture, as tracked by GFP marker gene. Morphologically similar cells were identified migrating through breaks in the tidemark of OA cartilage by electron microscopic studies. These cells expressed transcription factors Sox9 and runx2, differentiated into osteoblasts and adipocytes, and their chondrogenic potential was enhanced after downregulation of runx2, suggesting that they are derived from the osteoprogenitor lineage (Koelling et al. 2009). Although historically AC has been considered to have a poor intrinsic repair capacity, and progenitor cells appear to be able to spontaneously induce only improper fibrocartilagenous repair tissue, their presence in cartilage opens up novel possibilities for the future developments of cartilage repair in the late stages of OA.

1.4.4

Synovium

Synovial fibroblast-like cells were isolated from adult human synovial membrane (De Bari et al. 2001; Vandenabeele et al. 2003). Their progenitor nature was studied by five independent clones that were all capable of chondrogenic, osteogenic, and adipogenic differentiation. These cells contained specific lamellar bodies, and expressed surfactant protein A, both characteristic to type B synoviocytes, thus suggesting that they may originate from synovial lining. Distinct expression profiles between MSCs derived from intra-articular tissues (synovium, meniscus, and ligament) and extra-articular tissues (muscle, adipose tissue, and bone marrow) were observed in patient matched analysis of 47 000 human transcripts (Segawa et al. 2009). Intra-articular tissue MSCs and articular chondrocytes expressed significantly more PRELP, ECRG4 and OGN while extra-articular tissues expressed higher levels of DSP, NRG1, SERPINB2, LFNG, NOG, and NEF3. Comparison of the synovium-derived progenitor cells with those derived from bone marrow, periosteum, skeletal muscle, and subcutaneous adipose tissue in patient matched studies have indicated their superiority in chondrogenic differentiation (Sakaguchi et al. 2005). Synovial MSCs colony forming unit (CFU) was 100-fold of that in the bone marrow (Morito et al. 2008). Their superiority in proliferation rate and chondrogenic

24


differentiation was also supported by in vivo studies where MSCs from bone marrow, adipose tissue, and muscle were transplanted into cartilage defects together with periosteal patch in rabbits (Koga et al. 2008). Also, Fan et al. (2009) summarizes the superiority of chondrogenic properties of synovial MSCs over bone marrow MSCs, including higher expression of hyaluronan receptor CD44 and UDPGD, an enzyme vital for hyaluronan synthesis, and they also already express low levels of cartilage genes of COMP, aggrecan, type IX and XI collagen, and have a higher proliferation capacity than MSCs from other tissues. Synovium also originates from a common pool of progenitors with cartilage (Archer et al. 2003). These characteristics are advantageous in clinical applications, including that synovial membrane also easily self-regenerates, thereby allowing biopsies to obtain cells for autologous transplantation. Their high tendency to produce fibrocartilagenous tissue rather than hyaline cartilage remains yet a challenge to be overcome.

1.4.5

Synovial fluid

Jones et al. (2004) identified a presence of MSC population in synovial fluid of OA patients. Later they showed that also normal synovial fluid has a resident MSC population that increases at early OA (Jones et al. 2008). These cells are highly clonogenic and multi-potent both in young bovine and normal human joints. Synovial fluid cells represent a more homogenous pool in comparison with bone marrow MSCs. They were highly clonogenic cells with consistent chondrogenic capacity and were less adipogenic than bone marrow MSCs. Synovial fluid cells expressed higher levels of CD44 (hyaluronan receptor, a putative marker of enhanced chondrogenesis) than bone marrow MSCs, and lacked expression of CD271 (a low-affinity nerve factor receptor, characteristic marker of human bone marrow MSCs). Thus they are likely to originate from synovium (dislodged synovial fragments) or from the superficial AC layer rather than from bone marrow via circulation. Intra-articular ligament injury induced increased prevalence of synovial fluid MSCs and their adhesion onto the injured ligament (Morito et al. 2008). Also these cells were suggested to originate from synovium, as they were more similar to synovial MSCs than those derived from bone marrow, as compared by their morphologic and gene expression features. Reasons for synovial fluid MSC increase during ligament injury may be due to vessel injury-related promotion of cytokines and chemokines or to inflammation as was suggested by Jones et al. (2008).

1.4.6

Tendon and ligaments

Salingcarnboriboon et al. (2003) established mouse tendon-derived cell lines exhibiting pluripotent stem cell-like property with differentiation potential to


25

osteogenic, chondrogenic and adipogenic lineages, but these cells were not characterized for their surface antigens. Bi and collegues (2007) identified a stem/progenitor cell population with universal stem cell characteristics in mouse and human tendon. These cells resided in a unique niche where small PGs biglycan and fibromodulin were found to be critical components in organizing this niche and regulating the local BMP activity. Multi-potent MSCs have also been isolated from ligaments showing diversity in the differentiation potential between six independent clones (Huang et al. 2008). Only one clone out of six was tripotent, and when compared with bone marrow MSCs, anterior cruciate ligament (ACL) derived clones expressed more type I and III collagens, suggesting higher potential for ligament fibroblasts. Tripotent MSC populations with typical MSC properties and similar phenotype to bone marrow MSCs were also isolated and expanded from ACL and posterior cruciate ligament (PCL) (Cheng et al. 2009).

1.4.7

Meniscus

In a comparative patient matched study, progenitor cells from several intraarticular and extra-articular tissues, including menisci, were isolated and their differentiation potential and other MSC properties compared (Segawa et al. 2009). This study revealed distinct gene expression profiles between these tissue groups, thus suggesting that intra-articular sources may be more favourable for chondrogenic differentiation and cartilage repair than MSCs from extra-articular tissues. The existence of multi-potent MSCs in meniscus tissue has thus been presented, but to our knowledge, their properties or possible role in intrinsic healing have not been studied in detail.

1.4.8

Other tissues

In the periosteum, the cambium layer contains multi-potent progenitor cell population that readily differentiates into chondrogenic lineage (reviewed by O'Driscoll and Fitzsimmons 2001). The advantage of this differentiation has been used for a long time in surgical repair procedures covering cartilage lesions by periosteal flaps (Jaroma and RitsilaÈ 1987; O'Driscoll 1998). Although the method shows short-term benefit, the repair tissue degenerates in the follow-up and at present periosteal transplantation is not a recommended method for cartilage repair (Hoikka et al. 1990; Hunziker 2002). Basic research shows still progress. A recent study in a goat model with human periosteumderived progenitor cells showed highly clonogenic cells with differentiation potential into chondrogenic, osteogenic, adipogenic, and myogenic lineages (De Bari et al. 2006). In another study, a simple isolation technique for skeletal tissue repair purposes of periosteal multi-potential MSCs was described (Choi et al. 2008).

26


Intra-articular infrapatellar adipose tissue, i.e., Hoffa's fat pad was shown to contain highly clonogenic, multi-potential MSCs that were capable of maintaining chondrogenesis for a long time (Wickham et al. 2003; English et al. 2007). Also, the infrapatellar fat pad contained pericyte marker 3G5-positive clonogenic cells that expressed MSC markers with enhanced chondrogenic differentiation after expansion in presence of FGF-2 (Khan et al. 2008). As reviewed by El Tamer and Reis (2009) and Tapp et al. (2009), MSCs isolated from adipose tissue are multi-potent, but they differ from MSCs derived from other mesenchymal tissues in that they endogenously express lower levels of BMP2, -4, and -6 in comparison to bone marrow. Their chondrogenic potential can be significantly increased in vitro by supplementing the cultures with exogenous BMP in addition to TGF (Hennig et al. 2007). Skeletal muscle tissue contains two stem cell populations with a possible MSC character: satellite cells and muscle-derived stem cells. Satellite cells have been considered as precursor cells rather than stem cells and have been shown so far to have osteogenic properties (Hashimoto et al. 2008; Sun et al. 2008). Muscle-derived stem cells likely locate in the connective tissue regions of skeletal muscle or in the capillaries surrounding myofibers. Great variation has been shown to exist between different populations greatly depending on the isolation methods (O'Brien et al. 2002). Animal muscle-derived stem cells have been shown to differentiate at least into osteblasts, myofibroblasts, chondrocytes and hematopoietic lineages, in addition to myoblasts (Cao and Huard 2004; Usas and Huard 2007; El Tamer and Reis 2009; Kubo et al. 2009). A myoendothelial cell population expressing myogenic and endothelial markers was recently isolated from human muscle with potential to differentiate into myogenic, osteogenic, and chondrogenic lineages (Zheng et al. 2007).

1.5

Conclusions and future trends

In this chapter we have described the macromolecular structure of the knee joint, organization of the tissues, their cellular and extracellular matrix composition and interrelationships with the biomechanical properties of the tissues. The major emphasis has been AC. Also we have reviewed the current knowledge of the presence of MSCs and progenitor cells in human joint tissues and briefly discussed their possible input in the joint homeostasis and endogenous repair potential. Inhomogeneous structure, anisotropic and nonlinear mechanical properties are characteristic of joint tissues. In functional adaptation, the structure, composition, and properties of the joint tissues are conditioned to withstand the loads which they are exposed to. In healthy joints, this will lead to optimized joint function. In the knee joint, structure, composition, and mechanical properties of AC show significant topographical variation. The topographical variation in mechanical properties has been revealed also by in vivo indentation


27

measurements (Lyyra et al. 1999). Compression-tension non-linearity of AC is a further indication of tissue adaptation to the mechanical environment. Changes in the mechanical properties of joint tissues may indicate early pathology. In OA, equilibrium stiffness of cartilage can typically decrease by 50%. Minor degenerative changes in matrix, based on the scoring the histological integrity of the cartilage, have been shown to lead to extensive impairments in mechanical competence of AC (Laasanen et al. 2003). This has led to development of mechanical instrumentation for early in vivo diagnostics of cartilage degeneration. Compared with classical diagnostic techniques, these methods may provide minimally invasive ways for more sensitive diagnostics. Some techniques, e.g. quantitative ultrasound, may provide a method for simultaneous diagnostics of both AC and subchondral bone (Saarakkala et al. 2006). Potentially, small changes in tissue structure and composition may manifest themselves as more significant changes in tissue mechanical properties. This is an idea behind the concept of functional imaging (Julkunen et al. 2008). A combination of quantitative non-invasive imaging, such as magnetic resonance imaging (MRI), combined with the realistic theoretical model of joint function, helps to diagnose early tissue pathology and may enable functional analysis and prediction of the development of tissue properties in the future. This would be highly useful, for example, when assessing the outcome of cartilage repair operations. All joint tissues contain intrinsic repair potential in the form of resident tissue specific multi-potent progenitor cell populations. The presence of MSCs in the synovial fluid but also in the other tissues may have important function in the homeostasis of the joint, and particularly during traumatic or inflammatory conditions. The influence of stem cells in the tissues and synovial fluid may not be restricted to the direct effect during tissue remodeling or repair but they may also act as trophic mediators to assist these processes by secreting bioactive factors such as cytokines or factors that suppress immune recognition mechanisms (Caplan and Dennis 2006; Caplan 2009). Differences in the cell surface antigen presentation, gene expression profiles, in their effectivity to expand and self-renew in the culture, and potential to differentiate into different mesenchymal tissues are seen between MSCs derived from intra-articular and extra-articular tissues. There are differences also within the cell populations or single cell-derived clones from the same tissues, indicating the heterogeneous nature of these progenitor populations. MSC populations derived from intra-articular tissue are more homogenous than those from extra-articular sources, particularly from bone marrow. Synovial MSCs are superior over the others in their chondrogenic differentiation potential but they appear to favor development of fibrocartilagenous phenotype under conditions studied so far. Unfortunately, spontaneous repair by intrinsic MSCs seldom results in a formation of biomechanically adequate repair tissue. MSCs present in synovial fluid are increased in ligament injury and align with injured

28


tissue for regeneration, but perhaps unfavourable biomechanical and other factors prevent full healing (Strand et al. 2005; Morito et al. 2008). Further, chondrogenic progenitors are present in adult AC. This questions the prevailing opinion that the essential cause for poor repair capacity of AC is the shortage of intrinsic chondrogenic cells. It remains a challenge for future regenerative medicine to learn how to trigger intrinsic repair potentials of joint tissues, especially that of MSCs by taking advantage of the progenitor cell migration through tissues and homing to the repair site (Chamberlain et al. 2007; Koelling et al. 2009) and to induce synthesis of hyaline cartilage, instead of fibrocartilage.

1.6

Sources of further information and advice

1.6.1

Useful links

· Gray's Anatomy http://education.yahoo.com/reference/gray/subjects/ · Wheeless' Textbook of Orthopaedics: http://www.wheelessonline.com/ · Stem cell links http://stemcells.nih.gov/info/basics http://learn.genetics.utah.edu/content/tech/stemcells/

1.6.2

Books

Bronner F II and Farach-Carson M C (2007), Topics in Bone Biology, London, Springer. Cowin S C and Doty S B (2007), Tissue Mechanics, New York, Springer. JoÂzsa L G and Kannus P (1997), Human Tendons, Anatomy, Physiology, and Pathology, Champaign, IL, Human Kinetics. Nordin M and Frankel V H, (2001), Basic Biomechanics of the Musculoskeletal System, 3rd ed., Philadelphia, Lippincott Williams & Wilkins.

1.6.3

Book chapters

Bartl R and Frisch B (2009), `Biology of bone' in Bartl R and Frisch B, Osteoporosis, Diagnosis, Prevention, Therapy, Springer, Berlin, 7±28. Sandell L, HeinegaÊrd D and Hering T M (2007), `Cell biology, biochemistry, and molecular biology of articular cartilage in osteoarthritis' in Moskowitz R W, Altman R D, Hochberg M C, Buckwalter J A and Goldberg V M, Osteoarthritis. Diagnosis and medical/surgical management, 4th ed., Philadelphia, Lippincott Williams & Wilkins, 73±106. Zamorani M P and Valle M (2007), `Bone and joint' in Bianchi S, Martinoli C, Medical Radiology. Ultrasound of the Musculoskeletal System. Berlin, Heidelberg, New York, Springer, 137±85.


1.7

29

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The myofibroblast in connective tissue repair and regeneration B . H I N Z , University of Toronto, Canada

Abstract: Myofibroblasts contribute to the normal healing of connective tissues such as skin, bone and cartilage and assure tissue integrity by forming a mechanically resisting scar after injury of heart, lung, liver and kidney. Excessive extracellular matrix (ECM) secreting and contractile activities of the myofibroblast result in the development of fibrosis that dramatically impedes normal tissue function. Just like many other cell types, mesenchymal stem cells which are implanted to regenerate tissue are prone to this transformation and are in danger of generating scar instead of functional tissue. This chapter will define the characteristics of the myofibroblast with the ultimate aim of controlling its activity in tissue repair and regeneration. Keywords: -smooth muscle actin, fibrosis, stroma reaction to tumour, transforming growth factor beta (TGF 1), wound healing.

2.1

Introduction

When severely damaged tissues cannot be regenerated by the routine repair mechanism of the body or when physiological healing is imperfect, then regenerative medicine and tissue engineering are considered. Most cell therapeutic applications involve the isolation of autologous regenerative cells that are then expanded in culture and re-implanted with the aim of restoring organ function. Depending on the type and the structure of tissue to repair, different cell delivery strategies have been developed, ranging from direct injection into the damaged tissue (site-directed delivery), infusion by the intravenous and left ventricular pathways (systemic delivery), and delivery in various scaffolds (Caplan, 2007; Clark et al., 2007; Giordano et al., 2007; Karp and Langer, 2007; Lutolf and Hubbell, 2005; Pittenger and Martin, 2004; Robertson et al., 2008; Sands and Mooney, 2007; Segers and Lee, 2008; Weaver and Garry, 2008). In addition to choosing the appropriate delivery strategy, the success of regenerative medicine strongly depends on selecting the right cell type for implantation. The perfect regenerative cell must be able to replicate in culture for rapid cell population expansion and must exhibit (or develop) phenotypic features that are suitable to restore the function of the organ to repair. The `right cell' is not necessarily one that exhibits the required features in the preparative cell culture but the one that

40


develops and more importantly keeps these features in the body implant or graft and that attains/retains these features in the body. This is the basic dilemma of stem cell therapy: pluripotency is wanted because it enables stem cells to finally attain the characteristics of cells that construct the normal organ. However, when delivered for organ regeneration, pluripotent cells do not experience the normal organ but the hostile diseased environment which can drive their differentiation toward unwanted and even destructive phenotypes. If regenerative strategies fail, the outcome can be teratoma formation (implanted cells continue to proliferate and become tumorigenic) (Hentze et al., 2007; Rice and Scolding, 2008) or tissue contracture and deformation (implanted cells become fibrogenic). To better understand the challenges that regenerative cells face after engrafting injured organs we have to consider the body's inherent reparative machinery and the cells involved in physiological tissue repair. When organs and tissues are damaged such as after cardiac infarct, skin wounding or bone fracture the inherent repair mechanisms of our body have to fulfil two urgent tasks: (1) establishing tissue haemostasis, fighting inflammation and discarding debris which is carried out by immune and inflammatory cells and (2) providing mechanical tissue coherence by forming a scar which is the task of fibroblasts and so-called myofibroblasts (Baudino et al., 2006; Brown et al., 2005; Desmouliere et al., 2005; Gurtner et al., 2008; Werner and Grose, 2003). The myofibroblast is the most prominent cell phenotype, generated to populate and to repair injured tissues by secreting extracellular matrix (ECM) and organizing this ECM in a contractile process (Desmouliere et al., 2005; Hinz et al., 2007; Tomasek et al., 2002). Myofibroblasts severely impair organ function when their contractile and ECM protein secretory activities become deregulated as is the case in virtually all fibrotic diseases. After briefly summarizing the characteristic features and functions of the myofibroblast I will ask, and at least partly answer, the question of the myofibroblast origin. It becomes increasingly clear that myofibroblasts can arise from a plethora of precursor cells predominantly of mesenchymal but also from ectodermal origin (Hinz et al., 2007). It appears that damaged tissues can recruit myofibroblast precursors from several sources to satisfy the temporarily high demand of cells with tissue remodelling and repair activity. Most clinical approaches implanting differentiated or pluripotent stem cells or introducing biomaterials to engineer damaged and fibrotic tissue have to cope with the special cellular, chemical and mechanical environment created by the myofibroblast. Worse, regenerative cells themselves are at high risk of attaining the fibrogenic myofibroblast character being subjected to the `bad' neighbourhood of the scar. As a consequence, they can switch sides and become the enemies of regenerative medicine and tissue engineering by creating fibrotic scar tissue as opposed to the aim of restoring organ function. To prevent this development it is crucial to understand the general molecular pathways regulating evolution and function of the myofibroblast.

The myofibroblast in connective tissue repair and regeneration

2.2

41

Myofibroblasts: humble tissue construction workers

One has to bear in mind that the aim of the body response to injury, in contrast to regenerative medicine and tissue engineering, is to provide rapid repair even at the expense of losing tissue functionality. Indeed, a considerable level of scarring and fibrosis is required to preserve the mechanical stability of an injured organ against rupture. I will here concentrate on the processes that are involved in providing this mechanical stability and shed light on the major cell phenotype involved, the myofibroblast. The forces leading to remodelling and contraction of damaged tissues are generated within the wounded tissue itself. In the early 1970s Gabbiani and coworkers (1971) identified specialized fibroblasts as the active component in dermal wound contraction, which were named myofibroblasts to account for their ultrastructural similarity to smooth muscle cells (SMC) (Gabbiani et al., 1971). From a historical perspective it is important to point out that at the time of their discovery, the definition of `myofibroblast' was exclusively based on the co-existence of mesenchymal morphological features including a developed endoplasmic reticulum as well as SMC features such as actin filament bundles and contractile activity. Specific molecular markers had only been defined about ten years later. Further morphological features of the myofibroblast are high ECM synthesizing activity, development of cell-to-cell and cell-to-matrix adhesions (fibronexus), and secretion of growth factors (for reviews see Eyden, 2008; Hinz, 2007). Over the last three decades myofibroblasts have been found in a variety of physiological and pathological situations that are characterized by enhanced remodelling and tension production. Myofibroblasts can be of very heterogeneous origins as summarized in Section 2.4; however their development follows a well-established sequence of events. De novo myofibroblast differentiation in response to tissue injury is initiated by changes in the composition, organization and the mechanical properties of the ECM (Hinz and Gabbiani, 2003) and by various cytokines that are released by inflammatory and resident cells (Gurtner et al., 2008; Werner and Grose, 2003). The progress of myofibroblast differentiation can be separated into two main phases that are each characterized by specific cytoskeletal characteristics (Fig. 2.1). First, to re-populate cell-denuded damaged tissue, myofibroblast precursor cells acquire contractile bundles. These in vivo stress fibres generate sufficient forces to pull the cells forward during the migration process and to pre-remodel the ECM (Hinz et al., 2001b). To discriminate such activated and low contractile cells from quiescent fibroblastic cells which are devoid of any contractile features, the term `proto-myofibroblast' was proposed (Tomasek et al., 2002). It has to be noted that in standard culture most fibroblastic cells attain this phenotype by developing stress fibres composed of cytoplasmic actins on the rigid culture plastic surface that represents a considerable mechanical stimulus.

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2.1 The myofibroblast differentiation spectrum. Co-immunostaining for Factin (phalloidin, all lower image parts) and for -SMA (all upper image parts) discriminates between fibroblasts (left), proto- (middle) and differentiated myofibroblasts (right column). All three phenotypes occur sequentially in the granulation tissue of healing rat open wounds (Hinz et al., 2001b), and during the maturation (stiffening) of mechanically restrained collagen gels (Hinz, 2006). Myofibroblasts cultured on silicone culture substrates (elastic modulus indicated in kPa) fully differentiate on stiff, are proto-myofibroblasts on medium stiff and fibroblasts on very soft polymers. Similar control over myofibroblast differentiation is achieved by growing them on microcontactprinted (CP) arrays of very small (1 m length), medium long (4 m) and super-sized (20 m) adhesion islets (Goffin et al., 2006).


43

Mechanical stress is also prerequisite for the second stage of myofibroblast differentiation hallmarked by de novo expression of the actin isoform -smooth muscle actin (-SMA) and its incorporation into pre-existing stress fibres (Clement et al., 2005; Goffin et al., 2006; Tomasek et al., 2002). Expression of -SMA discriminates differentiated myofibroblasts from proto-myofibroblasts (Fig. 2.1). Indeed, neo-expression of -SMA by fibroblastic cells is the most widely used criterion to define the differentiated myofibroblast and to diagnose myofibroblast-related diseases. Besides serving as molecular marker, incorporation of -SMA into stress fibres significantly augments the contractile activity of fibroblastic cells and hallmarks the contraction phase of connective tissue remodelling (Hinz et al., 2001a). As elaborated in greater detail in Section 2.6, expression of -SMA is precisely controlled by the joint action of growth factors like transforming growth factor beta 1 (TGF 1), of specialized ECM proteins such as the extra domain A (ED-A) containing fibronectin (FN) splice variant ED-A FN and of the mechanical microenvironment (Tomasek et al., 2002). Under physiological conditions the contractile and secretory activities of myofibroblasts are terminated when the tissue is sufficiently remodelled and repaired. Under most circumstances `sufficient repair' signifies that the damaged tissue regains mechanical coherence but does not necessarily mean the restoration of functionality. At this point -SMA expression becomes down-regulated and myofibroblasts disappear by massive apoptosis, leaving the mature scar behind (Desmouliere et al., 1995). However, in excessive repair myofibroblast activity persists and results in tissue deformation by contracture. This is particularly evident in hypertrophic scars such as those developing after burns (Atiyeh et al., 2005), in scleroderma (Strehlow and Korn, 1998; Varga and Abraham, 2007) and in the palmar fibromatosis of Dupuytren's disease (Tomasek et al., 1999). Myofibroblast-generated contractures are also characteristic for fibrosis affecting vital organs such as liver (Desmouliere et al., 2003; Gressner and Weiskirchen, 2006), heart (Baudino et al., 2006; Brown et al., 2005; Virag and Murry, 2003), lung (Chiappara et al., 2001; Phan, 2002; Thannickal et al., 2004) and kidney (Lan, 2003). Cells with myofibroblastic phenotype further contribute to the development of atheromatous plaques after blood vessel injury (Bochaton-Piallat and Gabbiani, 2006). A number of different biomaterials have been shown to activate macrophages, which in turn contribute to the generation of myofibroblast by producing TGF 1 (Anderson et al., 2008; Li et al., 2007a). Myofibroblasts are further considered to be key elements in creating circumferential tissue constrictions that form around solid body implants (Comut et al., 2000; Suska et al., 2008) and contract silicone breast implants (Coleman et al., 1993; Rudolph et al., 1978; Siggelkow et al., 2003). Finally, myofibroblasts play a role in the process called stroma reaction to epithelial tumours which promotes cancer progression by creating a stimulating microenvironment for the transformed cells (Bhowmick and Moses, 2005; De Wever and Mareel, 2003; Desmouliere et al., 2004). This tumour-

44


promoting feature has raised the concept that myofibroblasts as one subpopulation of tumour-associated fibroblasts represent important targets of anticancer treatments (Albini and Sporn, 2007; Mueller and Fusenig, 2004). Given the diversity of organs that can host myofibroblasts it is intuitive to ask where these cells come from ± often with the aim of preventing their formation and destructive accumulation.

2.3

Know thy enemy: a quick guide to identify the myofibroblast

The original and most basic myofibroblast definition describes a cell of mesenchymal character joining fibroblastic and SMC features, which exhibits pronounced actin±myosin filament bundles and contractile activity. Cells missing these features cannot be functional myofibroblasts. On the other hand, no molecular marker has been described that unmistakably identifies a cell as being myofibroblast. One way to approach the problem of its identification is to consider that myofibroblasts are rarely present in normal tissue and predominantly arise in pathological and physiological repair processes. Under most circumstances, myofibroblasts differentiate from other cell types by de novo expressing stress fibres and -SMA (the exception of SMC will be discussed below) and can retain features of their precursors (Section 2.4). A warrant of typical proteins expressed by differentiated myofibroblasts and characteristic morphological features has been summarized elsewhere (Eyden, 2007, 2008; Hinz, 2007; Schurch et al., 2007); here I want to shed light from a slightly different angle: against which cell type do you want to discriminate the myofibroblast? Distinction from epithelial cells is straightforward by assessing expression of mesenchymal markers that are not expressed in epithelium, in particular the cytoskeletal protein vimentin. Generally, -SMA can be used as discriminator with the exception of myoepithelial cells that are also -SMA-positive. Myoepithelium however is negative for vimentin and in contrast to myofibroblasts expresses keratins, E-cadherin and desmoplakin (Bissell and Radisky, 2001; Lazard et al., 1993; Savera and Zarbo, 2004). Another special case is transformed epithelium that can lose epithelial characteristics and acquires mesenchymal features during epithelial-to-mesenchymal transition (EMT) (Kalluri and Zeisberg, 2006). The fraction of these cancer-derived cells that further differentiate to express -SMA are functionally and morphologically myofibroblasts. Once one has defined the criteria for the fibroblast (which is a daunting task in itself) the detection of -SMA-positive stress fibres suffices to distinguish fibroblasts from differentiated myofibroblasts. The same criterion can apply to sort myofibroblasts from other normal connective tissue cells, including chondrocytes and osteoblasts (Spector, 2001) as well as endothelial cells that all express vimentin. In addition, differentiated myofibroblasts in vivo and in


45

vitro can de novo express the cell±cell contact protein OB-cadherin that has not yet been reported on the surface of -SMA-negative fibroblasts (Hinz et al., 2004). In strict terms and according to the original definition of the myofibroblast, the development of -SMA-negative contractile stress fibres alone is a hallmark of myofibroblast differentiation because normal fibroblasts are noncontractile (Gabbiani et al., 1971). If both phenotypes co-exist in vivo, such as during dermal wound healing (Hinz et al., 2001b) and in lung alveolar septa in the context of lung fibrosis (Kapanci et al., 1990), `proto-myofibroblast' (SMA-negative stress fibres) and `differentiated myofibroblast' (-SMA-positive stress fibres) are the appropriate terms (Tomasek et al., 2002). In standard cell culture, in which fibroblasts inevitably form stress fibres, `myofibroblast' generally describes only the -SMA expressing cells. It is tedious to discriminate between myofibroblasts and SMC, for which SMA obviously fails to make the distinction (Fujimoto and Singer, 1987). In normal adult tissue, SMC express a number of late differentiation markers that are usually not in the repertoire of the myofibroblast, including smooth muscle myosin heavy chain (Benzonana et al., 1988), h-caldesmon (Eyden, 2007, 2008), and smoothelin (van der Loop et al., 1996). The muscle intermediate filament protein desmin is an often reliable and widely used exclusion criterion but can be expressed in myofibroblasts in some particular conditions (Hinz et al., 2001b; Skalli et al., 1986). Moreover, SMC normal vessels tissue do not only express desmin but also (or only) vimentin; both intermediate filament proteins are therefore not reliable markers for one or the other cell type (Frank and Warren, 1981; Gabbiani et al., 1981). Equally problematic is the differentiation between myofibroblasts and pericytes if only molecular markers are considered. Depending on their tissue location pericytes can express vimentin and desmin, are -SMA positive and smooth muscle myosin negative (Armulik et al., 2005; Eyden, 2007; Hughes, 2008). However, pericytes are characterized by their close interaction with endothelial cells and lack of contractile features in normal tissues (Eyden, 2007). Discriminating myofibroblasts and SMC as well as pericytes becomes practically impossible in conditions of smooth muscle injury and in cell culture, in particular if the provenance of the cells is unclear. For example, remodelling of injured arteries is thought to be predominantly driven by SMC from the media, de-differentiating into myofibroblasts but the contribution from adventitial fibroblasts to the myofibroblast population has also been suggested (BochatonPiallat and Gabbiani, 2006; Hao et al., 2006; Sartore et al., 2001; Zalewski et al., 2002). Similar uncertainty exists in explant and digestion cultures of tissue containing both connective tissue and smooth muscle. As during arterial remodelling, SMC here lose their late differentiation markers desmin, smooth muscle myosin and smoothelin and acquire a myofibroblastic and synthetic phenotype (Benzonana et al., 1988; Christen et al., 2001; Larson et al., 1984). On the other hand, gene expression profiling supported by protein biochemistry

46


demonstrated that some of the late SMC markers are expressed in fibroblasts after treatment with TGF 1, the most potent myofibroblast inducer (Chambers et al., 2003; Malmstrom et al., 2004). Hence, considering the expression profile of cytoskeletal proteins, the differentiated myofibroblast appears to exist in a continuous spectrum between fibroblasts and SMC. On this background, it seems appropriate to regard the myofibroblast as a phenotype rather than a cell type. Ultimately, one will have to determine what is more important: what a particular cell represents at the time of being assessed or where it came from.

2.4

Origins of the myofibroblast

2.4.1

Local precursors of the myofibroblast

Myofibroblasts are recruited from a variety of precursor cells whose nature depends on the injured tissue and the microenvironment (Hinz et al., 2007) (Fig. 2.2). In many organs, locally residing fibroblasts are considered to be the major source of -SMA positive myofibroblasts in response to injury and development of fibrosis, such as in the skin (Gabbiani, 2003; Hinz, 2007), in fibrotic reactions to body implants (Ariyan et al., 1978; Suska et al., 2008), in liver (Li et al., 2007b; Ramadori and Saile, 2004), kidney (Desmouliere et al., 2003; Qi et al., 2006), and in the stroma reaction to epithelial tumours (De Wever and Mareel, 2002; Desmouliere et al., 2004). EMT is another mechanism to generate fibroblasts and myofibroblasts from epithelial and endothelial precursors in tumour development (Kalluri and Zeisberg, 2006; Thiery, 2002), as well as in kidney fibrosis (Iwano et al., 2002; Kalluri and Neilson, 2003) and possibly lung fibrosis (Chilosi et al., 2003; Kim et al., 2006). EMT has further been demonstrated to contribute to fibrosis of heart (Zeisberg et al., 2007a) and liver (Zeisberg et al., 2007b). In fibrotic liver, hepatic stellate cells are another important source for myofibroblasts (Bataller and Brenner, 2005; Friedman, 2004b; Guyot et al., 2006). De-differentiation of SMC contributes to the generation of myofibroblasts in atheromatous plaques (Bochaton-Piallat and Gabbiani, 2006; Hao et al., 2006). In systemic sclerosis, vessel repair and dermal scarring, pericytes have been suggested to attain contractile myofibroblast features (Rajkumar et al., 2006; Sundberg et al., 1996). Although the repair processes of injured brain exhibit many specific features compared with other organs, astrocytes seem to develop a myofibroblastic phenotype in the glial scar (Moreels et al., 2008; Silver and Miller, 2004).

2.4.2

The circulating fibrocyte, another myofibroblast precursor ± or not?

Another source for reparative fibroblasts that has attracted great interest over the last decade is the so-called fibrocyte, a bone-marrow (BM) derived circulating


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2.2 Myofibroblast precursor cells. Locally residing fibroblasts appear to be the main myofibroblast progenitors in most tissues. In the liver, myofibroblasts are additionally recruited from hepatic stellate cells (HSC). In tumours and in fibrotic liver, lung, and kidney, differentiated myofibroblasts can arise from epithelial- and endothelial-to-mesenchymal transition (EMT). During atheromatous plaque evolution, myofibroblasts arise from de-differentiating smooth muscle cells (SMC). The relative contribution of bone marrow-derived circulating fibrocytes to the formation of differentiated myofibroblasts is unclear at present. Finally, MSC have been shown to acquire the differentiated myofibroblast phenotype in vitro and in vivo.

cell that is recruited to sites of organ injury, inflammation and fibrosis (Abe et al., 2001) (Fig. 2.2). Fibrocytes have been first characterized by the coexpression of fibroblast markers collagen types I and II and fibronectin, of monocyte markers CD13 (aminopeptidase N) and CD11b (M integrin), and haematopoietic progenitor markers CD34, CD45 (protein tyrosine phosphatase receptor type C) and CD105 (endoglin) (Bucala et al., 1994); a more complete characterization has recently been reviewed (Bellini and Mattoli, 2007; Metz, 2003). Early works estimated that fibrocytes make up 0.1±0.5% of the nonerythrocytic cell population circulating in the peripheral blood (Bucala et al., 1994), whereas later studies suggested that fibrocytes do not circulate in their mature form but differentiate from mononuclear precursor cells (Bellini and Mattoli, 2007; Gordon and Taylor, 2005). Mononuclear precursors of the fibrocyte express the CC chemokine receptor (CCR) CCR2 that is implicated in their recruitment to different tissues (Gordon and Taylor, 2005; Moore et al.,

48


2005) and that CCR2 becomes down-regulated during fibrocyte maturation at the tissue site (Abe et al., 2001). It is believed that the fibrocyte is a mandatory differentiation stage before attaining the fibroblastic phenotype in the target tissue (Bellini and Mattoli, 2007; Mattoli et al., 2009) (Fig. 2.2). In culture, fibrocyte-to-myofibroblast differentiation can occurs spontaneously to a certain extent and is inducible with endothelin-1 and TGF 1 (Hong et al., 2007; Schmidt et al., 2003). Up-regulation of myofibroblast markers including -SMA is correlated with down-regulation of the haematopoietic markers CD34 and CD45 (Schmidt et al., 2003). In an animal model of wound healing, loss of CD34 and CD45 fibrocyte markers occurs within hours of fibrocyte recruitment/maturation in inflamed tissue and renders it difficult to determine whether fibrocytes as such are able to differentiate into the myofibroblast in vivo or whether they preferentially localize to sites of myofibroblast accumulation (Mori et al., 2005; Quan et al., 2006). Leukocytespecific protein-1 (LSP-1) has been identified as another fibrocyte marker (Yang et al., 2002). Fibrocyte-derived fibroblastic cells were shown to express LSP-1 up to two months, when CD45 expression is mostly lost (Phillips et al., 2004; Wu et al., 2007b). It remains to be shown whether LSP-1 positive cells can co-express SMA or whether they play a supportive role. Indeed, fibrocytes were shown to promote myofibroblast differentiation from resident fibroblast during the healing of burns by secreting specific growth factors (Wang et al., 2007). To solve the question of whether fibrocytes can differentiate into -SMApositive myofibroblasts, a number of studies used irradiated wild-type mice, engrafted with BM obtained from GFP-expressing transgenic mice or from sexmismatched animals. BM-derived cells were associated in varying numbers with myofibroblast-containing lesions in animals subjected to fibrotic stimuli in different organs (Andersson-Sjoland et al., 2008; Forbes et al., 2004; Haudek et al., 2006; Ishii et al., 2005; Kisseleva et al., 2006; Phillips et al., 2004; Sakai et al., 2006; Schmidt et al., 2003; Wada et al., 2007), during vascular remodelling (Frid et al., 2006; Varcoe et al., 2006), in the context of tumour development (Direkze et al., 2004; Ishii et al., 2005), in lung fibrosis (Moeller et al., 2009), in chronic asthma (Wang et al., 2008) and following dermal wounding (Direkze et al., 2003; Fathke et al., 2004; Mori et al., 2005). On the basis of the used markers it is also possible that fibrocytes are involved in cardiac repair and pathogenesis of atherosclerosis (Fujita et al., 2007; Sata et al., 2002). One study has demonstrated that CD13/collagen I-positive fibrocytes isolated form wound granulation tissue contain BM-derived cells as well as cell populations expressing CD34, CD45 and -SMA; however, these markers were assessed separately (Mori et al., 2005). Because most studies did not simultaneously evaluate BM origin, the fibrocyte character and -SMA expression, fibrocyteto-myofibroblast differentiation in vivo is still a matter of debate (Hinz, 2007). Indeed, some studies appear to rule out that BM-derived fibrocyte progenitors contribute to myofibroblast formation in liver (Kisseleva et al., 2006) and lung


49

fibrosis (Hashimoto et al., 2004); in the latter, myofibroblast differentiation from fibrocytes was inducible by TGF 1 in culture. Fibrocytes are not the only cell types that originate from BM precursors to possibly giving rise to myofibroblasts. BM contains a small fraction of -SMA expressing cells that were considered as being myofibroblasts (Peled et al., 1991; Schmitt-Graff et al., 1989). Using a transgenic mouse that expresses GFP under the control of the -SMA promoter it was suggested that these cells are part of the stromal compartment of the BM with the capacity to circulate in the peripheral blood. These cells were not of hematopoietic character; however, fibrocyte (precursor) marker expression has not been assessed (Yokota et al., 2006). Another possibility is that these BM-derived cells represent a circulating fraction of mesenchymal stem cells (MSC) (Roufosse et al., 2004). Because a number of animal experiments and clinical trials have demonstrated the potential of MSCs to treat human diseases as a cell therapy and in tissue engineering, this cell type will be discussed in greater detail below.

2.5

Mesenchymal stem cells (MSC) and the myofibroblast phenotype: regeneration, repair or risk?

2.5.1

The regenerative potential of MSCs

For regenerative medicine, the population of BM-derived multipotent mesenchymal stromal cells is of major interest (Bianco et al., 2001; He et al., 2007). In culture these cells acquire a fibroblastic character which has been defined as colony-forming unit fibroblast (CFU-F) (Bianco et al., 2001; Friedenstein et al., 1970). The clonogenic cells among the CFU-Fs which exhibit self-renewal and multilineage differentiation character are generally referred to as BM-MSCs. It has to be noted that the stem cell identity of BM-MSC, i.e. the potential for selfrenewal, multilineage differentiation and reconstitution of functional tissue in vivo (Verfaillie, 2002), is not always consequently tested in studies that report to work with this cell type. For this reason, some authors prefer to use the term `mesenchymal precursor cell' instead (Roufosse et al., 2004). BM-MSC currently serve as the major source for experimental and clinical purposes (Caplan, 1991; Chamberlain et al., 2007; Giordano et al., 2007; Pittenger et al., 1999; Pittenger and Martin, 2004; Segers et al., 2006). Other MSC sources are umbilical cord blood (Erices et al., 2000; Lee et al., 2004), adipose tissue (Aust et al., 2004; Zuk et al., 2001), pancreas (Seeberger et al., 2006), pleural cavity (Metcalf, 1972), muscle and brain (Jiang et al., 2002), connective tissue of dermis and skeletal muscle (Young et al., 2001), exfoliated deciduous teeth, and the eye conjunctiva (Nadri et al., 2008). As stated above, MSC have also been suggested to circulate in the peripheral blood but this population is very difficult to identify and/or purify and their contribution to

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tissue regeneration is debated (Fernandez et al., 1997; Orlic et al., 2001b; Roufosse et al., 2004). It is believed that circulating MSC are capable of specifically targeting and entering sites of tissue injury (Fox et al., 2007). MSC in culture are generally identified by the expression of specific cell surface proteins such as CD105, CD90 (Thy1), CD44 (HA receptor), CD73 (SH3 or SH4), CD166 (ALCAM), CD29 ( 1 integrin), CD106 (VCAM) and Stro-1 (Minguell et al., 2001; Pittenger et al., 1999; Simmons and Torok-Storb, 1991). Hence, MSC are different from fibrocytes by being negative for monocyte surface proteins (CD13, CD11b) and haematopoietic markers (CD34, CD45) (He et al., 2007; Pittenger et al., 1999). MSC can differentiate into a variety of cell types that are of potential use for regenerative medicine and tissue engineering (Caplan, 2007); these include chondrocytes and osteocytes for cartilage and bone reconstitution (Bruder et al., 1998; Horwitz et al., 1999; Kadiyala et al., 1997; Noel et al., 2002; Pittenger et al., 1999), myoblasts for skeletal muscle repair (Ferrari et al., 1998), hepatocytes for liver regeneration (Petersen et al., 1999), cardiomyocytes and SMC to repair the cardiovascular system and to promote neo-vessel formation (Chen et al., 2004; Giordano et al., 2007; Orlic et al., 2001a; Psaltis et al., 2008; Rafii and Lyden, 2003; Strauer et al., 2002), and even neuronal cells to treat neurological disorders (Mezey et al., 2000; Picinich et al., 2007). MSC give rise to fibroblasts that can regenerate soft connective tissue (Pittenger and Marshak, 2001; Young et al., 1998) and tendon (Butler et al., 2008) and that are potential candidates for supporting skin repair after wounding (Wu et al., 2007b). In the context of wound healing, MSC-to-epidermal cell differentiation seems to be possible (Deng et al., 2005; Nakagawa et al., 2005; Sasaki et al., 2008; Wu et al., 2007a). In addition to directly regenerating tissue and organs, MSC can exhibit properties that modulate the body-inherent repair processes. Directly injected BM-MSC contribute to skin regeneration after wounding and supernatants from these cells stimulate tube formation of vascular endothelial cells and recruitment of macrophages to the wound, suggesting positive paracrine effects (Chen et al., 2008; Wu et al., 2007a). In co-culture, paracrine actions of BM-MSC stimulate proliferation and differentiation of skin epidermal cells (Aoki et al., 2004). Moreover, MSC are able to migrate to sites of damaged tissues where their immunosuppressive and immunomodulatory properties improve the tissue transplantation success (Le Blanc and Pittenger, 2005; Uccelli et al., 2008). Finally, MSC are immunologically immature and do not elicit inflammatory responses, which extends their possible use to gene delivery (Chamberlain et al., 2007; Pereboeva et al., 2003).

2.5.2

MSC-to-myofibroblast differentiation

Many of the potential therapeutic applications that have been proposed for MSC imply their engraftment into fibrotic tissue. In chronic fibrosis the high ECM


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secreting and contractile activities of endogenous myofibroblasts destroy the functional architecture of the remaining healthy tissue. Even if progression of fibrosis could be prevented, no efficient treatment exists to reverse this destruction and to reconstruct normal tissue. This is particularly evident in cardiac fibrosis following infarct and in pulmonary fibrosis (Gauldie et al., 2007; Phan, 2002; Wynn, 2008). MSC therapy has been proposed to regenerate fibrotic tissue (Ortiz et al., 2003) but it is presently unclear whether transplantation of undifferentiated MSCs can possibly fulfil this task. A number of recent findings rather suggest that the diseased microenvironment will drive regenerative MSC into fibrogenic myofibroblasts that will further distort the present ECM. What is the evidence for MSC-to-myofibroblast differentiation? One of the defining features of MSC in cell culture is their well-spread morphology and the formation of contractile stress fibres (Pittenger et al., 1999), which fulfils the criteria for the proto-myofibroblast phenotype (Tomasek et al., 2002). A number of studies have further reported spontaneous development of the differentiated myofibroblast phenotype in cultured BM-MSC, evidenced by de novo expression of -SMA and up-regulated contractile activity (Bonanno et al., 1994; Cai et al., 2001; Kinner et al., 2002b; Peled et al., 1991) (Fig. 2.2). The percentage of BM-MSC spontaneously expressing -SMA is gradually increasing with long-term culture (Charbord et al., 1985; Galmiche et al., 1993; Yokota et al., 2006). Moreover, MSC from different origins have been shown differentiate into myofibroblasts in response to stimuli that are known inducers of -SMA in fibroblastic cells. Proteomic profiling of BM-MSC revealed a myofibroblast differentiation program upon treatment with TGF 1 (Wang et al., 2004). Similarly, MSC derived from human adipose tissue de novo express -SMA upon treatment with lysophosphatic acid and with bradykinin in TGF 1 and Smad2/3 dependent manner (Jeon et al., 2008; Kim et al., 2008). Treatment with basic fibroblast growth factor (FGF-2) (Hankemeier et al., 2005) as well as application of cyclic mechanical strain (Kobayashi et al., 2004) induced -SMA expression in cultured BM-MSC and MSC differentiate into myofibroblasts in chondrogenic conditions (Hung et al., 2006). In co-culture with PDGF-Bactivated fibroblasts, BM-MSC are specifically recruited and differentiate into myofibroblasts in a process involving FGF-2 and CXCL5 (Nedeau et al., 2008). Application of mechanical stimuli drives MSC along a myogenic lineage, including expression of -SMA and other markers of the myofibroblast (Park et al., 2007). It has to be noted that not all -SMA-positive MSC in culture are necessarily myofibroblasts but may represent presumptive stages of differentiation into SMCs or pericytes (Bianco et al., 2001; Charbord et al., 1990). This distinction, however, is difficult to make because SMC attain the myofibroblast phenotype in culture (Bochaton-Piallat et al., 1992) and myofibroblasts can extend to the very far spectrum of SMC differentiation upon treatment with TGF 1 (Chambers et al., 2003).

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2.5.3


The threat of MSC-to-myofibroblast differentiation for regenerative medicine

What is known about the significance of MSC-to-myofibroblast differentiation in vivo? Recent studies evaluating the interaction between MSC and epithelial tumour cells in vivo and in vitro indicate that acquisition of the myofibroblast phenotype by MSC can reduce the success of the envisaged therapy and can even amplify the disease. Different groups have shown that systemically transplanted MSC target to the stroma environment of epithelial tumours (Hall et al., 2007; Hung et al., 2005; Menon et al., 2007; Studeny et al., 2004). This specific homing together with the immuno-inactivity of MSC was suggested to be useful for the tumour-specific delivery of anti-cancer drugs, cytokines and viruses (Komarova et al., 2006; Studeny et al., 2002) and to suppress tumour development as such (Khakoo et al., 2006). On the other hand, the tumour appears to activate MSC-to-myofibroblast differentiation in a fashion similar to carcinomaassociated fibroblasts (Mishra et al., 2009). BM-MSC cultured in tumourconditioned medium were shown to differentiate into -SMA-positive myofibroblasts (Mishra et al., 2008). MSC that have become activated by mildly invasive human breast carcinoma cells in culture enhance the metastatic potential of the cancer cells when injected subcutaneously; this effect is mediated in a feedback paracrine loop (Karnoub et al., 2007). Another detrimental effect of MSC-to-myofibroblast differentiation has been observed for organs that are sought to be regenerated with MSC therapy. Schneider and coworkers have tested the stability of the MSC character in a three-dimensional culture model of air-exposed dermal equivalent, similar to those used in skin tissue engineering (Schneider et al., 2008). In these keratinocyte-promoting growth conditions MSC did not trans-differentiate along the anticipated epithelial lineage but instead developed into fibrogenic myofibroblasts that contracted the ECM. Similarly, the high contractile activity of myofibroblastic MSC has been shown to deform scaffold matrices that are used to deliver MSC for musculoskeletal tissue engineering purposes (Kinner et al., 2002b). Excessive myofibroblast activity has further been demonstrated to have detrimental effects for the healing of these tissues (Premdas et al., 2001) although the in vivo function of myofibroblastic MSC in these conditions has not been tested yet. Yan and collaborators analysed the success of mouse BM-MSC transplantation as a treatment of lung injury induced by irradiation (Yan et al., 2007). MSC injected into the injured lung immediately after irradiation differentiated into functional lung epithelial and endothelial cells. In contrast, MSC injected 2 months after irradiation when fibrosis has developed, cells appeared as -SMApositive `myofibrocytes' that were involved in fibrosis progression (Yan et al., 2007). Similarly, injection of MSC into mice was shown to improve the outcome of acute renal injury, presumably by restoring the glomerular basement


53

membrane but not of the interstitial fibrosis following chronic renal disease (Ninichuk et al., 2006). Likewise, in a model of acute liver injury and fibrosis in immunodeficient mice, systemically transplanted human BM-MSC were shown to engraft in high numbers in the damaged liver. However, the wanted differentiation into hepatocytes occurred only at very low rates whereas most cells of human origin developed a myofibroblastic phenotype (di Bonzo et al., 2008). It appears likely that these cells will rather contribute to fibrosis than to regenerate the liver. Concomitantly, MSC reduced development of liver fibrosis when systemically transfused immediately after inducing fibrosis but not when delivered after 1 week of fibrosis development (Fang et al., 2004). Another major application of MSC is the potential repair of infarcted heart which already entered advanced clinical trials with variable success rates (Chamberlain et al., 2007; Giordano et al., 2007; Laflamme and Murry, 2005; Pittenger and Martin, 2004; Prockop and Olson, 2007; Segers et al., 2006; Shake et al., 2002). It has been acknowledged that the hostile environment of the infarcted heart, including ischaemia, inflammation and fibrosis is one important factor that reduces the success of endogenously and exogenously promoted heart regeneration (Segers and Lee, 2008). Fibrosis has been identified as a strong physical barrier for the entry of regenerative cells in zebrafish heart regeneration (Poss et al., 2002) and the fibrotic environment was shown to modulate regenerative cells in unwanted ways. BM-derived stem cells injected into the myocardial scar after infarct can lead to myocardial calcifications (Breitbach et al., 2007). On the other hand, MSC delivered to acutely infarcted heart were shown to attenuate development of fibrosis, presumably by exerting `trophic' paracrine effects that reduce myofibroblast differentiation of cardiac fibroblasts (Caplan and Dennis, 2006; Nagaya et al., 2005; Ohnishi et al., 2007). This effect could underlie the observation that injection of MSC early after cardiac infarct reduces the stiffness of the scar which should reduce its physical barrier function; softer environment is expected to reduce myofibroblast differentiation as discussed further below (Berry et al., 2006). In summary, is it conceivable that the timing of MSC transplantation is critical for the success of the regenerative therapy; when engrafting immediately after organ damage before the onset of fibrosis the local microenvironment will be very different from later fibrotic stages. In light of the fact that most patients are first seen by a clinician when lung and liver fibrosis has already progressed, transplanting MSC without additional measures into these organs appears a difficult and even dangerous strategy. This situation is different in the case of cardiac fibrosis following myocardial infarction because the onset of the damage is generally known.

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2.5.4


Possible benefits of MSC-to-myofibroblast differentiation for regenerative medicine

From the studies above it is amply clear that MSC have great potential to improve tissue regeneration when transplanted under the right conditions but can become detrimental when differentiating into fibrogenic cells. The question remains whether MSC-to-myofibroblast differentiation can exert beneficial actions and actually support tissue reconstruction. In a rabbit model of corneal wound healing, systemically transplanted BM-MSC were shown to home to the injured site and to improve corneal wound healing by differentiating into myofibroblasts (Ye et al., 2006). Similarly, GFP-expressing BM-MSC topically applied to open rat wounds differentiate into granulation tissue myofibroblasts and accelerated the wound healing progress (Yamaguchi et al., 2005). In a mouse skin wound model, intravenously injected BM-MSC were shown to recruit to the wound site and to accelerate wound healing by differentiating into different cell types, including -SMA-positive cells (Sasaki et al., 2008). Because rodents do not develop hypertrophic scars it is difficult to predict if such a treatment will also improve the quality of human skin wound healing. In conditions of experimental colitis, BM-derived MSC were shown to contribute to gut repair by replenishing the population of pericryptal myofibroblasts (Brittan et al., 2002, 2005). MSC-to-myofibroblast differentiation may also play a beneficial role for the repair of certain musculoskeletal tissues. Differentiated chondrocytes and osteoblasts, the cells involved in physiological cartilage and bone repair, pass over a controlled myofibroblast-stage which supports the healing process by priming the specific fibrous tissue architecture (Kinner et al., 2002a; Spector, 2001; Wang et al., 2000). Chondrocytes expressing -SMA populate the superficial layer of articular cartilage which is suggested to protect cartilage from damage (Kim and Spector, 2000). A similar role in preserving cartilage-like structure has been demonstrated for MSC subjected to chondrogenic differentiation in culture (Hung et al., 2006). MSC grown in chondrogenic culture in the presence of TGF 1 generated cartilagelike pellets with an annular surface that was populated by -SMA-positive cells. In contrast, removal of TGF 1 and consequent loss of the myofibroblast phenotype resulted in the loss of the structural integrity of the pellets (Hung et al., 2006). The question of whether the MSC-to-myofibroblast differentiation is beneficial or detrimental will depend on the specific repair demands in a given tissue and on the nature of the tissue itself. It appears that acute repair of connective tissue can be supported by MSC-derived myofibroblasts whereas myofibroblastic MSC will not directly contribute to the regeneration of other organs. In either case endogenous control over myofibroblast development is desired which requires profound knowledge of the control mechanisms and factors involved.


2.6

55

What drives myofibroblast differentiation?

To therapeutically counteract organ dysfunction caused by myofibroblasts it is crucial to understand the general molecular pathways regulating their evolution and function. I will here concentrate on the mechanisms that regulate myofibroblasts independently from a specific organ, disease or myofibroblast origin.

2.6.1

Mechanical stress regulates myofibroblast development

It emerges as a common theme that mechanical stress is one of the most potent factors controlling myofibroblast fate and development from different precursor cells (Fig. 2.1). After injury of connective tissues, mechanical stress results from the partial or total loss of the mechano-protective ECM architecture and the residing cells are directly exposed to the stress. To resist the mechanical load arising during tissue repair and remodelling processes and to prevent tissue rupture myofibroblast precursors develop tension on their own by building up mechano-resistant stress fibres. The ultimate goal of myofibroblast activity is to restore the mechanical integrity of the tissue by secreting and organizing new ECM, a process that is precisely controlled by mechanical feedback signals from the ECM, such as ECM stiffness. What is a `stiff' ECM in a physiological and pathological sense and how does ECM stiffness develop during tissue repair and remodelling? The stiffness of the provisional ECM laid down after acute wounding is low, with an elastic modulus (the physical unit of compliance) of ~100±1000 Pa. Fibroblastic cells subjected to similar mechanical conditions in vitro by growth on soft two-dimensional polyacrylamide gels do not develop stress fibres (Discher et al., 2005; Tamariz and Grinnell, 2002; Yeung et al., 2005) (Fig. 2.1). Hence, acquisition of the proto-myofibroblast phenotype and consequently of the differentiated myofibroblast is suppressed on soft substrates. In contrast, SMA-negative stress fibres start to develop on increasingly stiff twodimensional culture substrates exhibiting an elastic modulus of 3000±6000 Pa (Yeung et al., 2005). The threshold ECM stiffness for occurrence of -SMA in stress fibres ranges around 20 000 Pa as demonstrated for myofibroblasts cultured on silicone surfaces with tunable stiffness (Goffin et al., 2006) (Fig. 2.1). A comparable ECM stiffness of ~15 000 Pa activates hepatic stellate cells into -SMA-positive myofibroblasts in the appropriate growth conditions (Wells, 2005). The synthetic polyacrylamide and silicone substrates used in the studies above provide stable mechanical growth conditions; in contrast, reparative cells in vivo change and stiffen their own mechanical microenvironment. This situation can partly be reproduced in vitro using three-dimensional collagen gels with different stiffness and under different mechanical constraints (Grinnell, 2003). In mechanically unrestrained and/or newly polymerized collagen gels fibro-

56


blastic cells attain a stellate-like morphology and do not develop contractile features (Grinnell et al., 2003; Hinz, 2006). Resulting from the cell remodelling activity ECM stiffness increases over time which is reflected by the de novo development of stress fibres within hours of remodelling of anchored gels (Fig. 2.1) (Hinz, 2006). Exertion of cellular force in this phase is appreciated from the increasing ECM organization level in mechanically restrained (anchored) gels and from gel size reduction in unloaded (free floating) gels (Grinnell, 2000). Full myofibroblast differentiation, i.e. expression of -SMA, requires an even higher level of collagen gel stiffness that builds up with increasing culture time. Importantly, even the strongly pro-fibrotic cytokine TGF 1 is not effective to promote myofibroblast differentiation in soft substrate conditions (Arora et al., 1999; Goffin et al., 2006) (Fig. 2.1). A similar sequence of events that is linked with increasing ECM stiffness and stress has been demonstrated for in vivo healing of full thickness dermal wounds. Fibroblasts populating very early wound granulation tissue (1±3 days postwounding) are devoid of stress fibres (Hinz et al., 2001b) but develop actin filament bundles in 5±6-day-old wound granulation tissue (Hinz et al., 2001b). Expression of -SMA and occurrence of differentiated myofibroblasts is not observed before day 8 in experimental rat wounds (Darby et al., 1990; Hinz et al., 2001b), when ECM stiffness reaches values around 30 000 Pa (Goffin et al., 2006) (Fig. 2.1). Preventing wound closure by mechanically splinting the edges of experimental wounds accelerates expression of -SMA compared with normally healing wounds; stress release by removing the splint leads to reduced -SMA expression (Hinz et al., 2001b). In other fibrotic tissues and in granulation tissue toward the end of wound healing ECM stiffness values of greater than 50 000 Pa have been measured (Goffin et al., 2006). Increased stiffness of a damaged organ is not necessarily associated with scar formation but can precede and promote fibrosis as shown after liver injury (Georges et al., 2007). The mechanisms and intracellular signalling pathways through which tension controls -SMA transcription appear to involve Rho/Rho-associated kinase and have been reviewed elsewhere (Wang et al., 2006; Zhao et al., 2007). In addition to being regulated on the expression level, -SMA is considered as mechanosensitive protein. Reducing stress fibre tension by reducing substrate stiffness and/or by inhibiting intracellular contraction first results in the selective removal of -SMA from persisting stress fibres (Goffin et al., 2006) (Fig. 2.1). The fact that -SMA only localizes to stress fibres under significant mechanical load is believed to provide a mechanism for rapidly controlling myofibroblast contractile function (Goffin et al., 2006; Hinz, 2006). This cellular control mechanism provides a possible target to therapeutically counteract myofibroblast function and progression in excessive tissue contractures as elaborated in Section 2.7.


2.6.2

57

Latent TGF 1: pro-fibrotic cytokine with mechanosensory function

TGF 1 is the most potent myofibroblast-inducing factor and one of the strongest pro-fibrotic cytokines presently known. TGF 1 exerts its pro-fibrotic activities by mediating the inflammatory response, causing excessive ECM production, increasing the synthesis of tissue inhibitors of metallo-proteinases (TIMPs), decreasing protease synthesis and finally by inducing myofibroblast differentiation (Desmouliere et al., 1993; Grainger, 2007; Hinz, 2007; Leask and Abraham, 2004; Ruiz-Ortega et al., 2007; Taipale et al., 1998). The fact that cancer cells become insensitive to the growth-arresting action of TGF together with its proangiogenic effects and its potential to inducing EMT of cancer cells allocates TGF 1 also a central role in tumour development (Bierie and Moses, 2006; Pardali and Moustakas, 2007; Siegel and Massague, 2003). Because TGF 1 signalling also assures homeostasis of adult tissues by controlling proliferation of epithelial cells, endothelial cells, immune cells and fibroblasts (Feng and Derynck, 2005; ten Dijke and Arthur, 2007; Wakefield and Stuelten, 2007), global inhibition of TGF 1 is problematic as anti-fibrotic therapeutic strategy with many uncontrollable side-effects. On the other hand, the complex and diverse mechanisms leading to the activation of latent TGF 1 potentially provide the means for a cell-specific inhibition of TGF 1 action. Activation of latent TGF 1 requires its dissociation from the latency associated peptide (LAP) that is co-synthesized in complex with TGF 1 (Annes et al., 2003). The vast majority of cell types secrete TGF 1 as part of a large latent complex, consisting of TGF 1, LAP and the latent TGF 1 binding protein (LTBP-1) (Annes et al., 2003; Todorovic et al., 2005) (Fig. 2.3a). LTBP-1 is a member of the fibrillin family of ECM proteins that binds to several other ECM components, including fibrillin-1, FN, and vitronectin, thereby providing a reservoir of latent TGF 1 in the ECM (Annes et al., 2003; Todorovic et al., 2005). Activation of latent TGF 1 by its dissociation from LAP is promoted by various mechanisms which differ according to the cell type and the physiological context. Latent TGF 1 activation occurs upon proteolytic cleavage, by interaction with thrombospondin 1 and with the mannose-6-phosphate receptor (Annes et al., 2003; Jenkins, 2008) (Fig. 2.3b). Moreover integrins, as transmembrane components of cell-ECM adhesions, have been reported to play a major role in activating latent TGF 1 (Sheppard, 2005; Wipff and Hinz, 2008). Two principal mechanisms have been proposed and are experimentally supported how integrins can activate a growth factor. The first mechanism is sensitive to protease inhibitors and proposes integrins as common docking point for latent TGF 1 and its activating proteases. The second mechanism is independent from any proteolytic action and involves cell traction forces which are directly transmitted to the large latent complex via integrins (Jenkins, 2008; Sheppard, 2005; Wipff and Hinz, 2008) (Fig. 2.3c).

58


2.3 Activation of latent TGF 1. (a) TGF 1 is secreted in a ECM-bound large latent complex (LLC), consisting of TGF 1 and the latency associated peptide (LAP) that form the small latent complex (SLC), and the latent TGF 1-binding protein (LTBP-1. (b) Integrin-independent proteolytic activation of latent TGF 1 occurs at specific sites that are sensitive to proteolytic digestion (scissors), leading to TGF 1 release. (c) Integrins v 6, v 5 and v 3 have been shown to activate latent TGF 1 independently from proteolytic activity. They all recognize the integrin binding sequence RGD of the LAP moiety in the LLC. It has been proposed that when the LLC is covalently bound to a mechanically resistant ECM, cell traction forces exerted to LAP will result in a deformation change of the latent complex that liberates active TGF 1 (from Wipff and Hinz, 2008).

First evidence that integrins can directly activate latent TGF 1 independently from any proteolytic activity was provided for the epithelial integrin v 6 (Annes et al., 2004; Jenkins et al., 2006; Munger et al., 1999). Functional knockout of this integrin produces a phenotype in mice that closely resembles that of a TGF 1 knockout (Huang et al., 1996; Shull et al., 1992). Concomitantly, the lungs of 6 knockout mice are protected from fibrosis (Munger et al., 1999). Later, the integrins v 5, v 8, a yet unidentified 1 integrin and possibly v 3 integrin have also been reported to participate in activating latent TGF 1 (Sheppard, 2005; Wipff and Hinz, 2008). All integrins that contribute to


59

the activation of latent TGF 1 physically interact with the LAP portion of the large latent complex (Sheppard, 2005). Inhibiting the respective binding sequence in vitro abolishes latent TGF 1 activation in epithelial cells (Munger et al., 1999) and in myofibroblasts (Wipff et al., 2007); genetic deletion of this sequence in mice strongly resembles the TGF 1 knockout (Yang et al., 2007). The following model has been proposed for direct and integrin-mediated activation of latent TGF 1 by contractile myofibroblasts: (1) The integrin v 5 binds extracellularly to the LAP portion of the large latent complex and intracellularly to -SMA positive stress fibres. (2) The forces generated by contractile stress fibres pull on the latent complex via its integrin binding site. (3) Binding of LTBP-1 to the ECM provides mechanical resistance to the pulling which leads to opening of the complex and release/presentation of active TGF 1 (Fig. 2.3c). This model is supported by the findings that inhibition of integrin v 5 with function blocking antibodies diminishes latent TGF 1 activation by cultured myofibroblasts and reduces the fibrogenic character of fibroblastic cells (Asano et al., 2006). In addition, myofibroblasts activate latent TGF 1 as a function of their contractile activity (Wipff et al., 2007). Inducing myofibroblast contraction with thrombin, angiotensin-II and endothelin-1 increases latent TGF 1 activation; this effect depends on integrin binding to LAP (Wipff et al., 2007). Finally, to induce a putative conformational change in the latent TGF 1 complex by integrin-mediated pulling on the LAP, the ECM must provide mechanical resistance. Indeed, myofibroblasts only activate TGF 1 by integrinmediated contraction when cultured on substrates with a stiffness that corresponds to that of contracting fibrotic and granulation tissue but not when grown on substrates exhibiting the compliance of normal connective tissue (Wipff et al., 2007). This dependence of the availability of active TGF 1 on ECM stiffness would restrict generation and autocrine maintenance of the myofibroblast phenotype to a mechanical microenvironment that has been sufficiently pre-remodelled and stiffened for being efficiently contracted. Interfering with the integrins that are implicated in latent TGF 1 activation is one possible strategy to therapeutically counteract the harmful activity of active TGF 1 in a cell-specific manner, without impairing its beneficial effects on other cell types.

2.6.3

Other factors that modulate myofibroblast differentiation

Discussing all the intracellular signalling molecules, cytokines and ECM proteins that modulate myofibroblast differentiation and -SMA expression would by far exceed the scope of this chapter and the reader is referred to the respective literature (Hinz, 2007; Horowitz and Thannickal, 2006; Schurch et al., 2007; Wynn, 2007). Most differentiated myofibroblast-inducing factors act in synergy with TGF 1 signalling and are not effective in inducing -SMA on

60


their own (Desmouliere et al., 1993; Ronnov-Jessen and Petersen, 1993). Factors that stimulate the differentiation of myofibroblasts from their precursors without involving paracrine effects from other cell types, include CTGF (connective tissue growth factor) (Shi-Wen et al., 2008), IL-6 (Gallucci et al., 2006), Fizz1 (found in inflammatory zone) (Liu et al., 2004), galectin-3 (Henderson et al., 2006), osteopontin (Lenga et al., 2008; Mori et al., 2008), endothelin-1 (Jain et al., 2007; Shi-Wen et al., 2007), angiotensin II (Guo et al., 2001; Mezzano et al., 2001; Rosenkranz, 2004; Uhal et al., 2007), thrombin (Bogatkevich et al., 2003), possibly semaphorin 7A (Kang et al., 2007), NGF (nerve growth factor) (Micera et al., 2001) and cleavage of the urokinase receptor (Bernstein et al., 2007). Myofibroblast differentiation is further promoted by cell adhesion to specific ECM proteins including collagen type VI (Naugle et al., 2006), tenascin-C (De Wever et al., 2004; Tamaoki et al., 2005) and most importantly ED-A FN (Serini et al., 1998). In addition, pathogen-associated molecular patterns, such as bacterial lipoproteins, DNA and double-stranded RNA can bind to receptors on the surface of fibroblastic cells (Akira and Takeda, 2004) and have been shown to activate myofibroblasts in the intestine (Otte et al., 2003). Another important stimulating factor for myofibroblast differentiation appears to be the production of reactive oxygen species by NADPH oxidases (NOX) in fibroblastic cells (Shen et al., 2006). NOX are transmembrane proteins that regulate intracellular redox signalling by reducing extracellular molecular oxygen to superoxide generating downstream reactive oxygen species (Bedard and Krause, 2007). The predominant NOX isoform in fibroblasts is NOX4, which has been shown to mediate TGF 1-induced conversion of cardiac fibroblasts into myofibroblasts (Cucoranu et al., 2005). The vast majority of factors that were shown to exert a myofibroblast-inducing effect have been tested on fibroblasts from different tissue origins. It remains to be shown whether all identified myofibroblast precursor cells similarly respond to these factors. This appears the case for TGF 1, being an accepted myofibroblast promoter in fibroblasts (Desmouliere et al., 1993; Ronnov-Jessen and Petersen, 1993), hepatic stellate cells (Gressner and Weiskirchen, 2006), astrocytes (Moreels et al., 2008), epithelial cells (Masszi et al., 2003; Willis et al., 2005), fibrocytes (Hong et al., 2007) and MSC (Wang et al., 2004).

2.7

Lessons to be learned from the myofibroblast for the effective use of mesenchymal stem cells (MSC)

To suppress and/or reduce the contribution of engrafted MSC to the development of fibrotic scar, MSC delivery to diseased organs could be combined with the delivery of agents that down-regulate myofibroblast development. The search for anti-fibrotic drugs is intense and tissue engineering as well as regenerative medicine will benefit from the discovery of new therapies against


61

organ fibrosis (Brown et al., 2005; Friedman, 2004a; Gharaee-Kermani et al., 2007; Horowitz and Thannickal, 2006; Scotton and Chambers, 2007). At present, however, none of the countless factors that are implied in normal and pathological wound healing has been successfully targeted to significantly improve the tissue repair process in clinical applications. This is particularly true for therapies that imply growth factors which are subjected to rapid degradation in the aggressive wound environment and require precisely controlled timing of administration to achieve the wanted effects (Chan et al., 2006). Growth factors that antagonize myofibroblast development in culture and in animal models have been described, including IL-1 (Kanangat et al., 2006; Shephard et al., 2004), tumour necrosis factor- (TNF-) (Goldberg et al., 2007; Saika et al., 2006), TGF 3 (Shah et al., 1995) and interferon- (IFN- ) (Desmouliere et al., 1992). Using MSC as vehicle to deliver these growth factors specifically to fibrotic scars and simultaneously to regenerate the tissue can be one future strategy to improve tissue regeneration. However, much is left to be done before this utopian vision can be applied in clinics. It is unclear whether MSC respond to the above-mentioned factors in a similar fashion than fibroblasts and how the engrafted microenvironment will modulate their response. For instance, FGF-2 appears to support myofibroblast differentiation of MSC (Hankemeier et al., 2005; Nedeau et al., 2008) but antagonizes myofibroblast development of SMC, pericytes and fibroblasts (Cushing et al., 2008; Maltseva et al., 2001; Papetti et al., 2003). Another possibility to suppress development of the myofibroblast phenotype in MSC is to interfere with the mechanical feedback loop of high contractile activity and ECM stiffening that induces and maintains the fibrogenic cell character (Wipff et al., 2007) (see Section 2.6.1). This strategy has been proven successful to inhibit fibroblast-to-myofibroblast differentiation in different ways. First, reducing the stiffness of the microenvironment will lower the stress exerted on the MSC and possibly suppress its myofibroblast development. It is difficult to imagine how one could reduce the stiffness of a scar tissue in a controlled manner if MSC are to be delivered systemically. However, when using a scaffold delivery strategy the mechanical property of the biomaterial or synthetic material can be adjusted to control the fibrogenic behaviour of implanted MSC (Ghosh and Ingber, 2007). Second, if the mechanical properties of the microenvironment are not controllable, interfering with the mechanisms through which MSC perceive extracellular stress is another option. It has been shown that the level of substrate rigidity determines the size and molecular composition of cell±matrix focal adhesions that perceive and communicate extracellular mechanical signals to the cytoskeleton, leading to specific gene expression (Bershadsky et al., 2003; Ingber, 2003). By artificially reducing the adhesion area available for cell attachment using microcontact printing on rigid surfaces it is possible to `simulate' a soft environment for myofibroblasts. As a consequence, these cells lose -SMA expression in stress fibres and contractile

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capacity similar to growth of soft substrates (Goffin et al., 2006). Third, the level of myofibroblast intracellular tension can be specifically and directly reduced by targeting -SMA positive stress fibres using a competitive peptide strategy. Inhibiting myofibroblast contraction by long-term administration of the so-called SMA fusion peptide results in decreased collagen production and finally disappearance of the myofibroblast (Clement et al., 2005; Hinz et al., 2002). Whereas the latter option has not yet been tested for MSC, modulating substrate stiffness and cell adhesion area have already been shown to influence MSC differentiation into different lineages. Engler and collaborators could demonstrate that growth on differently compliant polymer surfaces induces early lineage differentiation of MSC (Engler et al., 2006). In otherwise identical culture conditions, adjusting ECM stiffness to the stiffness of their natural body environment guides MSC along the according differentiation pathway. On `brain-soft' gels with elastic modulus of ~1000 Pa, MSC express early neurogenic marker, growth on `muscle-stiff' substrates of ~11 000 Pa induces early myogenic factors and `bone-stiff' substrates of >34 000 Pa induces osteogenic differentiation (Engler et al., 2006). Similar observations were made by dictating the size of the surface area available for MSC adhesion. MSC are commitment to the adipocyte lineage when grown on small adhesion islands or into osteoblasts when plated on large islands; this mechanical restriction was even capable to override the effect of specific differentiating growth factors added to the medium (McBeath et al., 2004). Myofibroblastic differentiation of MSC on soft and micropatterned substrates has not been assessed in the appropriate fibrogenic conditions but the clear mechano-responsiveness of MSC suggests an effect of substrate stiffness also on the development of this phenotype.

2.8

Conclusions and future trends

Most studies that search to understand how myofibroblasts develop and how their activity is controlled are motivated by the desire to fight organ fibrosis, one of the major causes of death in Western countries. I have here evaluated the threat and/or potential that this cell phenotype represents for regenerative medicine with a specific focus on MSC-to-myofibroblast transition. It remains to be shown whether and when the myofibroblastic MSC is our friend or our enemy; in any case knowing your enemy raises the chances of success of tissue regeneration. This spirit is expressed in a very loose translation of the wellknown statement made by Sun Tzu in `The Art of War': `So it is said that if you know your enemies and know yourself, you will fight without danger in battles. If you only know yourself, but not your opponent, you may win or may lose. If you know neither yourself nor your enemy, you will always endanger yourself.'


2.9

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The structure of articular cartilage E . B . H U N Z I K E R , University of Bern, Switzerland

Abstract: Mature articular cartilage has a highly anisotropic structure which is indispensable for its mechanical competence. Immature articular cartilage has an isotropic structure which reflects its dual function as an articulating layer and a superficial growth plate. In this chapter, the structures of immature and mature articular cartilage are described, and the physiological mechanism underlying the evolution of the former into the latter is discussed. Inter-species differences in articular cartilage structure, and structural-functional correlations in humans, are also addressed. Key words: tissue resorption, tissue substitution, inter-species differences, anisotropic structure, isotropic structure.

3.1

Introduction

The abutting ends of the long bones that constitute a synovial joint are mantled by a layer of articular cartilage, which mediates the transfer of loads and permits the frictionless movements of the skeletal elements. This primarily mechanical function of the mature tissue is reflected in its highly anisotropic organization into several morphologically distinct zones. However, during the early phase of postnatal development, the tissue manifests an isotropic structure, which likewise has its functional corollaries. At this time, the tissue acts not only as a layer that mediates frictionless joint movement, but also as a surface growth plate for the rapid elongation and modelling of the epiphyseal bone. At a later phase of postnatal development, the isotropic organization of the tissue is transformed into the anisotropic architecture that typifies the adult organism. This transformation is achieved by a process of tissue resorption and substitution. During the growth phase of postnatal development, stem cells within the superficial zone feed the proliferating pool within the transitional and upper radial ones. Within the lower radial zone, the cells hypertrophy, and their extracellular matrix then undergoes mineralization prior to resorption. Towards the end of this phase, as the organism approaches skeletal maturity, the proliferative, but not the metabolic, activity of the cells ceases: they continue to remodel their extracellular matrix. Adult human articular cartilage is characterized by an extremely low numerical density of cells, which constitute less than 2% of the tissue volume.

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The mechanical properties of the layer are conferred chiefly by the abundant extracellular matrix. The structure of adult articular cartilage and the degree of anisotropy differ greatly between mammalian species that are commonly used as experimental animal models, for example the rabbit, the goat and the miniature pig. Not only the overall height of the layer, but also the relative heights of each zone vary tremendously for a given joint. The numerical density of cells within each zone is also subject to great diversity, although the total number contained within a unit area of tissue beneath the joint surface is remarkably constant. The fibrillar and macromolecular organization of each zone is likewise similar in the different mammalian species. However, the temporal course of postnatal growth and maturation differs greatly between these.

3.2

General structure and function of articular cartilage

The synovial joints of mammals comprise the abutting ends of mostly long bones, each of which is covered with a layer of articular cartilage. The articular cartilage layer is continuous with the underlying subchondral bone. The bony heads are surrounded by the synovial membrane and the joint capsule. Also associated with the joint are ligaments, tendons and muscle. A synovial space intervenes between the abutting layers of articular cartilage. This space is filled with a fluid, which is secreted by the lining cells of the synovial membrane. The abutting layers of articular cartilage, together with the intervening synovial fluid, permit the practically frictionless movement of the two associated bones and the transfer of loads between them. These biomechanical functions are reflected in the structure of articular cartilage. In the light microscope, this layer is revealed to be highly anisotropic in structure. The cells are organized into distinct vertical columns and horizontal strata relative to the articular surface. The first horizontal layer, namely, that closest to the articular surface, is referred to as the superficial zone. It consists of cells with an ellipsoidal or spindle-like form, whose long axis runs parallel to the articular surface, and which exist singly. The cells are embedded within a meshwork of fine collagen fibrils and fibres, which likewise run parallel to the articular surface. The superficial zone generally makes up one-thirteenth of the total height of the articular cartilage layer. It gives way to the transitional zone, which contains cells with a more rounded profile. These cells exist singly or in pairs. The collagen fibres form hemispherical arcades, which are continuous with vertically-running counterparts in the deeper zones. The transitional zone makes up about two-thirteenths of the total height of the articular cartilage layer. The superficial and transitional zones, being relatively poor in proteoglycans, stain weakly with cationic dyes. The transitional zone gives way to the radial zone, which is subdivided into an upper and a lower portion, each of which makes up about five-thirteenths of the total height of the articular cartilage layer. This


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zone is characterized by a gradual and marked increase in the size of the cells, which, in form, assume the appearance of oblate spheroids, and which are grouped into chondrons containing up to four vertically stacked chondrocytes. The concentration of proteoglycans increases continuously between the upper and the lower radial zones. Both regions are rich in vertically oriented collagen fibres, which course between the vertical columns of cells and which increase in thickness with increasing depth. The thick fibres at the base of the lower radial zone penetrate the underlying and fairly thin layer of calcified cartilage, which gives way to the subchondral bone. The interface between the layer of calcified cartilage and the subchondral bone plate has a characteristically undulating profile, which reflects the interdigitation of the two tissues and assists in anchorage. The upper border of the calcified cartilage layer is sometimes referred to as the tidemark. Its prominence as a line in conventionally processed tissue sections is an artefact, which is generated by decalcification. In the absence of decalcification, the mineralization front is represented as a discrete boundary between the unmineralized and the mineralized cartilage compartments (Hunziker et al., 1997). The anisotropic structure of mature articular cartilage has been described in detail by Benninghoff (1925). It is apparent in all synovial joints, irrespective of their size or of the animal species (Fig. 3.1). The maintenance of this highly anisotropic structure is indispensable for the functionality of the skeletal system as a whole (Godzinsky and Frank, 1990; Maroudas et al., 1985; Mow et al., 1990). Collagen, principally of types II, VI, IX and XI, is present in a non-soluble (polymerized) fibrillar form, although it can also exist in an unpolymerized, albeit fibril-associated, state (FAZIT-collagen) (Bruckner and Vanderrest, 1994), particularly type IX (Muller Glauser et al., 1986), and typically in adult tissue. Other collagen types (e.g., I, III and V) occur mainly in immature articular cartilage and at low levels. The intervening space between the collagen fibrils and fibres is occupied mainly by dermatan- and keratan-sulphatecontaining proteoglycans (Bayliss et al., 1983; Hardingham et al., 1992) and by fibromodulin (Hedbom and Heinegard, 1993; Hedlund et al., 1994). But a number of proteins and glycoproteins have also been identified within the extracellular matrix of mature articular cartilage. These include cartilage matrix protein (Hauser and Paulsson, 1994), cartilage oligomeric matrix protein, fibronectin, link proteins and osteonectin (Aeschlimann et al., 1995; Mundlos et al., 1992). As with the fibrillar collagens, these non-collagenous proteins and glycoproteins are distributed in a highly characteristic manner (Lorenzo et al., 1998; Wiberg et al., 2003). The extracellular matrix is generally recognized to be subdivided into three major compartments ± the pericellular, the territorial and the interterritorial (Fig. 3.2) ± which were first described by Meachim and Roy (1967) and Meachim and Stockwell (1973), and later characterized in detail by several groups of investigators (Eggli et al., 1985; Poole, 1993; Poole et al., 1982; Szirmai, 1963,

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3.1 Light micrograph of vertically-sectioned adult human articular cartilage (femoral condyle), illustrating its subdivision into superficial (S), transitional (T), upper radial (U), lower radial (L) and calcified cartilage (M) zones; the latter abuts on the subchondral bone plate (B). 100 m-thick saw-cut, surface-stained with basic Fuchsine, McNeal's Tetrachrome and Toluidine Blue O. Bar = 100 m (reproduced with permission from Hunziker, E.B. (1992) Articular cartilage structure in humans and experimental animals. In: Articular Cartilage and Osteoarthritis. K.E. Kuettner, R. Schleyerbach, J.G. Peyron and V.C. Hascall (eds.) Raven Press, New York, pp. 183±199).

1969; Szirmai and Doyle, 1961; Weiss et al., 1968). In these studies, the pericellular matrix coat was generally described to be free of fibrillar collagen (Eggli et al., 1988; Meachim and Roy, 1967; Weiss et al., 1968), but rich in soluble components, mainly proteoglycans. When articular cartilage is conventionally fixed in the absence of a cationic dye, as was the case in these former studies, proteoglycans are extracted from the pericellular matrix, which, as a consequence of the ensuing cell shrinkage, appears as an optically èmpty' lacuna (Davies et al., 1962; Freeman, 1973; Hunziker et al., 1982; Meachim and Roy, 1967). The pericellular space abuts on the territorial domain, which is characterized by a basket-like network of collagen fibrils that embraces not only individual chondrocytes, but also chondrocyte groups (chondrons) (Hunziker, 1992; Poole, 1993; Poole et al., 1987; Szirmai, 1963, 1969). The territorial


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3.2 Electron micrograph of a pair of chondrocytes (chondron) from the lower radial zone of adult human articular cartilage, processed by high-pressure freezing, freeze substitution and Epon embedding. At this relatively low level of magnification, the pericellular matrix compartment (PM) appears as a homogeneously stained mantle around each chondrocyte, in contrast to the territorial domain (TM), which forms a fibrillar coat around the chondron. Both of these compartments may vary considerably in width. The interterritorium (ITM), which is not clearly demarcated from the territorium, is generally distinguished from the latter by its higher proportion of parallel-oriented fibrils (see arrows): it occupies the bulk of the intercellular space. Arrowheads: fine cellular processes. Bar = 4 m (reproduced with permission from Hunziker et al., 1997).

matrix gives way to the interterritorial compartment, which constitutes the bulk of the extracellular space. It is characterized by a gradual increase in fibril diameter on moving away from any given chondrocyte (Bonucci et al., 1974; Davies et al., 1962; Dearden et al., 1974; Hedlund et al., 1994) and, on a more global basis, from the articular cartilage surface to the calcified tissue layer (spanning an eight- to ten-fold difference in the global case) (Davies et al., 1962; Ghadially, 1983; Hunziker, 1992). During the 1990s, improvements in cryotechnical tissue processing permitted a more precise and more detailed ultrastructural analysis of the extracellular matrix of articular cartilage (Hunziker et al., 1996, 1997; Studer et al., 1995, 1996). Using this methodology, it was possible to identify for the first time a network of cross-banded filaments, 10±15 nm in diameter, with a periodicity characteristic of collagen fibrils, throughout the truly vitrified substance (Fig. 3.3), even within the pericellular matrix compartment. This finding, which has been since neglected, should be of interest to tissue engineers, who are currently

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3.3 Electron micrograph of adult human articular cartilage, processed by highpressure freezing, freeze substitution and Epon embedding. Interterritorial matrix from the lower radial zone, illustrating part of a large longitudinallysectioned bundle of fibrils (LB), which traverse the picture diagonally from the lower left-hand edge to the upper right-hand one. The fibrillar units making up this bundle exhibit various diameters and are organized with a torsional twist around its longitudinal axis. The periodic banding of each fibril is aligned in register with that of its neighbour. A number of fine cross-banded filaments (F), with a diameter of 10±15 nm, can be seen dispersed throughout the entire extracellular space. Both filaments and collagen fibrils are mantled by an electron-lucent zone; this is most patent in cross-sectioned elements, where it is manifested as a halo, particularly when these units form bundles (demarcated by arrowheads). Bar = 0.5 m (reproduced with permission from Hunziker et al., 1997).

intent on demonstrating the usefulness of nanofibrillar meshworks in the engineering particularly of articular cartilage (Schindler et al., 2006), and they should bear in mind that nature has forestalled their `novel' idea. This fine crossbanded filamentous network has been identified in human as well as in bovine cartilage, and probably exists also in other mammalian species. Its existence adds to the complexity of the collagen architecture within articular cartilage (Fig. 3.4), and its presence has unknowingly influenced the conception of biomechanical models (Lai et al., 1993; Mow et al., 1980; Parsons and Black, 1977; Setton et al., 1993). The presence of this fine nanofibrillar meshwork may also help to account more satisfactorily for the diffusion characteristics of macromolecules within the extracellular space. Although the precise chemical composition of these filaments has yet to be determined, their cross-banded appearance and 67 nm periodicity afford strong indications that they are collagenous in nature (type-XI collagen is a very likely candidate). Other structural phenomena of collagen fibrils, which may be of relevance in a biomechanical context, include `kinking' and `brushing' (see Fig. 3.5). Also of relevance are variations in the internal structural make-up of individual fibres, which reflect differences in the types of collagen of which they are composed.


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3.4 Schematic representation of the matrix organization in adult articular cartilage. The pericellular compartment surrounds individual chondrocytes and is of variable width; it is free of fibrillar components but contains an abundance of isotropically arranged, cross-banded filaments which occur not only here but extend throughout the entire extracellular space. This domain abuts on the territorial compartment, which distinguishes the chondron as a distinct morphological entity. It contains a basket-like arrangement of collagen fibrils and, like the pericellular domain, is variable in width. The remaining, and bulk, portion of the extracellular space is referred to as the interterritorium; two subpopulations of fibrils and of fibril bundles are distinguishable here on the basis of their orientation: one exhibits the classic parallel arrangement, which distinguishes this compartment from the territorium and gives rise, on a broader scale, to the arcade-like architecture described by Benninghoff (1922, 1925); the other manifests a more random (isotropic) organization (reproduced with permission from Hunziker et al., 1997).

3.3

Dual function of immature articular cartilage during postnatal growth

During the postnatal development of a mammalian organism, the long bones undergo extensive growth and modelling in all regions, but particularly in the epiphysis. In the latter region, activities are governed by the joint cartilage, which thus acts not only as an articulating layer, but also as a surface growth plate for this portion of the bone during postnatal development (Fig. 3.6) (Carlsson et al., 1986). Hence, immature mammalian articular cartilage is sometimes referred to as an articular epiphyseal complex (Carlsson et al., 1985, 1986). The growth activities of the metaphysis and of the diaphysis are governed by the `true' growth plate, viz., by the physis (Hunziker et al., 1987). At the time of birth and during postnatal development, the layer of articular cartilage manifests a fairly isotropic structural organization (Hunziker et al.,

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3.5 Electron micrograph illustrating a portion of the interterritorial matrix compartment derived from the lower radial zone of adult human articular cartilage, processed by high-pressure freezing. Unstained ultra-thin cryosection. Cross-sectioned collagen fibrils (CF) exhibit variable size and shape. Their profile areas manifest a characteristic `staining' pattern, namely, a pale central region, a narrow, dark boundary and a perifibrillar electron-lucent halo, which features are more readily discerned in fibril bundles (demarcated by arrowheads). Longitudinally-sectioned fibrils, which exhibit a periodic banding of 67 nm, with a sub-banding of approximately 22.5 nm, sometimes exhibit brushing (Br), namely, a splitting up into finer elements; single fibrils are sometimes observed to undergo abrupt changes in their course, a phenomenon that is referred to as kinking (K). Note the occurrence of bubbling (B), which is induced by prolonged exposure of the section to the electron beam during microscopy. Magnification: 50 000. Insert: Electron-diffraction analysis of the section, revealing the central electron beam to be surrounded by diffuse concentric rings. This manifestation indicates that tissue water has been frozen in an amorphous (vitrified) state (reproduced with permission from Hunziker et al., 1997).

2007; Schenk et al., 1986), which bears little resemblance to that of the adult organism (Hunziker, 1992). In rabbits, the layer is already quite thick one month after birth (Hunziker et al., 2007), but the cells are distributed fairly randomly (Fig. 3.7a). Although the cells of the superficial zone tend to be oriented with their long axis running parallel to the articular cartilage surface, those of the underlying zones are arranged more isotropically. They exist singly or as small clusters, but with no preferential spatial orientation. During the ensuing (second) month, the structural organization of the articular cartilage layer does not change dramatically. However, the cells become more anisotropically arranged, and their numerical density increases (Fig. 3.7b). During the third postnatal month, which, in rabbits, marks the onset of puberty and the attainment of sexual maturity, a more dramatic change in the structural organization of the articular


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3.6 (a) Scheme representing postnatal growth of the epiphysis (E) of a long bone. The articular cartilage (A) acts not only as an articulating layer, but also as a surface growth plate for the longitudinal, radial and lateral growth of the epiphyseal bone (radially-oriented arrows). The true growth plate (G), which is located between the epiphysis (E) and the metaphysis (M), is responsible for the longitudinal growth of the metaphysis (M) and the diaphysis (D) (reproduced with permission from Hunziker et al., 2007). (b) Scheme illustrating the bidirectional replication of superficial-zone stem cells during the growth-activity phase of the articular cartilage layer. This slowly proliferating pool supplies daughter cells which can be displaced either horizontally (1) or vertically (2). Horizontally-displaced cells replenish the stem-cell pool and effect lateral growth of the articular cartilage layer. Vertically-displaced cells feed the rapidly proliferating pool of transitamplifying cells in the transitional and upper radial zones. These latter cells effect rapid clonal expansion in the vertical direction. Later, they hypertrophy and initiate matrix mineralization. Longitudinal bone growth is achieved both by rapid clonal expansion in the vertical direction and by cell hypertrophy (modified from Lavker and Sun, 2000) (reproduced with permission from Hunziker et al., 2007).

cartilage layer is apparent (Fig. 3.7c). The individual chondrocytes are highly oriented in space. Indeed, in the transitional zone and in the upper and lower radial zones, the anisotropy coefficient of the cells is comparable to that of chondrocytes in mature tissue (Fig. 3.8). A noteworthy change occurs also at the mineralization front. During the first and second postnatal months, only the longitudinal septa are mineralized (Fig. 3.9a). Hence, the region is still open. But at the end of the third month, by which time the animals have attained sexual maturity, the mineralization front is continuous, viz., the horizontal as well as the longitudinal septa are mineralized (Fig. 3.9b,c). This closure of the mineralization front coincides with the cessation of the growth activity of the articular cartilage layer. During the fourth to the eighth postnatal months, no noteworthy changes occur in the structural organization of the articular cartilage layer (Fig. 3.7d). At the ultrastructural level, the architecture of the collagen fibrils within the extracellular matrix changes in parallel with the temporal process of cellular

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3.7 Light micrographs of articular cartilage tissue derived from the medial femoral condyle of New Zealand white rabbits 1 month (a), 2 months (b), 3 months (c) and 8 months (d) after birth. The images illustrate the transition from an isotropic cellular organization 1 month after birth (a) to a highly anisotropic one by the third postnatal month (c). At this latter stage, the architecture resembles that in the adult animal (d). The change in structural organization is accompanied by a decrease in the overall height of the articular cartilage layer. One micrometre-thick sections stained with Toluidine Blue O. Scale bars: (a) 220 m; (b, c, d) 110 m (reproduced with permission from Hunziker et al., 2007).

reorganization. One month after birth, the fibrils are arranged randomly (isotropically) throughout the extracellular space (Fig. 3.10a) in all zones. By the end of the third month (Fig. 3.10b), and up until the eighth month (Fig. 3.10c), they are organized as in adult articular cartilage. Within the pericellular and the territorial matrix compartments, the fibrils are arranged in a basket-like fashion around the cells and cell groups. After short-term labelling of the articular cartilage layer at the different postnatal ages with tritiated thymidine, which tags solely the rapidly


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3.8 Graph depicting the anisotropy index (AQ) for articular cartilage chondrocytes (AQ cell diameter in the horizontal direction [D (hor)] divided by cell diameter in the vertical direction [D (vert)]) as a function of the tissue zone and the postnatal age of the New Zealand white rabbits. In the superficial zone, and especially in the transitional and upper radial zones, the anisotropy index decreases between the first and third postnatal months. The decrease in this quotient reflects the transition from an isotropic to an anisotropic organization of the cells. No further significant change in the anisotropy index occurs after the third postnatal month, which indicates that the mature structural organization of the chondrocytes is attained at this juncture (puberty). Mean values are represented together with the standard error of the mean (reproduced with permission from Hunziker et al., 2007).

proliferating cell pools, autoradiography reveals a positive reaction only in the transitional and the upper radial zones. Hence, it is within these zones that the cells undergo the rapid proliferation and clonal expansion that are required for the speedy growth and neoformation of articular cartilage tissue. The location of the slowly proliferating precursor cells, with an estimated cycling time of about 8 days, can be identified immunohistochemically after administering bromodeoxyuridine to the rabbits on a daily basis (via the drinking water) during the final 12 days prior to sacrifice. This analysis reveals a

3.9 Light micrographs of articular cartilage tissue derived from the medial femoral condyle of New Zealand white rabbits 1 month (a) and 3 months (b, c) after birth. Each image depicts the border between the hyaline articular cartilage layer and the mineralization front. One month after birth (a), this region is still open, since only the longitudinal septa are mineralized. The site is characterized by a high level of resorptive activity, as evidenced by the abundance of macrophages (M) and osteoclasts (O). By the third postnatal month (b), this resorption of the longitudinal (mineralized) septa has ceased. The mineralization front between hyaline and calcified cartilage (CC) is continuous (namely closed), since not only the longitudinal, but also the horizontal septa are mineralized. However, the calcified cartilage is subject to physiological remodelling by osteoclasts. The characteristically undulating course of the calcified cartilage layer (seen at higher magnification in (c)) is believed to improve its mechanical anchorage within the subchondral bone plate (Broom and Pole, 1982; Keinan-Adamsky et al., 2005). RC osteoclastic resorption channel; T bone tissue; arrowheads bone-cartilage interface. One micrometre-thick sections stained with Toluidine Blue O. Scale bars: (a, b) 30 m; (c) 15 m (reproduced with permission from Hunziker et al., 2007).


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3.10 Electron micrographs of chondrocytes within the radial zone of articular cartilage tissue derived from the medial femoral condyle of New Zealand white rabbits 1 month (a), 3 months (b) and 8 months (c) after birth. In 1-month-old rabbits (a), the collagen fibrils are distributed randomly (isotropically) and homogeneously throughout the entire extracellular space. No distinct compartmentalization of the matrix is apparent. In 3- (b) and 8-month-old (c) rabbits, the extracellular space is organized into distinct pericellular (P), territorial (T) and interterritorial (I) compartments. The narrow pericellular matrix compartment is highly electron-dense owing to the abundance of ruthenium-hexaamine-trichloride-precipitated proteoglycans. Within the territorial compartment, the collagen fibrils are arranged in a typically basketlike fashion around the cells. Within the interterritorium, which constitutes the bulk of the extracellular space, the collagen fibrils are oriented parallel to each other and in a predominantly longitudinal direction. Scale bars: (a) 10 m; (b, c) 5 m (reproduced with permission from Hunziker et al., 2007).

positive reaction within all cell nuclei of all zones, including the superficial one. Hence, the slowly proliferating pool of precursor cells (which resemble stem cells in this respect) is located within the superficial zone, in analogy to the resting zone (referred to in the older literature as the stem-cell zone) of a true growth plate (Hunziker et al., 1987; Kember, 1960).

3.4

Physiological mechanism underlying the evolution of a mature from an immature articular cartilage structure

Mature articular cartilage is characterized by a high degree of structural anisotropy, its cells being organized into well-defined vertical columns and horizontal septa, whereas immature articular cartilage is more isotropic in structure (Fig. 3.11). The mechanism underlying the evolution of the mature from the immature architecture has been only recently elucidated (Huniker et al., 2007). It was hypothesized that the articular cartilage layer of synovial joints underwent structural reorganization either by a process of internal tissue remodelling, or by one of controlled tissue resorption (at the vascular invasion front) that was synchronized with tissue renewal (on the basis of the proliferative activity of precursor cells within the superficial zone). The daily growth rate of the epiphyseal bone was determined in rabbits at monthly intervals from the first to the eighth postnatal months (Huniker et al., 2007) according to the tetracycline-labelling principle (Hulth and Olerud, 1962). The daily growth rate

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3.11 Schemes illustrating the structure of immature (left-hand picture) and mature (right-hand picture) articular cartilage tissue. During foetal and early postnatal life, the chondrocytes are distributed randomly (isotropically) in space. However, a vectoral gradient in cell size and shape is apparent: near the articular surface, the cells are small and horizontally flattened; with increasing depth, they become larger, rounder and ultimately (near the vascular invasion front) irregular in form. In adult articular cartilage, the chondrocytes are organized anisotropically into distinct vertical columns and horizontal zones (superficial/tangential, transitional, upper and lower radial, and calcified). Our data reveal this structural transformation to be achieved not by a process of internal remodelling, but by the resorption and neoformation of tissue. Arrows indicate the level of the mineralization front (namely, the tidemark) (reproduced with permission from Hunziker et al., 2007).

was highest 1 month after birth and decreased almost exponentially thereafter until the time of sexual maturity (between the third and fourth postnatal months). After the fourth month, growth activity ceased altogether (Fig. 3.12). The height of the articular cartilage layer (Fig. 3.13) decreased in parallel with the decrease in bone growth rate. The total bone length gain achieved during each of the first five postnatal months decreased with time (Fig. 3.14). However, during the growth phase (i.e., during the first three postnatal months), the monthly gain in bone length exceeded the height of the articular cartilage layer (from which that of the superficial zone had been subtracted). This finding indicates that immature articular cartilage does not become reorganized by a process of internal tissue remodelling. If this were the case, then the total bone length gain per month during the growth phase would be smaller than the corresponding height of the articular cartilage layer (after subtracting that of the superficial zone). Hence, immature articular cartilage must be completely resorbed and replaced by new tissue. The existing tissue is destroyed at the vascular invasion front. The superficial zone alone is spared, and serves as the source of the precursor cells which give rise to new tissue. One of the implications of these findings is that the postnatal structural reorganization of articular cartilage tissue occurs not at a fixed topographical location in space, but by a process of growth involving an elongation of the underlying bone. This circumstance is of importance in the engineering of articular cartilage within adult organisms. Adult articular cartilage lesions would be ideally repaired by tissue that manifests a high degree of structural anisotropy


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3.12 Graph depicting the daily growth rate of the epiphyseal bone as a function of the postnatal age of the New Zealand white rabbits. The growth rate declines precipitously up to the third postnatal month; by the fourth month, growth activity has ceased altogether. Mean values are represented together with the standard error of the mean (reproduced with permission from Hunziker et al., 2007).

3.13 Graph depicting the overall height of the articular cartilage layer as a function of the postnatal age of the New Zealand white rabbits. The height of the articular cartilage layer decreases precipitously and almost linearly up to the third postnatal month, at which juncture the mature structural organization of the tissue is achieved and the animals attain sexual maturity. Thereafter (between 4 and 8 months), the height of the articular cartilage layer does not change significantly. Mean values are represented together with the standard error of the mean (reproduced with permission from Hunziker et al., 2007).

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3.14 Bar graph comparing the net monthly length-gain in the epiphyseal bone (black columns) with the height of the articular cartilage layer from which that of the superficial zone has been substracted (grey columns) at each postnatal month. During the first 2 months, when the articular cartilage layer is undergoing structural reorganization, the net length-gain in the epiphyseal bone exceeds the height of the articular cartilage layer. This finding indicates that the articular cartilage layer is structurally reorganized not by a process of internal remodelling, but by the resorption of all zones except the superficial (stem-cell) one and their neoformation by appositional growth of the latter. If a process of internal remodelling were involved, the height of the articular cartilage layer (excluding the superficial zone) would exceed the net monthly length-gain in the epiphyseal bone. Mean values are represented together with the standard error of the mean (reproduced with permission from Hunziker et al., 2007).

from the very onset of the healing process, in order to ensure its longevity and mechanical competence (Wong et al., 1997). However, current cell-based approaches involve the implantation of immature cartilage, with a random distribution of cells (for review, see Hunziker, 2002). Since the structural reorganization of immature cartilage under physiological conditions does not take place at a fixed topographical position in space, it is highly unlikely that the current tissue-engineering approaches will lead to optimal repair results.

3.5

Inter-species differences in articular cartilage structure, and structure±function correlations in humans

A quantitative description of the three-dimensional structures (Cruz Orive and Hunziker, 1986) that make up the mature human articular cartilage layer is essential for a thorough understanding of its biochemical, biophysical and biomechanical properties, which determine its physiological functions. Such


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data are also necessary for the rational development of cartilage-engineering strategies. Furthermore, structural differences between mammalian species have to be borne in mind when selecting an animal model for the in-vivo testing of a potential repair strategy for human use. Currently, baseline structural data are available for only a few human joint types; most especially for the medial femoral condyle of young and middle-aged adults with no known pathological affections or history of joint disease (Hunziker et al., 2002; Shepherd and Seedhom, 1999). In this joint, the mean heights of the layers of hyaline and calcified cartilage, and of the subchondral bone plate, are 2±4 mm (coefficient of error: 22%), 0.13 mm (coefficient of error: 74%) and 0.19 mm (coefficient of error: 57%), respectively. In small and middle-sized experimental mammals, namely, in rabbits, sheep and goats, the hyaline cartilage layer in the corresponding joint is much thinner, viz., 0.25, 0.4 and 0.9 mm, respectively (Hulth and Olerud, 1962; Hunziker, 1999; Masoud et al., 1986). However, in larger mammals, such as bovine cows and horses, it is not thicker but of similar height. The calcified cartilage layer and the subchondral bone plate are, on the other hand, much thinner in humans than they are in rabbits, goats and sheep (Hulth and Olerud, 1962; Masoud et al., 1986). As may be seen from the data cited above, in the adult human medial femoral condyle, the heights of the calcified cartilage layer and the subchondral bone plate are characterized by very high coefficients of error. These great intra- and inter-individual variations around the mean heights reflect the undulating upperand lower-surface profiles of the calcified cartilage layer, which in turn reflect its interdigitation with the overlying hyaline cartilage layer and the underlying subchondral bone plate, and which facilitate anchorage. With respect to the junction between the hyaline and the calcified cartilage layers, this undulating profile could also enhance the diffusion of nutrients (from the former to the latter stratum). The volume density of chondrocytes within the articular cartilage layer of the adult human medial femoral condyle is overall very low (mean: 0.65%; coefficient of error: 9%), and decreases with increasing distance from the surface. Between the superficial zone and the lower portion of the lower radial zone, it drops by a factor of two (from 2.6% to 1.2%). Such low values have not been encountered in any other mammalian species. In rabbits and goats, for example, the overall value is about 12% (Hulth and Olerud, 1962; Hunziker, 1999). Among human bodily tissues, articular cartilage is unique in having such a low volume density of cells. In simple terms, the low volume density of chondrocytes indicates that the cells are very sparsely distributed, which in turn implies that the metabolic and synthetic activities of any given chondrocyte sustain a very large domain of the extracellular matrix. Given that articular cartilage is avascular, and that the chondrocytes thus depend upon the diffusion of oxygen and nutritients over a great distance (from blood capillaries within the synovium and the subchondral bone plate), it is not surprising that the metabolic activities of the cells are conducted chiefly along anaerobic pathways (Wong

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and Hunziker, 2001). The volume of the extracellular domain that is sustained by a single chondrocyte is 104 040 mm3 (coefficient of error: 4.9%) for the articular cartilage layer as a whole, which rises to 106 707 mm3 for the lower portion of the upper radial zone (Hunziker et al., 2002). These values are about ten-fold higher than those for the articular cartilage layer of the medial femoral condyle in rabbits (Eggli et al., 1988). And in the growth plate, such huge matrix domains have not been described for any mammalian species (Cruz Orive and Hunziker, 1986; Hunziker et al., 1987), including humans. Their great compass in human articular cartilage has an impact on the remodelling activity of the outskirts of the interterritorium, which is very low. The structure of the articular cartilage layer changes in response to the mode of biomechanical loading. A sustained increase in load accelerates tissue remodelling, which leads to a dramatic thinning of the articular cartilage layer (Vanwanseele et al., 2002). However, sustained decreases in load lead to only a minimal thickening of the layer (Eckstein et al., 2002; Shepherd and Seedhom, 1999). The adaptive potential of articular cartilage is probably limited by cellscale biophysical considerations. Since the activity of chondrocytes depends greatly on the diffusion of solutes through the avascular extracellular matrix, their metabolism is closely coupled with the local transport of oxygen and nutrients, which is partly governed by the organization of the matrix (Maroudas, 1975). This tenet is borne out by the finding that sustained loading of a weightbearing joint, which presumably induces local changes in the mechanical properties of the articular cartilage tissue, can give rise to highly circumscribed osteoarthritic lesions (Buckwalter and Mankin, 1998; Froimson et al., 1997; Shepherd and Seedhom, 1999). Furthermore, the metabolic activities of chondrocytes (Wong et al., 1996) and the biomechanical properties of the matrix (Chen et al., 2001) are known to vary between zones, and these differences represent the functional correlates of differences in structure. However, the thickness of human knee-joint cartilage varies between anatomical locations in a manner that is seemingly independent of the structural organization of the cells and the matrix (Quinn et al., 2005). Anatomical variations in knee-joint cartilage appear to be governed by factors such as the degree of congruency between the apposing bony elements (Simon et al., 1973) and topographical relationships with other tissues, for example, the meniscus (Shepherd and Seedhom, 1999; Vanwanseele et al., 2002). However, as aforeindicated, cartilage thickness is not trivially related to load bearing. A sustained increase in load accelerates tissue remodelling. This response can result in a decrease in joint surface area (Eckstein et al., 2002) and in site-specific differences in the mechanical properties of the articular cartilage (Froimson et al., 1997; Shepherd and Seedhom, 1999). The uniformity of cartilage structure in different anatomical regions of the knee joint, with different functional needs, indicates the existence of fundamental cell-scale constraints. For example, the metabolism of a given cell must be augmented if the matrix volume that it controls increases


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(assuming a constant rate of turnover); but it cannot do so indefinitely, owing to limitations in the transport of solutes both to and from the cell. A balance must be struck between decreased solute transport and increased mechanical stiffness, which is effected by an increase in matrix density. These cell-scale constraints may account for the observation that an increase in cartilage thickness is associated with a decrease in its mechanical stiffness (Froimson et al., 1997; Lyyra et al., 1999; Shepherd and Seedhom, 1999). The decrease in the cellularity of human articular cartilage that occurs with age (Mow et al., 1993) may contribute to its diminishing capacity for repair, since fewer cells govern an increasingly expanding volume of matrix. However, not only the numerical density of cells but also their organization influences the mechanical stiffness of articular cartilage: although foetal cartilage is characterized by a higher volume density of cells than is mature cartilage, its cells and matrix are organized isotropically, whereas in mature cartilage, they are arranged anisotropically; but the latter is mechanically stiffer than the former (Wong and Hunziker, 2001).

3.6

References

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Cruz Orive, L. M., and Hunziker, E. B. Stereology for anisotropic cells: application to growth cartilage. J Microsc 143: 47±80; 1986. Davies, D. V., Barnett, C. H., Cochrane, W., and Palfrey, A. J. Electron microscopy of articular cartilage in the young adult rabbit. Ann Rheum Dis 21: 11±22; 1962. Dearden, L. C., Bonucci, E., and Cuicchio, M. An investigation of ageing in human costal cartilage. Cell Tissue Res 152: 305±337; 1974. Eckstein, F., Faber, S., Muhlbauer, R., Hohe, J., Englmeier, K. H., Reiser, M., and Putz, R. Functional adaptation of human joints to mechanical stimuli. Osteoarthritis Cartilage 10: 44±50; 2002. Eggli, P. S., Herrmann, W., Hunziker, E. B., and Schenk, R. K. Matrix compartments in the growth plate of the proximal tibia of rats. Anat Rec 211: 246±257; 1985. Eggli, P. S., Hunziker, E. B., and Schenk, R. K. Quantitation of structural features characterizing weight- and less-weight-bearing regions in articular cartilage: a stereological analysis of medial femoral condyles in young adult rabbits. Anat Rec 222: 217±227; 1988. Freeman, M. A. R. Adult Articular Cartilage. London: Pitman Medical; 1973. Froimson, M. I., Ratcliffe, A., Gardner, T. R., and Mow, V. C. Differences in patellofemoral joint cartilage material properties and their significance to the etiology of cartilage surface fibrillation. Osteoarthritis Cartilage 5: 377±386; 1997. Ghadially, F. N. Fine Structure of Synovial Joints. A Text and Atlas of the Ultrastructure of Normal and Pathological Articular Tissues. London: Butterworth & Co. (Publishers) Ltd; 1983. Grodzinsky, A. J., and Frank, E. H. Electromechanical and physicochemical regulation of cartilage strength and metabolism. In: D. W. L. Hukins (ed.), Connective Tissue Matrix Part 2, pp. 91±126. London: The Macmillan Press Ltd; 1990. Hardingham, T. E., Fosang, A. J., and Dudhia, J. Aggrecan: the chondroitin sulfate/ keratan sulfate proteoglycan from cartilage. In: K. E. Kuettner, R. Schleyerbach, J. G. Peyron, and V. C. Hascall (eds.), Articular Cartilage and Osteoarthritis, pp. 5± 20. New York: Raven Press; 1992. Hauser, N., and Paulsson, M. Native cartilage matrix protein (CMP) ± a compact trimer of subunits assembled via a coiled-coil alpha-helix. J Biol Chem 269: 25747±25753; 1994. Hedbom, E., and Heinegard, D. Binding of fibromodulin and decorin to separate sites on fibrillar collagens. J Biol Chem 268: 27307±27312; 1993. Hedlund, H., Mengarelliwidholm, S., Heinegard, D., Reinholt, F. P., and Svensson, O. Fibromodulin distribution and association with collagen. Matrix Biol 14: 227±232; 1994. Hulth, A., and Olerud, S. Tetracycline labelling of growing bone. Acta Soc Med Ups 67: 219±231; 1962. Hunziker, E. B. Articular cartilage structure in humans and experimental animals. In: K. E. Kuettner, R. Schleyerbach, J. G. Peyron, and V. C. Hascall (eds.), Articular Cartilage and Osteoarthritis, pp. 183±199. New York: Raven Press; 1992. Hunziker, E. B. Biologic repair of articular cartilage. Defect models in experimental animals and matrix requirements. Clin Orthop Relat Res, S135±S146; 1999. Hunziker, E. B. Articular cartilage repair: basic science and clinical progress. A review of the current status and prospects. Osteoarthritis Cartilage 10: 432±463; 2002. Hunziker, E. B., Herrmann, W., and Schenk, R. K. Improved cartilage fixation by ruthenium hexammine trichloride (RHT). A prerequisite for morphometry in growth cartilage. J Ultrastruct Res 81: 1±12; 1982. Hunziker, E. B., Schenk, R. K., and Cruz-Orive, L. M. Quantitation of chondrocyte


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performance in growth-plate cartilage during longitudinal bone growth. J Bone Joint Surg Am 69: 162±173; 1987. Hunziker, E. B., Wagner, J., and Studer, D. Vitrified articular cartilage reveals novel ultra-structural features respecting extracellular matrix architecture. Histochem Cell Biol 106: 375±382; 1996. Hunziker, E. B., Michel, M., and Studer, D. Ultrastructure of adult human articular cartilage matrix after cryotechnical processing. Microsc Res Tech 37: 271±284; 1997. Hunziker, E. B., Quinn, T. M., and Hauselmann, H. J. Quantitative structural organization of normal adult human articular cartilage. Osteoarthritis Cartilage 10: 564±572; 2002. Hunziker, E. B., Kapfinger, E., and Geiss, J. The structural architecture of adult mammalian articular cartilage evolves by a synchronized process of tissue resorption and neoformation during postnatal development. Osteoarthritis Cartilage 15: 403±413; 2007. Keinan-Adamsky, K., Shinar, H., and Navon, G. The effect of detachment of the articular cartilage from its calcified zone on the cartilage microstructure, assessed by 2Hspectroscopic double quantum filtered MRI. J Orthop Res 23: 109±117; 2005. Kember, N. F. Cell division in endochondral ossification. A study of cell proliferation in rat bones by the method of tritiated thymidine autoradiography. J Bone Joint Surg (Br) 42: 824±839; 1960. Lai, W. M., Mow, V. C., and Zhu, W. B. Constitutive modeling of articular cartilage and biomacromolecular solutions. J Biomech Eng 115: 474±480; 1993. Lavker, R. M., and Sun, T. T. Epidermal stem cells: properties, markers, and location. Proc Natl Acad Sci USA 97: 13473±13475; 2000. Lorenzo, P., Bayliss, M. T., and Heinegard, D. A novel cartilage protein (CILP) present in the mid-zone of human articular cartilage increases with age. J Biol Chem 273: 23463±23468; 1998. Lyyra, T., Kiviranta, I., Vaatainen, U., Helminen, H. J., and Jurvelin, J. S. In vivo characterization of indentation stiffness of articular cartilage in the normal human knee. J Biomed Mater Res 48: 482±487; 1999. Maroudas, A. Biophysical chemistry of cartilaginous tissues with special reference to solute and fluid transport. Biorheology 12: 233±248; 1975. Maroudas, A., Ziv, I., Weisman, N., and Venn, M. Studies of hydration and swelling pressure in normal and osteoarthritic cartilage. Biorheology 22: 159±169; 1985. Masoud, I., Shapiro, F., Kent, R., and Moses, A. A longitudinal study of the growth of the New Zealand white rabbit: cumulative and biweekly incremental growth rates for body length, body weight, femoral length, and tibial length. J Orthop Res 4: 221± 231; 1986. Meachim, G., and Roy, S. Intracytoplasmic filaments in the cells of adult human articular cartilage. Ann Rheum Dis 26: 50±58; 1967. Meachim, G., and Stockwell, R. A. The matrix. In: M. A. R. Freeman (ed.), Adult Articular Cartilage, pp. 1±50. London: Pitman Medical; 1973. Mow, V. C., Kuei, S. C., Lai, W. M., and Armstrong, C. G. Biphasic creep and stress relaxation of articular cartilage in compression: theory and experiments. J Biomech Eng 102: 73±84; 1980. Mow, V. C., Ratcliffe, A., and Woo, S. L. Y. Biomechanics of Diarthrodial Joints. New York: Springer-Verlag; 1990. Mow, V. C., Ateshian, G. A., and Spilker, R. L. Biomechanics of diarthroidal joints ± a review of 20 years of progress. J Biomech Eng 115: 460±467; 1993.

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Muller Glauser, W., Humbel, B., Glatt, M., Strauli, P., Winterhalter, K. H., and Bruckner, P. On the role of type IX collagen in the extracellular matrix of cartilage: type IX collagen is localized to intersections of collagen fibrils. J Cell Biol 102: 1931±1939; 1986. Mundlos, S., Schwahn, B., Reichert, T., and Zabel, B. Distribution of osteonectin mRNA and protein during human embryonic and fetal development. J Histochem Cytochem 40: 283±291; 1992. Parsons, J. R., and Black, J. The viscoelastic shear behavior of normal rabbit articular cartilage. J Biomech 10: 21±29; 1977. Poole, A. R., Pidoux, I., Reiner, A., and Rosenberg, L. An immunoelectron microscope study of the organization of proteoglycan monomer, link protein, and collagen in the matrix of articular cartilage. J Cell Biol 93: 921±937; 1982. Poole, C. A. The structure and function of articular cartilage matrices. In: J. F. J. Woessner and D. S. Howell (eds.), Joint Cartilage Degradation. Basic and Clinical Aspects, pp. 1±35. New York: Marcel Dekker, Inc.; 1993. Poole, C. A., Flint, M. H., and Beaumont, B. W. Chondrons in cartilage: ultrastructural analysis of the pericellular microenvironment in adult human articular cartilages. J Orthop Res 5: 509±522; 1987. Quinn, T. M., Hunziker, E. B., and Hauselmann, H. J. Variation of cell and matrix morphologies in articular cartilage among locations in the adult human knee. Osteoarthritis Cartilage 13: 672±678; 2005. Schenk, R. K., Eggli, P. S., and Hunziker, E. B. Articular cartilage morphology. In: K. Kuettner, R. Schleyerbach, and V. C. Hascall (eds.), Articular Cartilage Biochemistry, pp. 3±22. New York: Raven Press; 1986. Schindler, M., Nur, E. K. A., Ahmed, I., Kamal, J., Liu, H. Y., Amor, N., Ponery, A. S., Crockett, D. P., Grafe, T. H., Chung, H. Y., Weik, T., Jones, E., and Meiners, S. Living in three dimensions: 3D nanostructured environments for cell culture and regenerative medicine. Cell Biochem Biophys 45: 215±227; 2006. Setton, L. A., Zhu, W. B., and Mow, V. C. The biphasic poroviscoelastic behavior of articular cartilage ± role of the surface zone in governing the compressive behavior. J Biomech 26: 581±592; 1993. Shepherd, D. E., and Seedhom, B. B. Thickness of human articular cartilage in joints of the lower limb. Ann Rheum Dis 58: 27±34; 1999. Simon, W. H., Friedenberg, S., and Richardson, S. Joint congruence. A correlation of joint congruence and thickness of articular cartilage in dogs. J Bone Joint Surg Am 55: 1614±1620; 1973. Studer, D., Michel, M., Wohlwend, M., Hunziker, E. B., and Buschmann, M. Vitrification of articular cartilage by high-pressure freezing. J Microsc 179: 321±332; 1995. Studer, D., Chiquet, M., and Hunziker, E. B. Evidence for a distinct water rich layer surrounding collagen II fibrils in articular cartilage extracellular matrix. J Structur Biol 117: 81±85; 1996. Szirmai, J. A. Quantitative approaches in the histochemistry of mucopolysaccharides. J Histochem Cytochem 11: 24±34; 1963. Szirmai, J. A. Structure of cartilage. In: A. Engel and T. Larsson (eds.), Thule International Symposium on Aging of Connective and Skeletal Tissue, pp. 163±184. Stockholm: Nordiska Boekhandelns; 1969. Szirmai, J. A., and Doyle, J. Quantitative histochemical and chemical studies on the composition of cartilage. J Histochem Cytochem 9: 611±618; 1961. Vanwanseele, B., Eckstein, F., Knecht, H., Stussi, E., and Spaepen, A. Knee cartilage of spinal cord-injured patients displays progressive thinning in the absence of normal


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joint loading and movement. Arthritis Rheum 46: 2073±2078; 2002. Weiss, C., Rosenberg, L., and Helfet, A. J. An ultrastructural study of normal young adult human articular cartilage. J Bone Joint Surg (Am) 50: 663±674; 1968. Wiberg, C., Klatt, A. R., Wagener, R., Paulsson, M., Bateman, J. F., Heinegard, D., and Morgelin, M. Complexes of matrilin-1 and biglycan or decorin connect collagen VI microfibrils to both collagen II and aggrecan. J Biol Chem 278: 37698±37704; 2003. Wong, M., and Hunziker, E. B. Articular cartilage biology and biomechanics. In: C. Erggelet and M. Steinwachs (eds.), Gelenkknorpeldefekte, pp. 15±28. Darmstadt: Steinkopff Verlag; 2001. Wong, M., Wuethrich, P., Eggli, P., and Hunziker, E. B. Zone-specific cell biosynthetic activity in mature bovine articular cartilage: a new method using confocal microscopic stereology and quantitative autoradiography. J Orthop Res 14: 424± 432; 1996. Wong, M., Wuthrich, P., Buschmann, M. D., Eggli, P., and Hunziker, E. B. Chondrocyte biosynthesis correlates with local tissue strain in statically compressed adult articular cartilage. J Orthop Res 15: 189±196; 1997.

4

Measuring the biomechanical properties of cartilage cells D . L . B A D E R and M . M . K N I G H T , Queen Mary University of London, UK

Abstract: Current techniques are described to deform chondrocytes and derive both quasi-static and viscoelastic parameters. The influence of intracellular structures, such as cytoskeletal elements and the nucleus, on these parameters is highlighted. The chapter also describes the metabolic response of chondrocytes to biomechanical conditioning. Output parameters, including cell proliferation and matrix synthesis, are estimated with respect to cell source, scaffolds and culture conditions. Key words: chondrocytes, loading techniques, cell mechanics, biomechanical conditioning, metabolic response.

4.1

Introduction

Mechanical loading is essential for the development, health and homeostasis of articular cartilage. This occurs primarily through a process of cellular mechanotransduction whereby the chondrocytes sense and respond to their mechanical environment by modulating the synthesis and catabolism of the extracellular matrix. Normal physiological loading of articular cartilage produces depthdependent compression of the tissue with associated deformation of the chondrocytes. Whilst the precise process of cellular mechanotransduction is as yet unclear, cell deformation is believed to be one of the primary stimuli. Hence it is of major importance to elucidate the deformation behaviour of cartilage cells in health and disease, which is governed by the inherent cellular and subcellular biomechanics. Measurements of cellular biomechanical parameters generally involve deformation of the cell surface, at least in part, by a known force or stress and simultaneous visualisation and measurement of the resulting cell deformation. Alternatively, a prescribed cellular deformation may be applied and the resulting force measured. Both approaches enable the force±displacement (F±x) relationship to be plotted for a single living cell, from which the apparent stiffness (k) may be calculated from the resulting gradient (k F=x). Frequently experimental data is combined with some sort of theoretical or computational model in order to derive a fundamental cell modulus. However, it is also


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necessary to take into account the fact that living cells exhibit characteristic viscoelastic behaviour. Hence any calculated cellular modulus is highly dependent on the temporal conditions of measurement. Finally, the process of mechanotransduction means that chondrocytes themselves actively respond to mechanical loading by remodelling intracellular structures resulting in changes in both cellular and intracellular biomechanics. Indeed it has been suggested that cells show behaviour similar to Wolf's Law in bone, namely that the biomechanical remodelling occurs in response to altered mechanical loading. There are numerous techniques to impart some form of deformation on living cells whilst simultaneously enabling direct microscopic visualisation and/or measurement of cell deformation (for review see Bader and Knight, 2008). This chapter reviews those techniques primarily used for analysing the biomechanics of chondrocytes. In addition, there have been many studies describing specialist loading rigs and bioreactor systems designed to apply in vitro mechanical stimuli to a variety of cell types in order to study their metabolic or injury responses (for review see Brown, 2000). This is further explored with reference to chondrocytes in the second part of this chapter, with particular emphasis on the influence of loading regimens, cell source and scaffold materials.

4.2

Measurement of chondrocyte biomechanics

4.2.1

Chondrocyte biomechanics in situ within cartilage explants

Various studies have investigated the chondrocyte deformation in cartilage explants subjected to mechanical loading in the form of compression or indentation. Consequently, a variety of loading rigs have been developed. These systems, such as the one shown schematically in Fig. 4.1, typically mount upon the stage of an inverted microscope enabling simultaneous visualisation of the resulting deformation to the tissue and cells. By visualising the tissue in both the unstrained and strained state and using the cells as displacement markers it is possible to calculate the local strain fields in terms of compressive, tensile and shear strains (Guilak et al., 1995; Schinagl et al., 1996). It is clear that the depthdependent mechanical properties of articular cartilage results in heterogeneous levels of local deformation, such that the local tissue strains will inevitably differ from the gross strain. Furthermore the local strain also differs from the cellular strain due to the mechanical properties of the pericellular matrix (PCM) associated with the chondrocyte, a functional unit termed the chondron (Poole et al., 1987). Cartilage deformation is transferred to the chondrocytes in two ways: (i) directly, via the deformation of the extracellular matrix (ECM), and (ii) indirectly, due to the compression-induced increase in extracellular osmolarity which produces a reduction in cell volume (Chao et al., 2005; Erickson et al.,

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2003; Hing et al., 2002). It is also possible that the compression-induced loss of cell volume in situ may be the result of mechanical deformation squeezing fluid from the cell, although this has not yet been established. In addition fluid flow during cartilage loading may also generate shear forces, which will impart a more subtle deformation at the cell surface. However, in order to determine cellular biomechanical properties based on cell deformation behaviour in cartilage explants, computational models have to be developed in which the physical properties of the cell can be adjusted to match the experimental data.


4.2.2

109

Micropipette aspiration

The most published approach for measuring chondrocyte biomechanics is that of micropipette aspiration, which is used to deform the membrane of the cell with a known suction pressure (for review see Hochmuth, 2000; Sato et al., 1987). The micropipettes, with inner diameters typically between 5 and 15 m, are coated with a silicone solution (Sigmacote, Sigma, MO) to prevent cell adhesion. A cell chamber containing approximately 1 mL of cell suspension permits the entry of the micropipette through a side wall. The micropipette is fixed to the stage of an inverted microscope and connected to a reservoir as shown in Fig. 4.1(a). Both the micropipette and the pressure control system need to be filled with a physiological saline solution, such as phosphate buffered saline (PBS). With the control of a hydraulic micro-manipulator (e.g. MO-203, Narishige, Tokyo, Japan), the micropipette is moved into contact with the cell surface and a tare pressure (typically 0.01 kPa) is applied to draw the chondrocyte against the mouth of the micropipette and to define the reference position. Aspiration pressure is then applied in a series of step increments, typically up to 5 cm H2O (0.49 kPa). At each pressure increment the cell is allowed to equilibrate for about 60±120 seconds, before a brightfield microscopy image reveals the extent of the cell deformation into the micropipette. This deformation is quantified by the aspirated length, L. A typical series of brightfield images of an isolated cartilage cell subjected to incremental levels of aspiration pressure is shown in Fig. 4.1(b)

4.1 (opposite) Micropipette aspiration. (a) Schematic diagram illustrating the micropipette aspiration system for quantifying cellular biomechanics. Suction pressure is applied to an isolated chondrocyte via a micropipette, by lowering a fluid-filled reservoir. A manometer or pressure sensor is used to provide a reading of the applied aspiration pressure. The syringes are used to fill the silicone tubing and to provide high positive pressure for releasing cells from the pipette. Cells are visualised during the aspiration process using an inverted microscope which may be connected to a confocal system (PBS phosphate buffered solution). (b) Representative brightfield and corresponding confocal microscopy images of a single isolated chondrocyte visualised during micropipette aspiration at pressures of 0, 1, 2, 3 and 4 cm of water. Scale bar indicates 5 m. Arrows indicate the aspiration of the cell into the micropipette. The cell was transfected with enhanced green fluorescent protein (eGFP) actin to examine mechanically induced changes in actin remodelling. (c) Corresponding plot of aspiration length, L, normalised to pipette radius, a, and plotted against aspiration pressure. A linear model has been fitted to the data with a gradient of 0.053 from which the cellular Young's modulus may be estimated at a value of 0.2 kPa. (d) Transient viscoelastic behaviour of a single chondrocyte immediately following a step increase in aspiration pressure to approximately 3.5 cm of water (data from Trickey et al., 2000). A theoretic model has been fitted to the data based on equation 4.2, thereby enabling the calculation of the instantaneous modulus, Ei, and the relaxation modulus, Er, at 0.41 and 0.24 kPa respectively (see text for details).

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alongside the corresponding confocal fluorescence images showing deformation of the actin cytoskeleton labelled using eGFP (enhanced green fluorescent protein) transfection. Prior to aspiration, the initial diameter of the chondrocye is measured, as well as the pipette inner diameter using calibrated brightfield microscopy. However, it is necessary to compensate for the distortion caused by the refractive index mismatch between the micropipette glass and the surrounding buffer solution. This is best achieved by multiplying the measured micropipette diameter by a factor of 0.92 (EngstroÈm et al., 1992). On the basis of the experimental data, the apparent Young's modulus can be determined using a theoretical elastic model previously developed to analyse the material properties of endothelial cells (Theret et al., 1988). In this model, the cell is assumed to be a homogeneous, elastic half-space material and the Young's modulus, E, is therefore given as: 3 P E 4:1 2 L=a where a and b are the inner and outer radii of the micropipette and is defined as the wall function with b ÿ a=a. This so-called `rigid punch model' employs a boundary condition of no axial displacement of the cell at the micropipette mouth, corresponding to 2:1 for the practical range used in a typical study. The Young's modulus can be determined from the slope of the linear regression of the normalised length L=a versus the negative pressure P as shown in Fig. 4.1(c). Further models and experimental approaches have been developed to calculate the viscoelastic properties of chondrocytes based on an analytical solution of micropipette aspiration (Sato et al., 1987). For this technique, the chondrocyte is aspirated into the micropipette with a single step aspiration pressure ranging from approximately 1 to 10 cm H2O. The aspiration length is then recorded over a time period of up to 300 seconds (Fig. 4.1d). The associated model assumes that the chondrocyte behaves as a homogeneous linear viscoelastic three-parameter solid half-space. Using this model the aspiration length, L, and the relaxation constant, can be predicted at time, t, based on the following equation: aP k2 4:2 1ÿ et= Lt k1 k2 k1

k1 k2 k1 k2

4:3

The viscoelastic parameters k1 , k2 and can be calculated by fitting experimental aspiration length data to equation 4.2 using a non-linear regression. The parameter k1 is termed the equilibrium or relaxation modulus (Er or E1 ), k1 k2 is the instantaneous modulus (Ei) and is the apparent viscosity. Using


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this viscoelastic model it is also possible to determine the apparent Young's model given by: E

3 2k1

4:4

It should be recognised that although both the elastic and viscoelastic models benefit from being simple, they neglect geometrical factors, such as finite cell dimensions, evolution of cell-micropipette contact region and curvature of the micropipette edges. Thus other models incorporating these geometric factors into a computational form have been developed, which will also account for the inhomogeneities in the cellular properties (Haider and Guilak, 2000). Extensive studies by Guilak's group have used pipette aspiration to investigate the biomechanics of chondrocyte isolated from both healthy and diseased human articular cartilage (Guilak et al., 1999; Jones et al., 1999; Trickey et al., 2004). These studies have quantified both the `pseudo' elastic properties and the viscoelastic time-dependent behaviour. The results of these and other micropipette aspiration studies suggest that chondrocytes have a Young's modulus of approximately 0.5 kPa, and instantaneous and relaxation moduli of 0.4 and 0.2 kPa respectively, as summarised in Table 4.1. Some studies suggest that cells from osteoarthritic (OA) cartilage are stiffer than those from normal tissue, particularly with respect to the equilibrium moduli (Trickey et al., 2000, 2004). In addition, this group has used the same experimental approach to investigate the biomechanics of the chondrocyte nucleus (Guilak et al., 2000; Vaziri and Mofrad, 2007). Indeed micropipette aspiration provides a valuable tool for examining intracellular biomechanical behaviour including nucleus deformation, cytoplasmic biomechanics (Bomzon et al., 2006; Ohashi et al., 2006) and cytoskeletal deformation and remodelling (Fig. 4.1b). Finally, pipette aspiration has also been used to quantify the biomechanical properties of the pericellular matrix associated with isolated chondrons. These studies estimate the Young's moduli of enzymatically isolated chondrons at approximately 25 kPa (Alexopoulos et al., 2005; Guilak et al., 2005). Such studies provide essential data for developing hierarchical models of articular cartilage biomechanics. However, it has to be appreciated that the isolation process may well damage the inherent biomechanics of the chondron, which may be substantially stiffer than the estimated values from these studies (Knight et al., 2001).

4.2.3

Atomic force microscopy (AFM)

Atomic force microscopy (AFM) systems, which were developed from a simple cell poking approach, are now available in both laboratory-based and commercial systems (e.g. Veeco Instruments). Their use in cell biomechanics involves the indentation of the cell surface with a small probe, whose movement

Table 4.1 Summary of compressive stiffness values for chondrocytes, as estimated from a range of different measurement techniques Reference

Modulus

Technique/model

Cell type

Micropipette aspiration Jones et al. (1999)

E 0:65 kPa

Elastic half space

Trickey et al. (2000, 2004), Guilak et al. (2002) Bader et al. (2002) Ohashi et al. (2006)

El 0:41 kPa Er 0:24 kPa E 0:81 kPa E 0:97 kPa

Viscoelastic half space

Human chondrocytes, knee, hip, ankle, elbow, normal and OA Human chondrocytes, knee, hip, normal and OA Adult bovine chondrocytes, MCPJ Adult bovine chondrocytes, MCPJ

K 0:02 N/m El 0:29±0.55 kPa Er 0:17±0.31 kPa

No model used Viscoelastic, isotropic surface

Adult bovine chondrocytes, MCPJ Porcine femoral condyle, superficial and deep zone cells

E 1:10 kPa El 8:00 kPa Er 1:09 kPa

Elastic half space Viscoelastic half space

Adult bovine chondrocytes, distal surface of first metatarsal

E 2:55 kPa El 2:47 kPa Er 1:48 kPa Ea 1:48 kPa El 1:06 kPa Er 0:78 kPa

Linear elastic model Viscoelastic model

Adult bovine chondrocytes, distal surface of first metatarsal

Compression in 3D scaffolds Freeman et al. (1994)

E 4 kPa

Bader et al. (2002) Knight et al. (2002)

Er 2:7 kPa E 3:2 kPa

Compression in agarose and elastic FEM Relaxation in 1% agarose Relaxation in 2% alginate

Atomic force microscopy Bader et al. (2002) Darling et al. (2006) Cytoindentation Koay et al. (2003) Cytocompression Leipzig and Athanasiou (2005)

Shieh and Athanasiou (2006)

Elastic half space Elastic half space

Linear biphasic model Viscoelastic half space

Adult bovine chondrocytes, distal surface of first metatarsal Swarm rat chondrosarcoma cells Adult bovine chondrocytes, MCPJ Adult bovine chondrocytes, MCPJ

E, Young's modulus; Er, relaxation (equilibrium)modulus; El, instantaneous modulus; Ea, aggregate modulus; FEM, finite element model; MCPJ, metacarpal phalangeal joint (proximal surface); OA, osteoarthritic chondrocytes.


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is controlled at a constant velocity (Radmacher et al., 1996; Sato et al., 2000). The tip is typically pyramidal or hemispherical in shape and is carefully moved towards the surface of an individual chondrocyte, imaged through a conventional light microscope. The tip can probe various locations on the cell surface with the force indirectly recorded under indentation control. Hence structural properties, in the form of the force±deformation relationship, can be obtained. Previous studies employing AFM to quantify chondrocyte biomechanics have demonstrated a typical force±displacement relationship with a characteristic toe-in followed by a more linear region with increasing indentation (Bader et al., 2002; Darling et al., 2006). The resulting stiffness of articular chondrocytes has been estimated from the gradient of the linear region with values ranging from 0.02 to 0.10 N/m (Bader et al., 2002), with cells from OA cartilage being significantly softer than those from normal tissue (Hsieh et al., 2008). However, interpretation of the results from AFM deformation is complicated by the tapered shape of its probe tip and its small size relative to the depth of indentation. Therefore to determine biomechanical material properties, such as the cell modulus from this experimental approach, finite element models have been developed (Costa and Yin, 1999). Using a theoretical solution for stress relaxation of a viscoelastic, incompressible, isotropic surface indented with a hard spherical indenter, previous AFM studies have estimated the chondrocyte modulus, as summarised in Table 4.1. In particular, the instantaneous modulus of isolated superficial zone cells has been estimated at a 0.55 kPa whilst the value for middle/deep zone cells is significantly lower at 0.23 kPa (Darling et al., 2006). It should be noted that these values are in broad agreement with those obtained using pipette aspiration.

4.2.4

Cytoindentation and cytocompression

A few studies, notably those from Athanasiou and co-workers, have developed specialist single cell cytoindentation (Koay et al., 2003; Shin and Athanasiou,1999) or cytocompression rigs (Leipzig and Athanasiou, 2005; Shieh and Athanasiou, 2002, 2006). Both approaches have been used to determine viscoelastic creep properties of individual isolated chondrocytes. However, as with micropipette aspiration and AFM, these techniques require assumptions to be made so that theoretical models may be used to derive the cellular biomechanical properties. The cytoindentation tests yield values for instantaneous and relaxation moduli of articular chondrocytes at 8.00 and 1.09 kPa respectively (Table 4.1) (Koay et al., 2003). These values are significantly greater than those obtained using micropipette aspiration or AFM. Similar elevated values for instantaneous and relaxation moduli are reported from cytocompression tests with superficial cells appearing stiffer than those isolated from the middle and deep zones (Shieh and Athanasiou, 2006). The results were broadly similar when derived using three different continuum models (Leipzig and Athanasiou, 2005) (Table 4.1).

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4.2.5


Compression of isolated chondrocytes in 3D scaffolds

One disadvantage with the techniques described above is that the nature of the applied localised cell deformation is distinctly different from that experienced in vivo during physiological loading of articular cartilage. However, to derive cellular biomechanical parameters from experiments with cartilage explants is also problematic, as discussed above. Therefore an alternative approach for examining cellular biomechanics involves compression of living chondrocytes in homogeneous 3D scaffolds. This can yield biomechanical data at both cellular and subcellular levels, associated with a more physiological gross chondrocyte deformation. Numerous studies have used the well-characterised chondrocyte-agarose model to investigate the role of cell deformation. This in vitro model system consists of isolated chondrocytes seeded within agarose gel, typically prepared at a concentration of 2±4% (w/v) (Lee and Bader, 1995, 1997). The cells adopt a spherical morphology with a cortical arrangement of the actin cytoskeletal similar to that observed in situ. Thus the isolated cells maintain their chondrocytic phenotype as shown by the synthesis of type II collagen and the proteoglycan, aggrecan. Compressive strain can be applied to cells seeded within agarose or other low modulus scaffolds, such as alginate, using relatively simple microscope-mounted loading rigs such as that shown in Fig. 4.2(a). These devices enable simultaneous visualisation of cells at different levels of applied gross compression. Studies using confocal microscopy to measure the deformation of viable chondrocytes in agarose demonstrate that gross compression results in deformation to an oblate ellipsoid morphology with significant lateral expansion and conservation of cell volume (Fig. 4.2b) (Lee et al., 2000b). Although this mode of loading is more physiological than that associated with micropipette aspiration, AFM or cytoindentation, the deformation behaviour differs from that in situ within cartilage explants where compression occurs with a reduction in cell volume (Section 4.2.1) (Guilak, 1994; Guilak et al., 1995). The viscoelastic properties of the scaffold relative to the cell can lead to temporal changes in cell deformation during either static or cyclic compression (Knight et al., 1998a). This phenomenon has been exploited to investigate cellular biomechanics by monitoring the reduction in cell deformation over a 60 minute period of static compression in 2% (w/v) alginate gel (GMB low viscosity, Kelco, UK) (Knight et al., 2002). Cell strain measurements were plotted against the corresponding viscoelastic stress relaxation in the gel, measured using a 2.5 N load cell. It was therefore possible to estimate the cell compressive modulus (E stress/strain) at a value of approximately 3.2 kPa (Table 4.1). A similar approach has also been used for deriving cell moduli from measurements of cell deformation in compressed agarose gels (Table 4.1) (Bader et al., 2002; Freeman et al., 1994). However, these approaches do not take into account the long-term viscoelastic behaviour of the cell, as opposed to


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4.2 Compression of chondrocytes in 3D scaffolds. (a) Schematic cross-section of loading rig for compression of a cartilage explants or a chondrocyte seeded scaffold. The device mounts upon the stage of an inverted microscope and allows simultaneous visualisation of cells during compression thereby enabling analysis of cellular and intracellular biomechanics (Knight et al., 1998b). The specimen is placed on a coverslip and hydrated in medium. Compression is applied via one or two sliding platens connected to stepping linear actuators controlled via a PC. (b) Representative confocal images of a single isolated chondrocyte visualised in an agarose construct at 0, 5, 10, 15 and 20% gross compression strain. The top row shows a cell labelled with calcein AM whilst the second row shows a separate cell labelled with mitotracker green and syto16 to label the mitochondria and nucleus respectively. Images demonstrate the deformation of the cell and intracellular structures during gross compression. Scale bar indicates 5 m. (c) The associated intracellular local strains distribution parallel to the axis of compression was calculated from the mitochondria images using digital image correlation. The magnitude and direction of the local strains are shown on a pseudocolour scale relative to the uncompressed images (Knight et al., 2006).

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that of the scaffold, as demonstrated in previous cell biomechanics studies (Darling et al., 2006; Trickey et al., 2004). Intracellular biomechanics may also be examined based on analysis of cell deformation in 3D scaffolds. Indeed, previous studies have investigated the biomechanics of the nucleus and mitochondria based on compression of chondrocytes in agarose gel (Bomzon et al., 2006; Knight et al., 2006; Lee et al., 2000b). However, with increasing time in culture, the matrix synthesised by the isolated chondrocytes forms a pericellular shell which is stiffer than the surrounding agarose and thus prevents cell deformation during gross compression, thereby providing indirect analysis of pericellular matrix biomechanics (Knight et al., 1998b, 2001).

4.3

Intracellular biomechanics

Whilst the measurement of cell deformation is essential for analysing gross chondrocyte biomechanics, an understanding of intracellular biomechanics provides additional, important information. In particular, it can generate a clearer understanding of both the structures which provide cells with their viscoelastic time-dependent biomechanical properties and the mechanotransduction signalling pathways through which mechanical loading is translated into an alteration in cell activity. Studies have reported that mechanical loading of cartilage explants induces cell deformation with associated distortion of cellular organelles including the rough endoplasmic reticulum (Szafranski et al., 2004), mitochondria (Knight et al., 2006), nucleus (Guilak, 1995) and potentially the primary cilium (Jensen et al., 2004), all of which may have a role in mechanotransduction. However, of all the intracellular structures, mechanical deformation of the nucleus has been most commonly examined since it may be involved in mechanotransduction through changes in gene expression and nuclear transport (Buschmann et al., 1996). Studies using pipette aspiration have estimated the viscoelastic biomechanical properties of isolated chondrocyte nuclei, with moduli values approximately 5±10 times stiffer than the surrounding cytoplasm (Guilak et al., 2000; Vaziri and Mofrad, 2007). Thus, the levels of nucleus deformation are typically less than that of the cell (Fig. 4.2b) (Guilak, 1995; Knight et al., 2002; Lee et al., 2000b). However, changes in the biomechanical properties of the nucleus may occur during differentiation or in the diseased state, with associated changes in mechanotransduction. In addition, the relative stiffness of the nucleus means that where cell deformation is sufficient to induce nuclear distortion, the nucleus is likely to provide a significant contribution to the gross biomechanical stiffness of the chondrocyte. Nucleus morphology and deformation are typically heterogeneous and nonuniform in nature (Knight et al., 2002). This needs to be considered when quantifying nucleus deformation ideally performed using live cell imaging. Cytoskeletal integrity is important for strain transfer to the nucleus (Djabali,


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1999; Janmey, 1998) and thus changes in cytoskeletal organisation will influence nucleus deformation and other aspects of intracellular biomechanics (Knight et al., 2002; Lee et al., 2000b). For the study of intracellular deformations, the optical sectioning capability of confocal laser scanning microscopy enables clear, blur-free imaging of intracellular structures. The scope of this technology has been greatly enhanced by the ever-increasing range of fluorescent compounds for labelling intracellular structures and organelles in living cells (see Molecular Probes: http:// probes.invitrogen.com/). Thus viable fluorescent markers are now available for organelles including the nucleus, mitochondria and endoplasmic reticulum. Whilst most of these compounds are cell permeable, labelling of other structures, such as the cytoskeletal protein networks, is possible using microinjection of fluorescent analogues (for review see Goldman and Spector, 2004). Alternatively the development of transfection techniques involving fluorescent tags, such as GFP, provides a powerful tool for visualising intracellular structural dynamics and biomechanics within living chondrocytes (Fig. 4.1b). The complexity and heterogeneity of intracellular structures, such as the cytoskeleton, frequently require advanced computational techniques, such as digital image correlation (DIC), to quantify the local biomechanics. DIC can analyse, with sub-pixel resolution, the movement and distortion within pairs of images and has been used for measuring deformation in a wide range of structures including articular cartilage (Chahine et al., 2004; Wang et al., 2003). More recently, the technique has been optimised for measuring the deformation and biomechanics of fluorescently labelled intracellular structures, visualised in living chondrocytes using confocal microscopy (Delhaas et al., 2002; Helmke et al., 2000, 2001, 2003; Hu et al., 2003). Intracellular displacements and strains can be automatically computed and graphically displayed in the form of pseudocolour maps (Fig. 4.2c). The technique is able to determine the local compressive and tensile strains as well as shear strains, area strains, von-Mises strains and the magnitude and direction of the principal strains. It should be noted that this approach does not distinguish between percentage changes in dimensions resulting from true biomechanical deformation, or strain, and inherent temporal movement. However, with appropriate experimental controls it is possible to use this information to generate computational models to describe biomechanical properties of intracellular elements (Bomzon et al., 2006).

4.4

Biomechanical conditioning of chondrocytes

4.4.1

Introduction

In addition to the many studies focused on chondrocyte biomechanics, others have proposed the use of in vitro biomechanical conditioning strategies for chondrocyte-seeded scaffolds as an essential feature for the long-term

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functionality of tissue engineered implants for cartilage repair (e.g. Guilak et al., 2001). This requires the development of suitable bioreactors, incorporating mechanical loading modules for use in a controlled biological environment. These can provide appropriate mechanical conditioning regimes to stimulate the formation of a functional neo-cartilage, thereby improving the efficacy and efficiency of production of tissue engineered implants for cartilage defects. Two fundamental approaches have been adopted to examine the response of chondrocytes to biomechanical stimuli in vitro, similar to those described earlier in the chapter. One involves the use of cartilage explants in which the chondrocytes are associated with ECM mimicking the in situ case (Gray et al., 1989; Jin et al., 2001; Sah et al., 1989). The alternative involves model systems incorporating isolated chondrocytes, which may be maintained in various culture systems, including pellet cultures, suspension cultures or monolayer cultures of either high or low cell densities, or the cells may be seeded within a 3D construct, typically comprising a hydrogel or a porous scaffold (Buschmann et al., 1995; Freeman et al., 1994; Kisiday et al., 2002; Lee and Bader, 1997). Both model systems have their proponents. It is evident that cartilage explants are appropriate for studies investigating fundamental mechanotransduction events associated with normal turnover and pathology. However, they are less suitable to examine individual extracellular components of mechanotransduction, such as cell deformation, due to the inherent coupling of mechanical and physicochemical processes in the charged ECM. Alternatively model systems, although non-physiological in nature, are generally considered appropriate for tissue engineered therapeutic strategies. Accordingly, a wide variety of model systems have been proposed in the literature (Table 4.2). One of the most popular systems involves chondrocytes seeded in 3D agarose constructs, as discussed earlier in the chapter. The system maintains a rounded chondrocyte phenotype over extended culture periods (Aydelotte et al., 1990; Benya and Shaffer, 1982; Hauselmann et al., 1994), by preventing the formation of actin stress fibres, which are evident when chondrocytes are cultured in monolayer. Thus agarose or similar model systems such as alginate, which can be characterized in mechanical terms (Knight et al., 1998a,b; Lee and Bader 1995), provide an ideal construct for the application of gross compression. Although compression of cell±agarose constructs will initiate transient changes in hydrostatic pressure and fluid flow, which have both been implicated in mechanotransduction, it is widely believed that cell deformation is the primary mediator. The effects of loading on cell morphology and deformation during the application of physiological levels of compressive strain has been discussed (Fig. 4.2b). Various factors have been shown to influence the level of cell deformation, including the presence and mechanical properties of the elaborated PCM, the modulus of the scaffold relative to that of the cells and matrix and the loading regime and viscoelastic properties of the scaffold (Bader et al., 2002; Knight et al., 1998a, 2002).

Table 4.2 Summary of experimental protocols used to assess in vitro biomechanical conditioning of chondrocytes in model systems Source

Cell source

Sample/scaffold material

Loading conditions/Culture conditions

Lee and Bader (1997), Lee et al. (1998a,b) Chowdhury et al. (2006)

Bovine (adult)

Agarose

15%, 0.3, 1 and 3 Hz, 48 h

Human

Agarose

Bonassar et al. (2001) Kisiday et al. (2002)

Bovine (calf) Bovine (calf)

Cartilage discs Agarose and self-assembling peptide gel

Buschmann et al. (1995) Buschmann et al. (1999)


Agarose Explants

Sah et al. (1989), Li et al. (2001) Mauck et al. (2000)

Bovine (calf)

Explants

Bovine (calf)

Agarose

Millward-Sadler et al. (1999) Guilak et al. (1994), Fermor et al. (2001) Altman et al. (2002)

Human normal or OA ± passaged Bovine (calf)/porcine

Monolayer

15%, 1 Hz, 48 h 10 ng mlÿ1 TGF 2%, 0.1 Hz, 48 h 2.5±3%, 1 Hz, 1 h on/1 h off, days 9±12 or 1 h on/7 h off, days 27±34 0.001 Hz to 1 Hz, 10 h 0.001 Hz, 23 h, days 4 and 6 0.01 Hz, 23 h, days 3 and 5 2.4%, 0.01 Hz (200 kPa) 2 h on/2 h off, 23 h, 0.88±1 mm compression 10%, 1 Hz, 3 1 h intermittent/day, 5 days/week for 4 weeks 0.33 Hz, 20 min

Explants

0.1, 0.5 MPa, 0.5 Hz, 24 h

Human bone marrow stromal cells ± passaged Bovine (calf)

Silk fibre matrices

0.0167 Hz, 2 mm compression

Martin et al. (2000) Hunter et al. (2002) Hunter and Levenston (2002a) Waldman et al. (2003, 2007)


Xie et al. (2007)

Young rabbit

Bovine

Polyglycolic acid

Compressive deformation for samples 1.3±3 mm thickness Collagen I Oscillatory 25 4%, 1 Hz, 24 h Either core explant with Dynamic compression at 10 4% at 0.3 or 1.0 Hz, agarose core or agarose alone after 3 or 15 days of pre-culture Porous calcium Static culture for 4 weeks ± various combinations polyphosphate of compression and/or shear strain at 1 Hz for 6 min, followed by 48 h recovery Microporous PLL±PCL Dynamic compression at 10% (continuous and elastomeric scaffolds intermittent) at 0.01±0.5 Hz

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4.4.2


Biomechanical loading systems

Conventional in vitro mechanical tests require the soft tissues to be kept in a moist environment and are performed on either a Universal materials test facility or a specially designed test system. However, if tissue explants or chondrocyteseeded 3D constructs are to be examined, viability must be maintained in an environment similar to a conventional CO2 incubator, whilst the system is subjected to either static or dynamic loading. With regards to commercial systems, that produced by Flexercell has gained widespread use for subjecting specimens to a range of tensile loading regimens. Although there is a compressive system equivalent, as reported by Graff and colleagues (2000), most researchers have used custom-made bioreactor systems to apply compression to chondrocyte cultures. Others describe bioreactors which incorporate shear, fluidinduced shear, hydrostatic pressure or a combination of loading modalities (Wernike et al., 2008). One of the few compression-type bioreactors available commercially (Zwick Testing Machines Ltd, Leominster, UK) has been used to apply static and dynamic loading to biomaterial constructs seeded with chondrocytes. The system, detailed over a decade ago (Lee and Bader, 1997), consists of a conventional loading frame with an hydraulic actuator-controlled vertical assembly, which enters a tissue culture incubator (Heraeus Instruments, Brentwood, UK). The assembly is connected to a central rod which is attached to a mounting plate located within a Perspex box, as shown in Fig. 4.3. The box, in turn, is placed on a circular platten fixed to the base of the loading frame. The mounting plate holds 24 loading pins, half of which are unconstrained to move vertically in harmony with the loading assembly. Each loading pin incorporates an 11 mm circular Perspex indenter, which applies compressive strain to samples located within separate wells of a 24-well tissue culture plate. An important aspect of any prolonged culturing period is the maintenance of cell viability. During a 48-hour culture period in the compressive cell strain system (Fig. 4.3), the chondrocyte viability has been shown to remain above 95% within both unstrained and strained constructs (Lee and Bader, 1997). Thus differences in metabolism could not be attributed to alterations in chondrocyte viability. Accordingly, this loading system has been used by the authors to examine key variables associated with the application of mechanical conditioning to chondrocyte-seeded 3D constructs. Specific studies have aimed at elucidating the influence of mechanical loading regimes, cell sub-populations and scaffold materials on the efficacy of mechanical conditioning strategies.

4.4.3

Mechanical loading regimens

An initial series of experiments on full-depth adult bovine chondrocytes seeded in agarose investigated the influence of both static and dynamic continuous


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4.3 Photograph (a) and schematic representation (b) of the compressive cell strain system.

compression at a strain amplitude of 15%. The normalised data for tritiated ([3H]) thymidine incorporation, a measure of cell proliferation, are presented in Fig. 4.4(b). It is evident that constructs subjected to dynamic strain at all three frequencies exhibited significant stimulation in comparison with unstrained control values, although there were no statistically significant differences between the dynamic frequencies. The corresponding data for glycosaminoglycan (GAG) synthesis values, normalised to unstrained control levels, are presented in Fig. 4.4(a). It can be seen that at the low frequency (0.3 Hz) dynamic strain inhibited GAG synthesis, while a frequency of 1 Hz induced a significant stimulation of GAG synthesis. At the higher frequency of 3 Hz GAG synthesis returned to unloaded control levels. These findings suggested that, using the chondrocyte±agarose system at the strain level of 15%, there is a frequency range which can induce GAG synthesis, with a lower cut-off frequency between 0.3 and 1 Hz and an upper cut-off between 1 and 3 Hz. This frequency will be dependent on the strain level, the mechanical properties of the matrix and the size and shape of the individual specimens. Indeed it has been established that increasing frequency will increase the rate of fluid flow in the periphery of the specimens, but will reduce the effective width of the peripheral ring in which it occurs (Sah et al., 1989). It is conceivable, therefore, that the influence of the central core may mask stimulatory effects within the peripheral ring at the higher frequency of 3 Hz. Although all loading regimens yielded an inhibition in protein synthesis, the analysis of data revealed an association between the frequency rate and the level of inhibition (data not presented, Lee and Bader, 1997). The findings also implied that each metabolic parameter may be influenced by dynamic strain regimens in a distinct manner, implying that the associated signalling mechanisms are uncoupled (Lee and Bader, 1997). A subsequent study examined the metabolic response of adult bovine cells from different zones of cartilage under dynamic compression (Lee et al., 1998a). Thus slices of cartilage from the uppermost 15±20% of the total uncalcified tissue

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4.4 GAG synthesis and [3H]-thymidine incorporation by full-depth bovine chondrocytes (a and b, respectively, and surface and deep bovine chondrocytes (c and d) embedded in agarose constructs and subjected to 15% dynamic compressive strain amplitude at various frequencies for 48 h. The values are presented as % change from unstrained control levels. Each value represents the mean and standard error of at least 12 replicates from at least two separate experiments. Unpaired Student's t-test results indicate differences from control values as follows: * p 0:05.

depth were removed from the proximal joint surface, and the `superficial cells' isolated and seeded into agarose constructs. `Deep cells' from the residual tissue were seeded into separate constructs. Both groups of constructs were cultured under a continuous compressive amplitude of 15% for 48 hours. Normalised GAG synthesis data, as presented in Fig. 4.4(c), revelaled that all three frequencies produced an inhibition in GAG synthesis by superficial cells, the differences being statistically significant at 0.3 and 3 Hz. With reference to the GAG synthesis by deep cells, 0.3 Hz produced a significant reduction, and 1 Hz induced a highly significant stimulation of 50% (Fig. 4.4c). By contrast, all three dynamic frequencies induced a significant increase in [3H]-thymidine incorporation by superficial cells (Fig. 4.4d). There was no statistically significant difference in the level of stimulation between the three dynamic strain regimens. However, [3H]thymidine incorporation by deep cells was not greatly influenced by the application of compressive strain. The only increase corresponded with the 0.3 Hz dynamic regime when compared with unstrained controls (Fig. 4.4d).


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Data from this study demonstrated that the control of GAG synthesis and proliferation in response to dynamic compression are not merely uncoupled, but occur in different sub-populations of chondrocytes within the full-depth cell isolate (Lee et al., 1998a). This conclusion raises the possibility of the involvement of distinct intracellular mechanotransduction mediators. One possible mediator is nitric oxide, which is known to influence both GAG synthesis and proliferation in chondrocytes, and may be modulated by physical stimuli. This has been examined by the authors in a series of papers (Chowdhury et al., 2001, 2003a, 2006; Lee et al., 1998b, 2000a), which have all indicated a downregulation of both nitric oxide and associated PGE2 by dynamic compression. Additionally in the presence of a pro-inflammatory cytokine, it has been shown that dynamic compression counteracts the IL1- -induced release of nitric oxide and PGE2 by superficial zone chondrocytes cultured in agarose constructs (Chowdhury et al., 2003a). This has important implications in the potential of exercise regimens, involving controlled biomechanical loading in vivo, for those subjects with inflammatory joint disease. It has been previously shown with different cell types that dynamic loading for short periods can stimulate cellular activity (Fermor et al., 2001; Robling et al., 2002; Rubin and Lanyon, 1992). This prompted an examination of the temporal response of full-depth bovine chondrocytes seeded in agarose constructs to both intermittent and continuous compression regimens (Chowdhury et al., 2003b). Intermittent compression was applied for 1.5 (denoted by I 1.5), 3 (I 3), 6 (I 6) and 12 (I 12) hours at 15% strain at 1 Hz with equivalent unloaded periods for a total of 48 hours. Each of the intermittent loading regimens, involving 86 400 duty cycles, resulted in significant increases in both sulphate and thymidine incorporation (Fig. 4.5). However there were clear differences in the optimal profiles over the 48-hour culture period. For example, there was a monotonic increase in stimulation of sulphate incorporation with increasing strain durations, such that differences were found to be statistically significant between the values at 1.5 h compared to those at 6 and 12 h (p < 0:01). By contrast, thymidine was maximal after 1.5 h of intermittent loading with values of 197% compared with unstrained controls. Longer bursts of cyclic compression were associated with a decrease in the absolute proliferative response (Fig. 4.5b). Similar findings were observed with extended periods of cyclic compression (Chowdhury et al., 2003b). It was postulated that dynamic compression acts as a competence and/or progression factor for DNA synthesis in chondrocytes. Once the cell has been stimulated to enter the cell cycle it will take several days to progress through the complete cycle. Thus stimulation of cell proliferation within the timescale of this study is a unique event, as opposed to repeated stimulation, which is required to up-regulate proteoglycan synthesis. It is proposed that the frequency of dynamic strain does not affect the response, as it appears from the continuous compression data that a finite number of cycles of dynamic strain are sufficient for stimulation. This suggests that the cells are temporally processing the mechanical stimulus.

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4.5 The percentage change from unstrained control values for 35 SO 4 incorporation (a) and [3H]-thymidine incorporation (b), by chondrocytes seeded in 3% agarose and subjected to different periods of intermittent compression of 15% dynamic strain at a frequency of 1 Hz. Error bars represent the mean and SEM of 24 replicates from two separate experiments (based on Chowdhury et al., 2003b).

A similar approach was adopted in a recent study, which examined the effects of both intermittent and continuous compression on young rabbit chondrocytes seeded in microporous elastomeric scaffolds made from a poly--caprolactone± poly-L-lysine (PLL±PCL) co-polymer cultured for up to 6 days (Xie et al., 2007). Both ECM and appropriate genes were monitored. Data indicated a mechanical-induced up-regulation in both GAG and collagen secretion up to a maximum at day 3. The mRNA expression of collagen type II was noted to be up-regulated with intermittent stimulation in the short term although, over an extended time period, there was a redundancy of stimulation leading to a downregulation of cell biosynthesis. The benefits of intermittent compression were also demonstrated for bovine chondrocytes grown on porous ceramic substrates (Waldman et al., 2003). After a free swelling culture period of 4 weeks, the cultures were subjected to short bursts (400 duty cycles at 1 Hz) of either compression or shear loading. Biochemical analysis indicated an up-regulation in both proteoglycan and collagen content, which was more significant under dynamic shear stimulation at 2%.


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This work was extended to examine the intermittent effects of combinations of compression and shear stimulation (Waldman et al., 2007). The findings again revealed an increase in up-regulation of ECM components with stimulation, which correlated with an increase in the mechanical integrity of the cultured constructs. A recent study (Wernike et al., 2008) examined the influence of dynamic mechanical stimulation at a different oxygen tensions. They used a custom-built joint stimulator, which permitted the application of both axial compression and simulated surface torsion. Calf chondrocytes were seeded in porous polyurethane scaffolds and subjected to 1 hour stimulation per day for up to 34 days in either a normoxic or anoxic (5% O2) environment. There were clear differences in results of both gene and protein analyses in the two conditions. For example, under reduced oxygen tension, there was an increase in mRNA levels of type II collagen and aggrecan, with an associated increase in GAG/ DNA content. Mechanical compression yielded an enhanced level of Type II collagen gene. Interestingly a combination of biomechanical stimulation and hypoxia produced a significant down-regulation of type I collagen gene expression, suggesting that their combination could prove an effective tool for maintaining chondrocyte phenotype in culture.

4.4.4

Cell source

Autologous chondrocyte implantation (ACI or ACT) represents a wellestablished approach to repair full or partial thickness cartilage defects (Brittberg et al., 1994). The technique is detailed in Chapter 9. To review briefly, chondrocytes are isolated from a small biopsy, removed from a low load-bearing site at the periphery of the joint surface. The chondrocytes are then expanded, typically ten-fold, in monolayer culture for approximately 4 weeks to ensure the practicality and financial viability of the technique (Brittberg et al., 1994). While these methods are extremely effective at inducing cell proliferation, chondrocytes are known to dedifferentiate and adopt a fibroblastic phenotype in monolayer culture and this process is only slowly reversible (Benya and Shaffer, 1982; deHaart et al., 1999; Mayne et al., 1976). Additionally, the induction of cell proliferation during expansion will increase the telomeric age of the cells, which, in turn, may influence their response to biomechanical stimulation. Accordingly, a number of cell-related factors need to be addressed, particularly given the diverse nature of the experimental protocols employed in previous studies (Table 4.2), which ultimately can provide only phenomenological information. Nonetheless, a number of studies by the authors and others have attempted to address these issues in a systematic fashion. One study examined the effects of continuous dynamic compression to agarose constructs containing full-depth bovine chondrocytes isolated from either the femoral condyle and the patella groove of the equine knee joint

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(Wiseman et al., 2003). These tissue locations were selected to represent areas experiencing different loading conditions, as identified during normal articulation of the joint (Athanasiou et al., 1991) and thus correspond to the sites of lesion and donor tissue, respectively, in the ACI protocol (Brittberg et al., 1994). Results indicated differences between the response from the two selected tissue locations, in terms of both the absolute levels of proteoglycan synthesis and cell proliferation and also changes induced by dynamic compression. Additionally, nitric oxide was inhibited by the application of loading for cells isolated from both anatomical regions for all equine samples tested. However the heterogeneous response may have been compounded by the use of tissue of different ages and unknown exercise history, a factor which is highly relevant for the translation of tissue engineered cartilage repair systems from the laboratory to the clinic. The influence of passage in monolayer on the response of full-depth bovine chondrocytes to the application of dynamic compression was examined (Wiseman et al., 2004). The chondrocytes were either seeded directly into 3D agarose constructs or were passaged up to four times at weekly intervals prior to seeding into the constructs. It is well established that cells at passage 2 and beyond express a fibroblastic phenotype even when cultured in alginate beads, as evidenced by the presence of type I collagen and the absence of type II collagen staining (Wiseman et al., 2004). On application of dynamic compression (15% strain at 1 Hz), chondrocytes at early passages (P1 and P2) exhibited a stimulation of both proteoglycan synthesis and cell proliferation, as illustrated in Fig. 4.6(a) and (b), respectively. However, beyond passage 2 (P3± P4) the reverse was evident with dynamic compression acting to inhibit these factors, which are essential for the re-population of defect sites and the formation of a cartilaginous neo-tissue. Thus the study demonstrated a clear relationship between passage number and mechanical sensitivity, with mechanicalinduced stimulation only evident during early passage. Many studies have demonstrated that the provision of specified differentiation factors, such as fibroblast growth factor (FGF) and transforming growth factor beta (TGF- ), either during expansion or subsequent culture in 3-D, can influence the re-expression of the chondrogenic phenotype (deHaart et al. 1999; Jakob et al., 2001; Lemare et al., 1998; Yaeger et al., 1997). This was further investigated in a study by Chowdhury et al. (2004) utilising monolayerexpanded human chondrocytes surplus to requirement for clinical ACI repair procedure (expanded and supplied by Verigen AG, Leverkusen, Germany). The cells were seeded into agarose constructs and subjected to dynamic compression during incubation in either standard medium comprising Dulbecco's modified eagle's medium (DMEM) + 20% fetal calf serum (FCS), or a defined chondrogenic medium comprising DMEM + insulin plus transferring and selenous acid (ITS) + 10 ng mLÿ1 TGF- . Absolute levels of both proteoglycan synthesis and cell proliferation were elevated during incubation in the presence of the con-


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4.6 Sulphate incorporation (a) and [3H]-thymidine incorporation (b) by freshly isolated bovine chondrocytes (designated passage 0) and chondrocyte passaged in monolayer between one and four times. The cells were subsequently embedded in agarose constructs and subjected to 15% dynamic compressive strain amplitude at 1 Hz for 24 hours. The values are presented as % change from unstrained control levels. Each value represents the mean and standard error of at least 12 replicates. Unpaired Student's t-test results indicate differences from control values as follows: * p < 0:05 (based on Wiseman et al., 2004).

ditioned medium. In addition, a further up-regulation was achieved by application of dynamic compression (Fig. 4.7). Importantly, during incubation in DMEM + 20% FCS alone, dynamic compression failed to stimulate either metabolic parameter, similar to the response demonstrated for monolayerexpanded bovine chondrocytes reported above P2 (Fig. 4.6). These data provide further evidence for a link between the expression of a chondrocytic phenotype

4.7 Sulphate incorporation (a) and [3H]-thymidine incorporation (b) by human monolayer-expanded chondrocytes embedded in agarose constructs and subjected to 15% dynamic compressive strain amplitude at 1 Hz for 48 hours (black) or remained unstrained (white). The constructs were maintained in DMEM + 20% FCS or a defined medium comprising DMEM + ITS + 10 ng mLÿ1 TGF- . Each value represents the mean and standard error of at least 12 replicates. Unpaired Student's t-test results indicate differences from control values as follows: * p < 0:05 (based on Chowdhury et al., 2004).

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and a beneficial response to dynamic compression. In addition this study demonstrates the importance of interactions between biophysical and biochemical stimuli, an understanding of which is essential for the successful utilisation of biomechanical conditioning strategies.

4.4.5

Scaffold materials

As indicated in Table 4.2, there are a wide range of scaffold materials which have been employed to examine the metabolic effects of chondrocytes when subjected to dynamic loading. It might be supposed that the cell±scaffold interactions in agarose gels are substantially different from those in protein scaffolds (Hunter and Levenston, 2002b). In the former the cells preferentially bind to the synthesised PCM as it is deposited, while in protein scaffolds, cell adhesion molecules such as integrins, will enable direct interactions between the cell and the scaffold. Cell receptor binding is well known to alter both mechanical behaviour and the manner in which cells respond to mechanical conditioning. Indeed in the study involving chondrocytes seeded in fibrin glue, results suggested that early sustained oscillatory compression, inhibited both cell proliferation and matrix accumulation (Hunter et al., 2002). These results are in marked contrast to those found in other systems (Lee and Bader, 1997; Sah et al., 1989). An alternative series of scaffolds proposed for cartilage tissue engineering is based on poly(ethylene glycol) (PEG) hydrogels (Bryant and Anseth, 2001), which exhibit a range of mechanical properties, for example compressive stiffness ranging from 60 to 670 kPa, and controlled degradation profiles. Previous work has reported that such photo-crosslinkable matrix maintains chondrocyte viability and promotes deposition of both proteoglycans and type II collagen (Elisseeff et al., 2000). The effects of changes in the hydrogel crosslinking density on the metabolic response of chondrocytes to continuous dynamic compression of 15% at a 1 Hz frequency for 48 hours were investigated (Bryant et al., 2004). Adult bovine chondrocytes were seeded into two PEG dimethacrylate (PEGDM) gels crosslinked with final concentrations of 10% and 20% (w/w). An increase in crosslinking density resulted in an inhibition in cell proliferation and proteoglycan synthesis. The normalised data for both sulphate and thymidene incorporation and nitrite production are presented in Fig. 4.8. Dynamic compression only marginally influenced GAG synthesis in the 10% gel, although for the 20% gel, there was a marked decrease in PG production. Cell proliferation was inhibited in both crosslinked gels but particularly in the highly crosslinked gel. By contrast, nitrite release was slightly increased as a result of dynamic stimulation. These trends in the metabolic response are in marked contrast to those found for 3% agarose constructs seeded with bovine chondrocytes (Fig. 4.8). It is clear that the interactions between cells and scaffold materials must be well characterised before successful strategies for biomechanical conditioning can be adopted.


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4.8 Sulphate incorporation (a) and [3H]-thymidine incorporation (b) by full depth bovine chondrocytes embedded in 3% agarose constructs (grey) or constructs comprising 10% (white) or 20% (black) PEGDM. All constructs were subjected to 15% dynamic compressive strain amplitude at 1 Hz for 48 hours. The values are presented as % change from unstrained control levels. Each value represents the mean and standard error of at least 12 replicates. Unpaired Student's t-test results indicate differences from control values as follows: * p < 0:05.

4.5

Future trends

Chondrocyte biomechanics has developed considerably in recent years in terms of both experimental and computational modelling techniques. With regards to the latter, future work is likely to involve the development of more sophisticated triphasic cell biomechanical models at a variety of hierarchical levels associated with tissues, cells, intracellular structures, such as the nucleus or cytoskeletal networks, and individual proteins, such as stretch activated ion channels. This approach involving different length scales will provide a more realistic model of chondrocyte biomechanics and response to loading, including the associated activation of putative mechanoreceptors. In addition future biomechanical models must also take into account the active biological response to mechanical loading, such as mechanically induced remodelling of the actin cytoskeletal, with resulting changes in chondrocyte biomechanics. Thus it should be possible to develop the equivalent of Wolff's Law for single chondrocytes and to incorporate previous loading history into any model of cellular biomechanics.

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In addition new experimental techniques such as magnetic twist cytometry and scanning acoustic microscopy, may be adapted for cellular and intracellular biomechanics. However, these and existing measurement techniques are designed to examine isolated chondrocytes only. Therefore a major challenge involves the development of minimally invasive experimental techniques that are able to determine the viscoelastic properties of chondrocytes within the native cartilage PCM microenvironment and for cells subject to more appropriate physiological loading. To extend this further the ultimate goal would be to quantify cellular biomechanics in vivo during physiological joint loading, to aid the development of new techniques for diagnosis and treatment of cartilage disease and injury. The biomechanical conditioning studies highlight the complex interplay between stimuli and metabolic parameters, which appear to be distinctive and uncoupled. Nonetheless provided suitable monitoring systems are available, there remains a possibility of fine-tuning the mechanical stimulation to elicit a specific cellular response during the in vitro conditioning period. Future developments will include the use of mini-bioreactor systems that permit the application of defined mechanical loading regimes to constructs maintained in an uninterrupted and defined oxygen environment. These constructs might contain either chondrocytes and/or chondrons and be stimulated for extended time periods. This permits the provision of a more physiological mechano/ metabolic environmental conditions that will be a powerful tool for tissue engineering and for studying pathophysiological processes by providing more relevant 3D model systems that could be used for drug discovery applications. Future studies will interrogate key signalling pathways that are known to be both mechano- and oxygen-sensitive. These include the pathophysiological release of nitric oxide and prostaglandin E2 by chondrocytes (Chowdhury et al., 2006, 2008) and the potential of mesenchymal stem cells to differentiate into the chondrogenic lineage (Campbell et al., 2006; Terraciano et al., 2007). The adoption of a systematic approach can ultimately result in the definition of underlying mechanistic parameters leading to the derivation of predictive strategies, incorporated within computational models, to control the optimisation of neo-cartilage formation.

4.6

Acknowledgements

The authors acknowledge the invaluable assistance of Professor David Lee, Dr Tina Chowdhury, Dr Toshiro Ohashi and other colleagues, who have contributed to much of the experimental work described within this chapter.

4.7

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5


F . D E L L ' A C C I O , Barts and The London, Queen Mary's School of Medicine and Dentistry, UK and T . L . V I N C E N T , Kennedy Institute of Rheumatology, UK

Abstract: It is widely accepted that cartilage injury leads to osteoarthritis (OA), although the mechanisms by which this occurs are still poorly understood. Research in this area has been somewhat neglected in recent years, but it was a highly fashionable academic pursuit in the 18th, 19th and early 20th centuries and many seminal observations were made during this time. These included the findings that acute cartilage injury induces an active chondrocytic response, involving both degradative as well as synthetic processes that resemble OA. There was also evidence of a repair response, determined to be both from the substance of the tissue and from the underlying bone marrow. In patients, injury to cartilage is defined as either direct, e.g. following intra-articular fracture, or indirect by repetitive wear on the tissue, with age or following joint destabilisation. Although both are associated with the risk of developing OA, this risk is variable and there are emerging data to suggest that some focal cartilage lesions not only do not progress, but may actually heal spontaneously. Recent in vitro studies have begun to unravel the molecular basis for these responses, and these are identifying potentially important pathways which may be involved in driving OA, as well as those that stimulate cartilage repair. Key words: joint surface injury, osteoarthritis, explantation, intrinsic repair, impact load.

5.1

Introduction

Very little is known about how tissues respond to injury, apart from the wellestablished activation of the clotting cascade upon vascular damage: this is a highly orchestrated process that is initiated by platelets binding to the damaged vascular endothelium. Activated platelets regulate the catalytic activities that initiate thrombin generation, which is then sustained at the site of injury by the recruitment of circulation monocytes and neutrophils. Monocytes are capable of driving thrombin production through expression of tissue factor, a protein also expressed by subendothelial cells and activated endothelium which, when in contact with factor VIIa activates the clotting cascade zymogens. The combination of damaged endothelium, the activated surface of the platelet and activated leukocytes provide the optimal environment for locally contained control of haemostasis.1

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In skin, wounding evokes a coordinated cellular response which is critical upon resident cells as well as invading immune cells, which are essential for resolution of the healing response.2,3 Clearly, skin wounding also causes endothelial damage, as it is a vascular tissue, but whether there are other triggers that initiate this response, is not known. What triggers injury responses in tissues that are avascular is very poorly understood. Articular cartilage injury is considered to be one of the most important risk factors for osteoarthritis (OA); disease is increased in aged joints (chronic wear and tear-associated injury), and occurs prematurely in individuals who have sustained injury to the cartilage surface or to other associated structures within the joint. Damage to the menisci and cruciate ligaments is thought to lead to increased wear across the articular surfaces through induction of joint instability. The response of articular cartilage to injury was a highly fashionable pursuit in the 18th and 19th centuries as well as the early 20th century, but it is surprising how òut of vogue' this area of research has become. This is perhaps surprising in view of the increasing prevalence of OA in our society associated with increased longevity and the obesity epidemic. As funding bodies focus their attention to neglected prevalent diseases such as OA, so it is likely that research into basic mechanisms of cartilage injury will increase. This chapter will address what is known about the natural history of clinical cartilage injury in vivo, and will discuss the work that has been undertaken to understand tissue responses in animal models of in vivo injury, and in vitro injury systems.

5.2

Clinical in vivo cartilage injury

In vivo injury of cartilage is often referred to as joint surface injury (JSI) and results in joint surface defects (JSD). It includes a broad range of conditions involving damage to the articular cartilage, with or without involvement of the subchondral bone. Such lesions are very common, being reported in over 60% of all arthroscopic procedures.4,5 Chronic JSDs represent a clinical problem because (1) they can be symptomatic and disabling, with pain and/or locking of the joint, and (2) they predispose to further cartilage loss and development of OA.6 Chondral lesions vary greatly in their morphology and topography, and this variation likely influences their outcome and clinical manifestations. Broadly speaking, lesions can be divided into being localised, such as those due to trauma, or diffuse, such as those that characterise OA. A second important division is that between superficial, partial thickness cartilage defects, which do not involve the subchondral bone, and full-thickness lesions which cross the ostoechondral junction (Outerbridge grade 3 or 4). Superficial cartilage defects, particularly if linear, without tissue loss, have a poor repair capacity. Although much emphasis has been given to this factor in the field of cartilage biology, in clinical practice such defects are rarely considered an indication for chondral


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surgery, since they are often asymptomatic and the evidence available today does not indicate that they predispose to cartilage loss and OA progression.7±10 Indeed, chondral surgery, such as autologous chondrocyte implantation or microfracture, is usually reserved for chronic, symptomatic, full thickness chondral or osteochondral defects, especially where injury is the most likely cause of the symptoms.10±13

5.2.1

The aetiology of JSI

The most important risk factors in the development of cartilage loss in OA development are age, trauma, malalignment, meniscal or cruciate ligament injury, and a family history.14±17 Such factors point towards mechanical factors contributing to JSD development. Although we generally attribute meniscal injury as causing OA through joint destabilisation, the clinical data for this are missing. For instance joint instability in benign hypermobility syndrome does not predispose to OA, nor is it the case that individuals with repair to their menisci have a reduce risk of developing OA compared with those who have not had a repair (Lohmander, personal communication). Trauma has traditionally been regarded as the most important aetiological factor in the development of focal chondral or osteochondral defects.18 However, in a large study of 1000 consecutive arthroscopies, 39% of patients with a focal defect in their knee cartilage failed to remember a previous traumatic episode to their joint.5 In addition, it has been shown that up to 43% of healthy subjects without a family history of OA have knee chondral lesions as evaluated by magnetic resonance imaging (MRI).19 These data point to the fact that chondral or osteochondral defects are more common than previously thought and that a traumatic episode may not always be apparent from the history.

5.2.2

The natural history of JSI

Over 250 years ago Hunter stated: Ìf we consult the standard Chirurgical Writers from Hippocrates down to the present Age, we shall find, that an ulcerated Cartilage is universally allowed to be a very troublesome disease and when destroyed, it is never recovered.'20 Of course, this statement most likely referred to severely symptomatic lesions that had acquired a chronic and disabling course, leading patients to the attention of a surgeon. Through the years, however, such a paradigm has extended to all chondral injuries. This is mainly because of the lack of diagnostic tools capable of identifying, and prospectively following up smaller and, in particular, acute lesions. It is also the case that published studies are capable of fuelling intuitive preconceptions; reporting, for instance, that the risk of developing OA by the age of 65 is 13% in individuals with a history of trauma compared with 6% in those without a history of trauma.21 As a consequence the cartilage biology/repair literature

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often concludes that `cartilage lesions never heal spontaneously', and that they necessarily `predispose to osteoarthritis'. The recent development of cell-based therapies for the repair of chondral lesions such as microfracture, mosaicplasty and autologous chondrocyte implantation (ACI) has generated interest in the natural history of these lesion to define indications for such invasive procedures.12 Improved cartilage imaging using MRI has contributed significantly to the generation of longitudinal data in all groups of patients. In 1996, Messner and Maletius8 reported that 22 out of 28 young athletes who had an isolated chondral injury in a weight-bearing part of their knees diagnosed by arthroscopy had good or excellent knee function at 14 years follow-up as evaluated clinically and radiographically. No specific treatment had been preformed except Pridie drilling (similar to microfracture) in three cases and occasional debridement. Twenty-one patients were able to return to pre-injury level sports activities. Although at the end of follow-up 12 patients had some radiographic joint space reduction. No control group (without JSD) was included and therefore we do not know whether joint space reduction would have occurred in the absence of a JSD. In this study, all patients had an isolated Outerbridge grade 2 (most cases) or 3 chondral defect (diameter > 1 cm), without any damage to other joint structures including menisci, ligaments and the remaining cartilage. No patient had instability, or a previous history of knee surgery. What we learn from this study is that isolated chondral or osteochondral lesions, in young active patients, in otherwise healthy knees, have a favourable natural history leading to long-term functional restoration. We do not know whether (good) structural repair is required for functional outcome or whether these lesions become asymptomatic or repair with scar tissue. In either case, such a good outcome after 14 years follow-up in 78% of the lesions, suggest that an aggressive approach for all such lesions is not justified. The study discussed above focused on isolated (osteo) chondral defects in otherwise normal knees. However, isolated chondral defects are present in only 36.6% of symptomatic knees requiring arthroscopy. In the majority of cases, other lesions, including those to menisci4 or ligaments,9,22 co-exist. In a longitudinal study, Shelbourne et al.9 asked the question whether the presence of a chondral injury detected in young athletes undergoing ACL reconstruction modifies the clinical outcome. In this study they selected two groups of such patients, one that had a single chondral or osteochondral injury at the time of arthroscopy, and an age and sex matched group who had no chondral injury. The cartilage injury was left untreated, and the clinical outcome was monitored for 8.7 years, clinically and radiographically. Throughout follow-up, the patients with a chondral injury had more subjective symptoms that those without. This difference was statistically significant, but small in size and more than 79% of patients returned to pre-injury levels of sports activities involving jumping, twisting and pivoting. The radiological score was not different in the two groups. There was no correlation between the size of the defect and the


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outcome, although the size of some defects might have been underestimated since lesions were not debrided. Finally, they reported that, in each individual patient, the severity of symptoms fluctuated significantly during the follow-up. Again, there was no information as to whether structural repair of the chondral injury was a prerequisite for good clinical outcome. The recent improvement in imaging of the articular cartilage with 3D fast spin-echo, or fat suppressed spoiled gradient-echo MRI have allowed detection of chondral lesions diagnosed at arthroscopy with a sensitivity approaching 95%, and specificity sometimes reaching 100%.23±27 MRI has therefore allowed monitoring of chondral defects to obtain prospective clinical and structural outcomes in symptomatic and asymptomatic groups. These studies have yielded surprising results. Ding et al.28 reported a longitudinal study in which the presence and the natural history of chondral defects had been studied by MRI in asymptomatic subjects with a family history of joint replacement, and in an age and sex matched population without family history of OA. This study revealed that 43% of the subjects without a family history of OA and 57% of subjects with a family history of OA19 had chondral defects. At 2.3 years follow-up, 33% of all subjects had a worsening of the defects as graded by MRI, 37% an improvement and the rest remained stable. A worse outcome was associated with female sex, age and body mass index at baseline. Although factors associated with the reproducibility of the MRI grading may have contributed to the defect variation, in general, measurement error was considered to be very low. Importantly, only 18% of the subjects with a cartilage defect had a history of knee trauma. These data show three very important points. Firstly, that chondral defects, including full thickness ones, are often asymptomatic; secondly, that the majority of these lesions may not be related to traumatic injury as previously thought. Thirdly, and most importantly, a number of these lesions may improve (and possibly heal) spontaneously. The presence of chondral defects in these patients predicted a rate of cartilage loss of 2±3% instead of 1±2% of subjects without chondral defects.7 Since the rate of cartilage loss is an independent predictor of joint replacement in patients with OA,29 it is arguable that at least a number of such asymptomatic defects may predispose to OA. In a separate paper, Davies-Tuck et al. reported on the natural history of similar chondral lesions in a cohort of patients with OA.30 In this cohort, chondral injuries worsened in 81% of the cases and improved in only 4% over 2 years. In a similar prospective study, Wluka et al. showed that the presence of cartilage defects in patients with OA was associated with disease severity, correlated with the rate of cartilage loss within 2 years and was a predictor of joint replacement within 4 years.31 We could summarise these data by saying that cartilage defects are often but not always due to acute mechanical injury, and can complicate and accelerate the course of OA, but may be present in otherwise normal knees, where, at least

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in some cases, they may accelerate the physiological rate of cartilage loss that takes place after the age of 40 years. Importantly, particularly in the absence of OA, some of these defects may undergo at least a partial healing. Although age, female sex and body mass index are generally associated with progression, it is not presently possible to predict the outcome of one individual defect. These considerations have important clinical consequences in deciding when to treat a chondral defect. Although general common sense would suggest that chronic, symptomatic, isolated defects are a good indication for interventions such as microfracture or autologous chondrocyte transplantation, in most cases we do not have good evidence that defects are associated with progression to OA. Nor do we know whether asymptomatic isolated defects go on to become chronic and predispose to cartilage loss and OA. There is therefore a clinical need for criteria or biomarkers that could predict the outcome of JSD accurately. Such tools would not only be precious for the clinician, but would also improve the sensitivity of clinical trials and therefore avoid the floor effect due to a number of patients who might improve spontaneously in the absence of treatment.

5.2.3

Chronic joint injury

Chronic injuries include repetitive trauma, excessive loading, as well as direct and indirect altered biomechanics. All these factors have been associated with cartilage loss, pain and disability.14 Subjects exposed to heavy work such as farming and repetitive lifting of heavy weight, as well as athletes practising sports involving high levels of twisting and torsional loading are at higher risk of developing OA.32±36 It is worth stressing, however, that both heavy work and sport activities also predispose to meniscal and ligament injuries, which are independent risks for cartilage loss.37 The way in which damage to menisci and ligaments predisposes to OA is thought to be due to loss of cartilage protection, and creation of joint instability, leading to increased wear of the articulating surfaces.14,38,39 Certainly there is evidence that patients with such lesions have an increased risk of developing OA.37 Indeed, even simple extrusion of the medial meniscus is a risk factor in for progression of knee OA, and is predictive of joint replacement.38,39 Malalignment is another important risk factor for cartilage loss and OA.37,40 Cartilage loss occurs in dysplastic joints in which weight-bearing is not evenly distributed.14,40 Interestingly, even in the case of chronic cartilage ulcers in adult individuals with varus knees, correcting malalignment by tibial osteotomy was able to partially restore the articular surface,40±44 though mostly by fibrocartilage.40 How acute or chronic mechanical injury results in progressive breakdown of the cartilage tissue is not completely understood. Ultimately, the degradation of the cartilage matrix is believed to be mediated by enzymes such as


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metalloproteinases, aggrecanases and other proteolytic enzymes, which may be produced by inflamed synovium, by infiltrating haematopoietic cells,45 and also by injured chondrocytes.45±47

5.3

Animal models of cartilage injury

The development of animal models has contributed enormously to our understanding of the pathophysiology of cartilage injury. Injury in such models can be described as direct or indirect, and are discussed below.

5.3.1

Direct injury to the articular surface

Despite the earlier observations of Hunter in the 18th century, Kolliker and Paget were the first to recognise the ability of cartilage lesions to mend. At the time they attributed repair to cells outside the articular cartilage (reviewed in Campbell48). These early observations heralded an explosion in experimental in vivo cartilage injury models, which became a fashionable scientific pursuit for the next 150 years. Many of the seminal papers in this field are derived from these early years. Peter Redfern in 1851 (reprinted in 196949) was one of the first to describe the tissue response following incisions to the articular cartilage of dogs. Redfern's observations apparently contradicted the concensus view, by demonstrating that cuts in articular cartilage were capable of ùniting with a highly cellular, disorganised, granular mass'. He described a reactive chondrocyte phenotype; superficial cells were enlarged and those close to the lesion showed evidence of proliferation. He concluded that there was `no longer . . . the slightest doubt that wounds in articular cartilages are capable of perfect union by formation of fibrous tissue out of the texture of the cut surfaces'. Whilst these findings have held true over the years, his interpretation of the repair response was probably incorrect. Shands in 1931 observed that there was a significant difference in repair responses between full thickness defects, where the cut extended across the osteochondral junction, and superficial lesions where the cut did not breech the junction.50 In the 1970s, Meachim proposed that there were two responses to articular cartilage injury; an intrinsic and extrinsic repair response. Extrinsic repair was stimulated by bone marrow cells when the cut had breeched the osteochondral junction, and it was these cells that contributed to what we now regard as a fibrocartilage scar.51 Intrinsic repair was by cells of the cartilage and was more effective at generating a scar-free hyaline cartilage between cut surfaces, even though Bennett and co-workers recognised that `the powers of such regeneration are feeble and not always demonstrable'.52,53 Calandruccio went further to suggest that both sorts of lesions were capable of stimulating intrinsic repair, but that intrinsic repair was suppressed in the presence of a dominant extrinsic response from the bone marrow.54 What these studies as well as those of others clearly indicated was that there was

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considerable variability in repair responses in individual animals which could not, and has not, been explained. Nonetheless, these studies collectively demonstrated that cartilage tissue was able to stimulate a repair response. In 1963, Meachim performed a comprehensive study following the effects of superficial scarification of articular cartilage in adult rabbits. On histological examination of the tissue and by sulphate incorporation, he found an early increase in metachromasia around superficial chondrocytes near the cuts, and subsequent loss of inter-territorial proteoglycan over the next few weeks. Cell clusters were apparent at 16 weeks especially in the deep layers.51 Cell clustering was shown to be due to chondrocyte division by Mankin55 who concluded that this represented intrinsic repair, albeit late, and unlikely to be contributing to healing in the presence of a more rapid extrinsic reaction. More recent studies have confirmed these original observations but few have added much more to how cartilage repair is controlled and coordinated. Shapiro et al.56 generated small osteochondral defects in young rabbit cartilage, which resulted in invasion of bone marrow cells into the lesion within 2 weeks of injury. Interestingly, the process that led to restoration of the articular surface within 24 weeks involved a coordinated and polarised morphogenetic process highly reminiscent of endochondral bone formation during development. This experiment suggested that mechanisms similar to those governing embryonic skeletogenesis were reactivated during postnatal repair processes. This finding is supported by the observation that mutations or allelic variants of several morphogens and growth factors known to play a role in skeletal development are associated with an increased risk of OA.57±62 Wei et al.63 studied repair responses in rabbits of different ages, from adolescent to fully mature, and reported results similar to that of Shapiro et al. in young rabbits. In adult rabbits the outcome of repair was considerably worse, thereby paralleling clinical data in humans. The role of inflammatory cells in cartilage injury was suggested by Hembry et al. when they demonstrated macrophage recruitment to the damaged surface.45 They proposed that these cells were responsible for release of MMP9, which probably activated other metalloproteinases of joint origin, causing activation of chondrocytic MMP-3 and MMP-13, as well as aggrecanases, at the site of injury. Matrix metalloproteinase (MMP) inhibitors such as tissue inhibiting metalloproteinase 1 (TIMP-1) were also induced by injury suggesting a tight regulation of proteolysis. The subchondral bone in adult animals has also been shown to react promptly to cartilage injury with activation of intense remodelling. In a goat model, Vasara et al. reported that, even partial thickness lesions caused extensive bone remodelling with distortion of the trabeculae and reduced bone volume.64 Towards the end of the 20th century a number of groups tested the responses induced by injurious loading of the joint. This might be regarded as more physiological in vivo damage. Donohue et al. impact loaded closed canine joints


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and studied the histological response 2, 4 and 6 weeks later.65 They found that there was clonal expansion of cells in the cartilage, with increased vascular invasion and increased water content. They also calculated a 40% reduction in proteoglycan associated with collagen. A similar closed joint transarticular load to the patella was used by Thompson et al. who extended the period of observation to 24 weeks post-injury. They described the presence of fissures in the cartilage, and late loss of proteoglycan in the tissue associated with deep clefts.66 Haut and co-workers added that there was significant softening of the articular cartilage early post-impact,67 and in a follow-up study.68 They determined that there was increased tissue thickness in the first 5 months following injury, but by 36 months a significant (45%) reduction in tissue thickness was measured. They also documented a significant increase in the thickness of the subchrondral bone. Whilst the histological chronology of articular cartilage lesions has been well characterised over the past 200 years, the molecular and cellular mechanisms that control such events are far from clear. We may be able to attribute this in part to the recent decline in the use of such in vivo models at a time when molecular tools were becoming increasingly sensitive. As experimental JSI was becoming unfashionable, many were turning their attention to indirect models of cartilage damage in vivo.

5.3.2

Indirect cartilage injury

Modern approaches to look at cartilage injury in vivo have tended to involve indirect injury, through, for instance, joint destabilisation. Such models have the advantage of more closely mimicking clinical osteoarthritis. Joint instability can be induced through surgery (a list of some such models is given in Table 5.1), or through enzymatic treatment of the joint with collagenase,69 which is thought to weaken the supporting ligaments. It is worth stressing that all of these models, to a greater or lesser extent, induce joint inflammation, and so cartilage injury is due to a combination of inflammation and mechanical factors. Table 5.1 Surgical models of osteoarthritis Animal

Method of induction

Rabbit

Medial meniscectomy70 Meniscectomy and cruciate transaction71

Dog

Cruciate transection (Pond-Nuki)72

Guinea pig

Gluteal resection (myectomy and tendotomy)73

Mouse

Partial meniscectomy and transaction of med collateral ligament74 Meniscotibial ligament transaction (DMM)75

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One recently described model deserves detailed attention. This murine model, which is induced by destabilisation of the medial meniscus (DMM), was described by Glasson et al. in 2005.75 A robust and predictable degeneration of the articular cartilage is seen in male mice within 4±8 weeks of transection of the medial menisco-tibial ligament. Synovitis is rarely present, and bone changes are a late feature. As meniscal injury is a strong risk factor for the development of OA in humans, this model is highly clinically relevant.37,39 There are a number of advantages in using this model over other historical ones: · · · · · · ·

100% penetrance (all male mice develop disease); cartilage loss from 4 weeks post-surgery; mice are cheap and easy to house; mouse experimentation is less emotive than that using higher species; can combine model in mice which have been genetically modified; less inflammatory than other historical models of cartilage degeneration; could be used for drug screening.

Of paramount importance for understanding pathogenesis of OA and early responses to cartilage injury is the fact that this model allows one to combine OA progression in genetically modified mice. This was done to good effect by Glasson et al., who demonstrated that mice deficient in ADAMTS5, but not ADAMTS4 (aggrecan degrading enzymes in cartilage) were partially protected from osteoarthritis.75 This not only told us that cartilage degeneration was dependent upon aggrecanase activity, but also that ADAMTS5 was the principal cartilage aggrecanase in mice. The model has provided functional data on a large number of molecules in addition to ADAMTS5.76 Partial protection was seen in mice deficient in IL-1 beta, although unexpectedly MMP9 null mice developed accelerated cartilage degradation after DMM surgery, suggesting that it was protective in vivo. Quoting Sonya Glasson, àlthough the results in the mouse will not always transpose to the human condition, the track record of mouse knockouts corresponding to the human phenotype have been excellent'.76

5.4

In vitro cartilage injury

Much recent interest has focused on the responses of articular cartilage to injury in vitro. For the sake of simplicity we will focus on direct mechanical injury rather than chemical or inflammatory. There are a number of ways of inducing cartilage injury including explantation, re-cutting, impact loading and high magnitude (low velocity) loading. Each of these will be addressed in more detail below.

5.4.1

Explantation

Explantation is perhaps the `purest' injury in cartilage, because it is closest to how injury might be perceived in vivo. The articular surface of cartilage is


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normally bathed in synovial fluid which contains plasma proteins as well as some of the cellular constituents of blood. It is hardly surprising that washing plasma proteins from cartilage explants (which occurs when explants are cultured in medium) will change the response of the tissue to different stimuli. Although direct comparisons of explantation versus re-cutting cartilage explants in vitro have not been performed, some observations have been made. For instance, there is a strong activation of the extracellularly regulated kinsae (ERK), p38 and c-jun-terminal kinase (JNK), members of the mitogen activated protein (MAP) kinases upon explantation.77 The activation of the ERK pathway is due to release of FGF2, which also occurs when explants are rested then recut.78 Activation of JNK and p38 does not occur on re-cutting,78 and does not appear to be due to the release of a soluble factor (J. Saklatvala, personal communication). Activation of inflammatory MAP kinases upon explantation is sufficient to activate inflammatory response genes such as IL-1 and at the mRNA and protein level.77

5.4.2

Re-cutting

Re-cutting cartilage is usually performed when the cartilage has been explanted into medium (either serum containing or serum free) and rested for a variable amount of time ± anywhere from 12 hours to 7 days. The rationale for this experimental system is to allow any cellular response which occurred upon explantation to return to baseline before the cartilage is re-cut. The timing of the re-cutting is probably critical as a recent gene array study has demonstrated that it takes 7 days for chondrocyte gene expression to return to in vivo (resting) levels.79 In this study the authors rested human articular cartilage in medium for 7 days prior to re-cutting. They identified 690 genes that were significantly regulated at least two-fold following re-cutting. These included genes previously reported to be differentially expressed in OA versus normal cartilage, or having allelic variants genetically linked to OA. A systematic analysis of the Wnt signalling pathway revealed up-regulation of Wnt-16, down-regulation of FRZB, and up-regulation of Wnt target genes. They were able to demonstrate increased Wnt pathway activity in wounded as well as osteoarthritic cartilage, by demonstrating accumulation of cellular beta catenin.

5.4.3

Impact loading

Conservative mechanobiologists make a clear distinction between injurious mechanical loading and impact loading.80 The latter is characteristically a high velocity load with consequent cellular and matrix damage in the absence of significant tissue strain (the amount the tissue deforms). Such injury to cartilage is a feature of acute mechanical trauma to cartilage, say following a road traffic accident, or a fall from a height. Load is often delivered via a drop tower and is

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usually a single impact. Such injury causes cell loss associated with microscopic splitting in the superficial layer of the cartilage and early surface delamination.81 This same group also studied the effect of impact load on sulphate incorporation and loss of proteoglycan from the matrix. They found that in the 2 weeks following impact, 25±40% of sulphated proteoglycan was lost from the tissue. Although recovery was associated with an increase in proteoglycan synthesis, synthetic responses were suppressed early following injury.82

5.4.4

Low velocity injurious load

Low velocity injuries tend to be characterised by high strain (say 50%) with relatively low velocities, e.g. 100%/s. Such cartilage injury also causes significant proteoglycan loss in the first few days following load.83 Loss of proteoglycan was determined to be independent of MMP activity in these early stages, although later losses could be suppressed by the presence of MMP inhibitors.84 Microarray analysis of injured cartilage showed strong upregulation of MMP3 and ADAMTS5, as well as the negative regulator of metalloproteinase activity, suggesting that these enzymes may be responsible for active proteolytic degradation of tissue proteoglycan.47 Another response to cartilage injury that has been extensively investigated in vitro is cell death. From the early observations of Redfern, Bennett and others, cell death along the cut surface has been a robust feature. Tew et al. determined that cell death in explants following trephine injury was due to a combination of apoptosis and necrosis. They also were able to show increased cell proliferation in the regions adjacent to the cut surface.85 The same group went on to show that cartilage wounding responses were essentially similar between immature and mature cartilage,86 but that increased cell death was a feature of blunt wounds rather more so than those generated by a scalpel blade.87 Others have shown similar results in cartilage explants subjected to injurious load.88,89 Quinn et al. observed a mixture of cell death and enlarged viable cells adjacent to the dead chondrocytes, which, by autoradiography, showed increased proteoglycan synthesis. There is little doubt that such a loading regime induces significant cellular and tissue damage. Stevens et al. studied the pattern of secreted proteins 5 days following injurious load of bovine explants. They identified a number of proteins by mass spectrometry of sodium dodecyl sulphate±polyacrylamide gel electrophoresis (SDS-PAGE) separated proteins. Their main findings were the release of a large number of intracellular proteins indicating significant cell injury, as well as release of fragments of matrix proteins such as type VI collagen, dermatan sulphate proteoglycan 3 and fibronectin implying matrix damage.90


5.5

149

Conclusions

Chondral injury in patients is still poorly understood, although recent emerging data suggest that small chondral defects may have the capacity to heal spontaneously. It remains the case that joint surface injury is a risk factor for development of OA probably through both the mechanical disruption of the tissue, cell death, as well as through activation of proteolytic pathways. The early in vivo injury observations of Redfern, Calandruccio, Meachim, Mankin and others have made an unrivalled contribution to our understanding of how cartilage responds to injury. In view of the advances in modern science over recent years it is perhaps surprising that our understanding of the cellular and molecular events of cartilage injury is not greater. A small number of novel molecular pathways of cartilage injury have been described. Such molecules are likely to be important in the development of molecular tools to support repair and, at the same time, may help identify those patients who would benefit from chondral surgery/tissue engineering. An exciting challenge in such studies will be finding strategies that not only stimulate production of hyaline cartilage, but that will allow integration of neocartilage into the existing matrix.

5.6

References

1 Bouchard, B.A., and Tracy, P.B. 2001. Platelets, leukocytes, and coagulation. Curr Opin Hematol 8: 263±269. 2 Martin, P., and Leibovich, S.J. 2005. Inflammatory cells during wound repair: the good, the bad and the ugly. Trends Cell Biol 15: 599±607. 3 Stramer, B.M., Mori, R., and Martin, P. 2007. The inflammation-fibrosis link? A Jekyll and Hyde role for blood cells during wound repair. J Invest Dermatol 127: 1009±1017. 4 Curl, W.W., Krome, J., Gordon, E.S., Rushing, J., Smith, B.P., and Poehling, G.G. 1997. Cartilage injuries: a review of 31,516 knee arthroscopies. Arthroscopy 13: 456±460. 5 Hjelle, K., Solheim, E., Strand, T., Muri, R., and Brittberg, M. 2002. Articular cartilage defects in 1,000 knee arthroscopies. Arthroscopy 18: 730±734. 6 Ding, C., Cicuttini, F., and Jones, G. 2008. How important is MRI for detecting early osteoarthritis? Nat Clin Pract Rheumatol 4: 4±5. 7 Ding, C., Cicuttini, F., Scott, F., Boon, C., and Jones, G. 2005. Association of prevalent and incident knee cartilage defects with loss of tibial and patellar cartilage: a longitudinal study. Arthritis Rheum 52: 3918±3927. 8 Messner, K., and Maletius, W. 1996. The long-term prognosis for severe damage to weight-bearing cartilage in the knee: a 14-year clinical and radiographic follow-up in 28 young athletes. Acta Orthop Scand 67: 165±168. 9 Shelbourne, K.D., Jari, S., and Gray, T. 2003. Outcome of untreated traumatic articular cartilage defects of the knee: a natural history study. J Bone Joint Surg Am 85-A Suppl 2: 8±16. 10 Smith, G.D., Knutsen, G., and Richardson, J.B. 2005. A clinical review of cartilage repair techniques. J Bone Joint Surg Br 87: 445±449.

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28 Ding, C., Cicuttini, F., Scott, F., Cooley, H., Boon, C., and Jones, G. 2006. Natural history of knee cartilage defects and factors affecting change. Arch Intern Med 166: 651±658. 29 Cicuttini, F.M., Jones, G., Forbes, A., and Wluka, A.E. 2004. Rate of cartilage loss at two years predicts subsequent total knee arthroplasty: a prospective study. Ann Rheum Dis 63: 1124±1127. 30 Davies-Tuck, M.L., Wluka, A.E., Wang, Y., Teichtahl, A.J., Jones, G., Ding, C., and Cicuttini, F.M. 2008. The natural history of cartilage defects in people with knee osteoarthritis. Osteoarthritis Cartilage 16: 337±342. 31 Wluka, A.E., Ding, C., Jones, G., and Cicuttini, F.M. 2005. The clinical correlates of articular cartilage defects in symptomatic knee osteoarthritis: a prospective study. Rheumatology (Oxford) 44: 1311±1316. 32 Anderson, J.J., and Felson, D.T. 1988. Factors associated with osteoarthritis of the knee in the first national Health and Nutrition Examination Survey (HANES I). Evidence for an association with overweight, race, and physical demands of work. Am J Epidemiol 128: 179±189. 33 Buckwalter, J.A., and Lane, N.E. 1997. Athletics and osteoarthritis. Am J Sports Med 25: 873±881. 34 Croft, P., Coggon, D., Cruddas, M., and Cooper, C. 1992. Osteoarthritis of the hip: an occupational disease in farmers. BMJ 304: 1269±1272. 35 Felson, D.T., Hannan, M.T., Naimark, A., Berkeley, J., Gordon, G., Wilson, P.W., and Anderson, J. 1991. Occupational physical demands, knee bending, and knee osteoarthritis: results from the Framingham Study. J Rheumatol 18: 1587±1592. 36 Yoshimura, N., Sasaki, S., Iwasaki, K., Danjoh, S., Kinoshita, H., Yasuda, T., Tamaki, T., Hashimoto, T., Kellingray, S., Croft, P., et al. 2000. Occupational lifting is associated with hip osteoarthritis: a Japanese case-control study. J Rheumatol 27: 434±440. 37 Lohmander, L.S., Englund, P.M., Dahl, L.L., and Roos, E.M. 2007. The long-term consequence of anterior cruciate ligament and meniscus injuries: osteoarthritis. Am J Sports Med 35: 1756±1769. 38 Ding, C., Martel-Pelletier, J., Pelletier, J.P., Abram, F., Raynauld, J.P., Cicuttini, F., and Jones, G. 2007. Meniscal tear as an osteoarthritis risk factor in a largely nonosteoarthritic cohort: a cross-sectional study. J Rheumatol 34: 776±784. 39 Ding, C., Martel-Pelletier, J., Pelletier, J.P., Abram, F., Raynauld, J.P., Cicuttini, F., and Jones, G. 2007. Knee meniscal extrusion in a largely non-osteoarthritic cohort: association with greater loss of cartilage volume. Arthritis Res Ther 9: R21. 40 Koshino, T., Wada, S., Ara, Y., and Saito, T. 2003. Regeneration of degenerated articular cartilage after high tibial valgus osteotomy for medial compartmental osteoarthritis of the knee. Knee 10: 229±236. 41 Fujisawa, Y., Masuhara, K., and Shiomi, S. 1979. The effect of high tibial osteotomy on osteoarthritis of the knee. An arthroscopic study of 54 knee joints. Orthop Clin North Am 10: 585±608. 42 Matsui, N., Moriya, H., and Kitahara, H. 1979. The use of arthroscopy for follow-up in knee joint surgery. Orthop Clin North Am 10: 697±708. 43 Odenbring, S., Egund, N., Lindstrand, A., Lohmander, L.S., and Willen, H. 1992. Cartilage regeneration after proximal tibial osteotomy for medial gonarthrosis. An arthroscopic, roentgenographic, and histologic study. Clin Orthop Relat Res 277: 210±216. 44 Schultz, W., and Gobel, D. 1999. Articular cartilage regeneration of the knee joint after proximal tibial valgus osteotomy: a prospective study of different intra- and

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Regenerative medicine and biomaterials in connective tissue repair extra-articular operative techniques. Knee Surg Sports Traumatol Arthrosc 7: 29±36. Hembry, R.M., Dyce, J., Driesang, I., Hunziker, E.B., Fosang, A.J., Tyler, J.A., and Murphy, G. 2001. Immunolocalization of matrix metalloproteinases in partialthickness defects in pig articular cartilage. A preliminary report. J Bone Joint Surg Am 83-A: 826±838. Flannelly, J., Chambers, M.G., Dudhia, J., Hembry, R.M., Murphy, G., Mason, R.M., and Bayliss, M.T. 2002. Metalloproteinase and tissue inhibitor of metalloproteinase expression in the murine STR/ort model of osteoarthritis. Osteoarthritis Cartilage 10: 722±733. Lee, J.H., Fitzgerald, J.B., Dimicco, M.A., and Grodzinsky, A.J. 2005. Mechanical injury of cartilage explants causes specific time-dependent changes in chondrocyte gene expression. Arthritis Rheum 52: 2386±2395. Campbell, C.J. 1969. The healing of cartilage defects. Clin Orthop Related Res 64: 45± 63. Redfern, P. 1851 reprinted in 1969. On the healing of wounds in articular cartilage. Clin Orthop 64: 4±6. Shands, A.R. 1931. Regeneration of hyaline cartilage in joints. Arch Surg 22: 137. Meachim, G. 1963. The effect of scarification on articular cartalige in the rabbit. J Bone Joint Surgery 45B: 150±161. Bennett, G.A., and Bauer, W. 1935. Further studies concerning the repair of articular cartilage in dog joints. Bone Joint Surg 17: 141. Bennett, G.A., Bauer, W., and Maddock, S.J. 1932. A study of the repair of articular cartilage and reactions of normal joints of adult dogs to surgically created defects of articular cartilage, `joint mice', and patellar displacement. Am J Pathol 8: 499. Calandruccio, R.A., and Gilmer, W.S. 1962. Proliferation, regeneration, and repair of articular cartilage of immature animals. J Bone Joint Surg 44A: 431. Mankin, H.J. 1962. Localisation of tritiated thymidine in articular cartilage of rabbits. II: Repair in immature cartilage. J Bone Joint Surg 44A: 688. Shapiro, F., Koide, S., and Glimcher, M.J. 1993. Cell origin and differentiation in the repair of full-thickness defects of articular cartilage. J Bone Joint Surg Am 75: 532±553. Chapman, K., Takahashi, A., Meulenbelt, I., Watson, C., Rodriguez-Lopez, J., Egli, R., Tsezou, A., Malizos, K.N., Kloppenburg, M., Shi, D., et al. 2008. A metaanalysis of European and Asian cohorts reveals a global role of a functional SNP in the 50 UTR of GDF5 with osteoarthritis susceptibility. Hum Mol Genet May 15; 17, 10: 1497±1504. Loughlin, J., Dowling, B., Chapman, K., Marcelline, L., Mustafa, Z., Southam, L., Ferreira, A., Ciesielski, C., Carson, D.A., and Corr, M. 2004. Functional variants within the secreted frizzled-related protein 3 gene are associated with hip osteoarthritis in females. Proc Natl Acad Sci USA 101: 9757±9762. Loughlin, J. 2006. Osteoarthritis linkage scan: more loci for the geneticists to investigate. Ann Rheum Dis 65: 1265±1266. Miyamoto, Y., Mabuchi, A., Shi, D., Kubo, T., Takatori, Y., Saito, S., Fujioka, M., Sudo, A., Uchida, A., Yamamoto, S., et al. 2007. A functional polymorphism in the 50 UTR of GDF5 is associated with susceptibility to osteoarthritis. Nat Genet 39: 529±533. Southam, L., Rodriguez-Lopez, J., Wilkins, J.M., Pombo-Suarez, M., Snelling, S., Gomez-Reino, J.J., Chapman, K., Gonzalez, A., and Loughlin, J. 2007. An SNP in the 50 -UTR of GDF5 is associated with osteoarthritis susceptibility in Europeans and with in vivo differences in allelic expression in articular cartilage. Hum Mol Genet 16: 2226±2232.


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62 Valdes, A.M., Van Oene, M., Hart, D.J., Surdulescu, G.L., Loughlin, J., Doherty, M., and Spector, T.D. 2006. Reproducible genetic associations between candidate genes and clinical knee osteoarthritis in men and women. Arthritis Rheum 54: 533±539. 63 Wei, X., Gao, J., and Messner, K. 1997. Maturation-dependent repair of untreated osteochondral defects in the rabbit knee joint. J Biomed Mater Res 34: 63±72. 64 Vasara, A.I., Hyttinen, M.M., Lammi, M.J., Lammi, P.E., Langsjo, T.K., Lindahl, A., Peterson, L., Kellomaki, M., Konttinen, Y.T., Helminen, H.J., et al. 2004. Subchondral bone reaction associated with chondral defect and attempted cartilage repair in goats. Calcif Tissue Int 74: 107±114. 65 Donohue, J.M., Buss, D., Oegema, T.R., Jr, and Thompson, R.C., Jr 1983. The effects of indirect blunt trauma on adult canine articular cartilage. J Bone Joint Surg Am 65: 948±957. 66 Thompson, R.C., Jr., Oegema, T.R., Jr., Lewis, J.L., and Wallace, L. 1991. Osteoarthritic changes after acute transarticular load. An animal model. J Bone Joint Surg Am 73: 990±1001. 67 Haut, R.C., Ide, T.M., and De Camp, C.E. 1995. Mechanical responses of the rabbit patello-femoral joint to blunt impact. J Biomech Eng 117: 402±408. 68 Ewers, B.J., Weaver, B.T., Sevensma, E.T., and Haut, R.C. 2002. Chronic changes in rabbit retro-patellar cartilage and subchondral bone after blunt impact loading of the patellofemoral joint. J Orthop Res 20: 545±550. 69 Rudolphi, K., Gerwin, N., Verzijl, N., van der Kraan, P., and van den Berg, W. 2003. Pralnacasan, an inhibitor of interleukin-1beta converting enzyme, reduces joint damage in two murine models of osteoarthritis. Osteoarthritis Cartilage 11: 738± 746. 70 Moskowitz, R.W., Davis, W., Sammarco, J., Martens, M., Baker, J., Mayor, M., Burstein, A.H., and Frankel, V.H. 1973. Experimentally induced degenerative joint lesions following partial meniscectomy in the rabbit. Arthritis Rheum 16: 397±405. 71 Telhag, H., and Lindberg, L. 1972. A method for inducing osteoarthritic changes in rabbits' knees. Clin Orthop Relat Res 86: 214±223. 72 Pond, M.J., and Nuki, G. 1973. Experimentally-induced osteoarthritis in the dog. Ann Rheum Dis 32: 387±388. 73 Layton, M.W., Goldstein, S.A., Goulet, R.W., Feldkamp, L.A., Kubinski, D.J., and Bole, G.G. 1988. Examination of subchondral bone architecture in experimental osteoarthritis by microscopic computed axial tomography. Arthritis Rheum 31: 1400±1405. 74 Clements, K.M., Price, J.S., Chambers, M.G., Visco, D.M., Poole, A.R., and Mason, R.M. 2003. Gene deletion of either interleukin-1beta, interleukin-1beta-converting enzyme, inducible nitric oxide synthase, or stromelysin 1 accelerates the development of knee osteoarthritis in mice after surgical transection of the medial collateral ligament and partial medial meniscectomy. Arthritis Rheum. 48: 3452± 3463. 75 Glasson, S.S., Askew, R., Sheppard, B., Carito, B., Blanchet, T., Ma, H.L., Flannery, C.R., Peluso, D., Kanki, K., Yang, Z., et al. 2005. Deletion of active ADAMTS5 prevents cartilage degradation in a murine model of osteoarthritis. Nature. 434: 644± 648. 76 Glasson, S.S. 2007. In vivo osteoarthritis target validation utilising geneticallymodified mice. Curr Drug Targets 8: 367±376. 77 Gruber, J., Vincent, T.L., Hermansson, M., Bolton, M., Wait, R., and Saklatvala, J. 2004. Induction of interleukin-1 in articular cartilage by explantation and cutting. Arthritis Rheum. 50: 2539±2546.

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Vincent, T., Hermansson, M., Bolton, M., Wait, R., and Saklatvala, J. 2002. Basic FGF mediates an immediate response of articular cartilage to mechanical injury. Proc Natl Acad Sci USA 99: 8259±8264. 79 Dell'accio, F., De Bari, C., Eltawil, N.M., Vanhummelen, P., and Pitzalis, C. 2008. Identification of the molecular response of articular cartilage to injury, by microarray screening: Wnt-16 expression and signaling after injury and in osteoarthritis. Arthritis Rheum 58: 1410±1421. 80 Aspden, R.M., Jeffrey, J.E., and Burgin, L.V. 2002. Impact loading of articular cartilage. Osteoarthritis Cartilage 10: 588±589; author reply 590. 81 Jeffrey, J.E., Gregory, D.W., and Aspden, R.M. 1995. Matrix damage and chondrocyte viability following a single impact load on articular cartilage. Arch Biochem Biophys 322: 87±96. 82 Jeffrey, J.E., Thomson, L.A., and Aspden, R.M. 1997. Matrix loss and synthesis following a single impact load on articular cartilage in vitro. Biochim Biophys Acta 1334: 223±232. 83 Patwari, P., Cheng, D.M., Cole, A.A., Kuettner, K.E., and Grodzinsky, A.J. 2007. Analysis of the relationship between peak stress and proteoglycan loss following injurious compression of human post-mortem knee and ankle cartilage. Biomech Model Mechanobiol 6: 83±89. 84 DiMicco, M.A., Patwari, P., Siparsky, P.N., Kumar, S., Pratta, M.A., Lark, M.W., Kim, Y.J., and Grodzinsky, A.J. 2004. Mechanisms and kinetics of glycosaminoglycan release following in vitro cartilage injury. Arthritis Rheum 50: 840±848. 85 Tew, S.R., Kwan, A.P., Hann, A., Thomson, B.M., and Archer, C.W. 2000. The reactions of articular cartilage to experimental wounding: role of apoptosis. Arthritis Rheum 43: 215±225. 86 Tew, S., Redman, S., Kwan, A., Walker, E., Khan, I., Dowthwaite, G., Thomson, B., and Archer, C.W. 2001. Differences in repair responses between immature and mature cartilage. Clin Orthop Relat Res 391: 142±152. 87 Redman, S.N., Dowthwaite, G.P., Thomson, B.M., and Archer, C.W. 2004. The cellular responses of articular cartilage to sharp and blunt trauma. Osteoarthritis Cartilage 12: 106±116. 88 Loening, A.M., James, I.E., Levenston, M.E., Badger, A.M., Frank, E.H., Kurz, B., Nuttall, M.E., Hung, H.H., Blake, S.M., Grodzinsky, A.J., et al. 2000. Injurious mechanical compression of bovine articular cartilage induces chondrocyte apoptosis. Arch Biochem Biophys 381: 205±212. 89 Quinn, T.M., Schmid, P., Hunziker, E.B., and Grodzinsky, A.J. 2002. Proteoglycan deposition around chondrocytes in agarose culture: construction of a physical and biological interface for mechanotransduction in cartilage. Biorheology 39: 27±37. 90 Stevens, A.L., Wishnok, J.S., Chai, D.H., Grodzinsky, A.J., and Tannenbaum, S.R. 2008. A sodium dodecyl sulfate-polyacrylamide gel electrophoresis-liquid chromatography tandem mass spectrometry analysis of bovine cartilage tissue response to mechanical compression injury and the inflammatory cytokines tumor necrosis factor alpha and interleukin-1beta. Arthritis Rheum 58: 489±500.

6

Understanding osteoarthritis and other cartilage diseases T . A I G N E R , Medical Center Coburg, Germany, N . S C H M I T Z , University of Leipzig, Germany È D E R , University of Erlangen-Nurnberg, Germany and S . S O

Abstract: The most relevant pathology of the articular cartilage is osteoarthritis (OA), i.e. degenerative joint cartilage destruction. Other important conditions are osteochondrosis dissecans, a focal subchondral bone destructive process leading to focal cartilage defects, as well as crystallopathies, which affect the joints in particular. Inflammatory conditions originating mostly from the synovial membrane as well as developmental malformations (chondrodysplasias) are not discussed. Besides the considerations of pathogenetic events and the description of pathological changes taking place during the disease process, a major emphasis will be put on the outline and discussion of grading systems of cartilage degeneration and repair. Key words: cartilage, osteoarthritis, osteochondrosis dissecans, grading, staging, typing, crystallopathies.

6.1

Introduction

The major relevant pathology of articular cartilage is degenerative joint disease, òsteoarthritis' (OA), which deals with all various forms of joint cartilage destruction. OA is in most cases primary, i.e. there are no clear reasons for its development and progression. Less frequently, OA is secondary due to inflammatory and endocrine conditions. However, (inflammatory) rheumatoid diseases are not within the scope of this chapter: they are primarily involving the synovial membrane (synovitis) and potentially bone and only secondarily involve the articular cartilage, largely similar to osteoarthritic cartilage degeneration. One important initiating event ± which might also be considered to lead to secondary OA ± is repetitive or focal trauma. Inflammatory events directly related to articular cartilage do not exist. Thus, for example, purulent arthritis, a rather rare condition, which can, however, have dramatic destructive consequences to the joints, is thought to result from septic granulocytic inflammation within the joint space. An important condition ± in particular as a potential target for repair ± is osteochondrosis dissecans, a focal subchondral bone destructive process leading to focal cartilage defects. Crystallopathies represent interesting conditions

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special to joints and the articular cartilage within the body. In contrast to most other tissues, there are no known neoplasias originating from articular cartilage. Besides considerations of pathogenetic events and the description of pathological changes taking place during the disease process, a major emphasis will be put on the outline and discussion of grading systems of cartilage degeneration and repair.

6.2

The normal joint

Joints are highly specialized organs (Fig. 6.1) that allow repetitive pain-free and largely frictionless movements. These properties are provided by the articular cartilage and its extracellular matrix, which under physiological conditions is capable of sustaining high cyclic loading. Articular cartilage covers the joint surfaces and is mainly responsible for the unique biomechanical properties of the joints. Joints are, however, complex composites of different types of connective tissue including (subchondral) bone, cartilage surfaces, ligaments and the joint capsule. Thus, the capsule, together with the ligaments, is extremely important for the mechanical stability of the joint as a whole. If malaligned, the cartilage is loaded abnormally and degenerates rather dramatically, as seen in misalignment syndromes of the joints (e.g. genu valgum et varum). All the different joint tissues together provide their own functional capacities in order to allow the correct functioning of the joint. The articular cartilage is a highly specialized and uniquely designed biomaterial that forms the smooth, gliding surface of the diarthrodial joints. It

6.1 Schematic representation of the main structures of a healthy (left side) and degenerated (right side) joint in osteoarthritis: in particular the articular cartilage is lost or severely thinned, the (subchondral) bone is sclerotic, the joint capsule thickened and the synovial membrane activated.

Understanding osteoarthritis and other cartilage diseases

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is largely an avascular, aneural and alymphatic matrix, which is synthesized by sparsely distributed resident cells ± the chondrocytes. The major constituents of the articular cartilage matrix are collagens, proteoglycans as well as the very heterogeneous group of non-collagenous non-proteoglycaneous proteins.1 In fact, most of the physiological (wet) weight of articular cartilage comes from water bound to the proteoglycans (namely aggrecan). The synovial capsule and in particular the synovial membrane (i.e. the synovial lining cell layer) represent important portions of the joint as an organ. As already mentioned, it is the capsule together with the ligaments which provide the mechanical stability of the joint and determine the flexibility of the possible movement. Overflexibility (e.g. after traumatic ligament rupture) clearly increases the risk for joint degeneration with time.2 The synovial membrane with its metabolically highly active surface cells (synoviocytes) plays a crucial role in nourishing the chondrocytes as well as removing metabolites and (matrix) degradation products form the synovial space. The synoviocytes maintain the basic metabolic homeostasis of the joints. Furthermore, the synoviocytes produce large amounts of hyaluronic acid, which provides the joint surfaces with its gliding capacity.

6.3

Major cartilage pathology and pathobiology

Most prominent conditions of articular joint cartilage are OA, whether primary or secondary to trauma, inflammation, etc., osteochondrosis dissecans and crystal deposition disease. Exceptionally, chondrodysplasia in particular of the hips occurs (for review, see Daneshpouy et al., 20023), which at least in part leads to early OA development.4 Many of them are, however, lethal and/or lead to severe malformation of the body.

6.3.1

The pathology and pathobiology of OA

Osteoarthritis, the degeneration of the joints, is the most common disabling condition in the Western world. Clinically, degeneration affects mostly the large weight-bearing joints of the legs (i.e. hips and knees), but can in principle affect any joint of the body including, notably, the finger joints. Osteoarthritis is not a single disease entity, but represents a disease group with rather different underlying pathophysiological mechanisms. In this respect, primary osteoarthritis has to be distinguished from secondary forms of the disease, which are due to traumatic events, endocrine or metabolic disorders, etc. Clinically, pain and loss of joint functioning are the major issues leading to a significantly reduced quality of life for patients suffering from the disease. Primary OA of the large weight-bearing joints is generally the result of an imbalance between applied mechanical stress and the physicochemical ability of the articular cartilage to resist this stress. In the end, osteoarthritis results from

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the destruction and failure of the extracellular matrix, the functional element of articular cartilage. However, osteoarthritis is a disease of the joint as an organ system (Fig. 6.1), not only of the articular cartilage. This includes all connective tissues within and around the joints as well as the respective musculature, the nervous system and even portions of the body more remote from the joint site, such as the central nervous system. The latter is particularly important for the symptomatic aspects of the disease and in fact the innervation and the processing are important to pain, the major symptom of the disease process. In terms of joint tissues, clearly the synovial capsule including the synovial membrane plays a very important role in the scenario of joint functioning and tissue maintenance. Overall, recognition that the joint does not only consist of articular cartilage, but also a number of adjacent tissues is not only important for the understanding of joint physiology, but also joint pathology. All these tissues are more or less affected by degenerative changes or their consequences (Fig. 6.1). In conventional radiology, in particular changes to the bone and (indirectly via the loss of joint space) to articular cartilage are visible (Fig. 6.2a). This, however, changed dramatically with the introduction of magnetic resonance imaging (MRI) into joint imaging.5,6

6.2 (a) Radiographic appearance of hip osteoarthritis displaying distorted joint architecture, loss of joint space as well as osteophyte formation in the joint margins. (b) Arthroscopic picture of a cartilage defect of the femoral condyle within the knee joint (courtesy Dr Eger, Rummelsberg). (c±d) Macroscopic appearance of femoral condyles of the knee: normal (c) and severely damaged (d).


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The cartilage matrix Macroscopically, hyaline articular cartilage is a rather unruffled white to yellowish overlay coating the joint surface (Fig. 6.2c). The synovial fluid makes it appear sloppy and provides its gliding properties. Microscopically, hyaline cartilage consists of evenly stained (`hyaline') proteoglycan-rich ground substance with the cartilage cells (`chondrocytes') lying sparsely in between. The cells represent less than 5% of the total volume of articular cartilage, but are of obvious importance for the maintenance of the tissue. Chondrocytes are surrounded in most parts by a specialized pericellular matrix forming a biomechanical and biochemical interface in between the rigid interterritorial matrix and the cells. The mechanical properties of articular cartilage largely depend on the biochemical composition of the extensive interterritorial (extracellular) cartilage matrix. Macroscopically, osteoarthritic cartilage is often yellowish or brownish and is typically soft. The surface shows roughening in the early stages and overt fibrillation and matrix loss in the later stages until the eburnated subchondral bone plate is visible (Fig. 6.2b,d). These changes are visualized in more detail on the histological level (Fig. 6.3b,d,e; a is normal articular cartilage for comparison). Besides the total destruction of matrix areas, the degradation of matrix molecules also plays an important role preceding and driving the final loss of the respective matrix areas (Fig. 6.3d,e: loss of toluidine blue staining reflecting the loss of proteoglycans in damaged cartilage areas). Apart from the degradation of molecular components, destabilization of supramolecular structures also takes place. For example, destabilization of the collagen network results in microscopically and finally macroscopically visible matrix destruction. Both mechanical wear and enzymatic degradation appear to play a pivotal role during the disease process. Together, these result in the destruction of cartilage matrix on the molecular (e.g. proteoglycan depletion) and the macromolecular (e.g. network loosening), the microscopic (e.g. fissuring) and the macroscopic (e.g. cartilage tear) levels. The destruction of articular cartilage and the loss of its biomechanical function are largely due to the destruction and loss of the interterritorial cartilage matrix, which result from an imbalance between degradation and de novo synthesis of matrix components on the molecular and supramolecular level in spite of the compensatory attempts of the chondrocytes. The cartilage cells (chondrocytes) Despite the importance of the extracellular matrix for the functioning of articular cartilage, the cells are not `functionless' in connective tissues, as they are the only viable players within the tissue. Thus, they are centrally responsible for the balanced turnover of the extracellular matrix, which is necessary for maintenance of the integrity of the extracellular cartilage. During the

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6.3 Conventional histology shows fibrillation and matrix loss in osteoarthritic cartilage (b) compared to the normal (a). In severely damaged areas nearly all articular cartilage is destructed (e). Also a moderate (d) to severe (e) loss of proteoglycans is found as visualized by toluidine blue staining (d,e). Besides changes in articular cartilage, changes in the subchondral bone are also prominent, namely thickening of the bone trabeculae (f, osteoarthritic; c, normal).

osteoarthritic disease process, the cellular reaction patterns are altered. At first sight these are rather pleomorphic, but can be basically summarized in three categories. First, the chondrocytes can undergo cell death or they can proliferate to compensate for cell loss or to increase their synthetic activity. Secondly, chondrocytes activate or deactivate their synthetic-anabolic activity by increasing or decreasing anabolic gene expression. Lastly, chondrocytes undergo phenotypic modulation implicating an overall severely altered gene expression profile of the cells in the diseased tissue. In addition, osteoarthritic chondrocytes are heterogeneous and nearly all observed cellular changes are region- and zonespecific, and also dependent on the degradation stage. One straightforward explanation for osteoarthritic cartilage degeneration would be a mere loss of viable cells at the beginning and during the disease process (for review, see Aigner and co-workers7,8). However, this has only limited impact on the pathology of early osteoarthritis or ageing of human articular


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cartilage.9 The only zone in which a large number of empty lacunae were found was the calcified cartilage layer. The continuous progression of cartilage calcification in osteoarthritic cartilage might explain, at least partly, the increasing number of empty lacunae reported, particularly in high grade lesions.10,11 One striking phenomenon in many OA cartilages is the focal proliferation of chondrocytes, forming so-called chondrocyte `clusters': this clearly reflects the increased proliferative activity within the diseased tissue and might represent an attempt of cells to compensate for an increased need of cellular activity or even cell death (similar to other tissues). Overall, however this increased proliferative activity might not be of much help to maintain tissue homeostasis as these cell clusters are often anabolically inactive12 and in fact, might represent rather further empty holes destabilizing the cartilage matrix. Also, increased proliferation might lead to (focal) shortening of the telomeres of the cells which might itself be problematic for cell integrity.13 The synovial membrane, the joint capsule and inflammation OA research traditionally concentrates on the understanding of events within the degenerated articular cartilage as the tissue in which the initiating events presumably take place. Synovial changes are generally interpreted as largely secondary to the degeneration of the articular cartilage14±16 and not pathogenetically involved to a relevant extent in the disease process. However, the synovial capsule and, in particular, the synovial lining cells represent an important portion of the joint as an organ. Also, thickening of the collagen plate within the joint capsule, a typical feature in many osteoarthritic patients at least in the late stages reduces significantly the movement properties of their joints. Thus, capsular fibrosis is centrally responsible for joint stiffening, which is, after pain, the second biggest symptomatic issue of osteoarthritic joint degeneration. Overall, four types of synovial reaction pattern can be distinguished:17 presumably, in most cases the earliest event is hyperplastic synoviopathy, which is mainly characterized by synovial hyperplasia (i.e. villus formation) as well as proliferation18,19 and activation of the synovial surface cells (i.e. synovial lining layer, synoviocytes). At the end, detritus-rich synovitis is often observed, which is characterized by a lot of cartilage and bone debris as well as fibrinous exudate and some granulating inflammation. Few granulocytes might be observed as well as a giant cell rich foreign body reaction. This is at this stage also mostly combined with fibrous thickening of the joint capsule, a feature which in some cases also dominates in earlier stages of the disease and is then called fibrous OA synoviopathy (this might include hypertrophic changes as well). A forth pattern of synovial reaction in OA is inflammatory OA synoviopathy, which is characterized by significant lymphoplasmacellular infiltrates resembling that one found in rheumatoid conditions (but less pronounced). Spotty lymphocyte aggregates might also occur in other variants. Granulocytes are not part of the

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spectrum of OA synoviopathy (except minor infiltrates already mentioned in detritus-rich synoviopathy). The subchondral bone Another important tissue often neglected in osteoarthritis research is the subchondral bone,20 although it is unclear yet whether pathological changes within the subchondral bone tissue (e.g. sclerosis) can precede changes in the articular cartilage (e.g. bone mass as risk factor of osteoarthritis21) or subchondral bone changes are secondary adaptation processes following changes in the biomechanical properties of the cartilage.22 Significant changes in terms of increased thickness of the subchondral bone plate as well as underlying trabecules are already apparent in the early stages of the disease (Fig. 6.3c,f). Thus, active new bone formation is found at multiple foci in early- to mid-stage patients.23 In later stages, severe bone remodelling processes take place, in particular in areas of advanced destruction of the overlying articular cartilage. Apart from extensive bone sclerosis, significant aseptic bone necrosis is a common feature of advanced osteoarthritic joint degeneration. In areas of total cartilage destruction (i.e. eburnated bone plate) synovial fluid gets access to the bone marrow and induces fibrocytic and even chondrometaplastic changes of mesenchymal precursor cells. This leads to the characteristic `cartilage-nodules' or `tufts', which are frequently found in these areas in late stage disease. At least in moderate to advanced lesions, the changes in the subchondral bone represent one tissue responsible for the osteoarthritic joint pain.24,25

6.3.2

Post-traumatic cartilage lesions ± intra-articular bleeding

Trauma, in particular microtrauma, is thought to be one core causing event for the development of osteoarthritis. Obviously, people undertaking sporting activity and also heavy occupational loading are at high risk of developing general joint cartilage degeneration. Major traumatic events, such as intraarticular fracturing and share traumata leading to cartilage flakes, are major issues both in terms of symptoms as well as long-term outcome. They might require direct surgical or arthroscopical intervention. Also post-traumatic bleeding into the joint cavity might negatively influence cartilage and chondrocyte26,27 integrity, though this clearly depends on the extent and its duration. Such events are particularly a problem if there exists an increased bleeding susceptibility such as in haemophilic disease. In this condition, significant joint damage including the articular cartilage occurs, though part of it is thought to be mediated through synovial changes rather than a direct effect on the articular cartilage and the cells.28 Nevertheless, blood has been shown to damage chondrocytes in vitro29 and in vivo.30


6.3.3

163

Osteochondrosis dissecans

In general, osteochondrosis dissecans is a condition in which ± as indicated by the name ± an osteochondral flake is torn off the joint surface. This, in principle, can occur in any joint, but is most frequently encountered in the medial femoral condyle. Pathogenetically, the necrosis of the subchondral bone compartment is the core event, which then leads to the weakening of the subchondral bone plate strength with subsequent dissociation from the remaining bone. The overlying articular cartilage might be rather vital and largely intact or may show alternatively moderate to severe degenerative alterations. As the definition, the subchondral bone part of an osteochondral flake, however, is fully necrotic. As a consequence of the destruction of part of the articular surface, often a more extended or generalized joint destruction occurs and thus, osteochondrosis dissecans is a classical cause of secondary osteoarthritis. The underlying condition of osteochondrosis dissecans is so far largely unknown: one general speculation is an articular trauma leading to a vascular occlusion and thus subchondral bone necrosis. However, there might be many other factors involved, including a familial predisposition as described at least for some cases.31 Whatever osteochondrosis dissecans finally is, it is a condition affecting primarily the bone and only on a second level the articular cartilage and the joint as such. This, however, occurs in a dramatic way in most cases.

6.3.4

Crystal deposition disease

There are two major forms of crystal deposition disease in the articulating joints: deposition of urate crystals (i.e. gout) and the deposition of pyrophosphate crystals (pseudogout, chondrocalcinosis). Both can cause significant symptomatic disease, but gout is clearly usually symptomatic and pseudogout in most cases a clinically silent process mostly associated with (osteoarthritic) cartilage degeneration. Gout Gout, a severely symptomatic disease mainly affecting the joints, has been known for a long time. It mostly affects older men and is characterized mainly by very painful synovitis or inflammation in the periarticular soft tissues, though any organ except the brain might be involved in gouty `tophus' formation. Gout occurs in repetitive phases (gouty àttacks') and often first affects the feet (often the first metatarsophalangeal joint). Typically, the urate levels are increased in the serum. The precipitation of urate crystals in the soft tissue is often following alcoholic excess or calory-rich food intake. However, the exact reason of the urate deposition, e.g. preferentially in certain joints and less often in others is not yet known. Histologically, gouty tophi are characterized by needle-shaped crystals showing a typical negative birefringence in polarized light microscopy

164


6.4 Urate crystal deposition (gout) is characterized by more or less extensive deposition of strongly birefringent crystalline material within an amorphous matrix (a) hematoxyline±eosine, (b) polarized light microscopy (wonderful crystalline structure from different area)). (c, d) calcium pyrophosphate crystallopathy (chondrocalcinosis) shows the deposition of hardly birefringent short needle-like crystalline material (c, hematoxylin±eosine; d, detail photographed with polarized light microscopy).

(Fig. 6.4b). Sodium urate crystals are embedded into an amorphous eosinophilic mass. The area of crystal deposition is surrounded by a more or less dense mononuclear (histiocytic) infiltrate with multi-nuclear giant cells of the foreign body type scattered in between. The strongly birefringent crystals also make this condition easily distinguishable from the small, hardly birefringent crystals present in chondrocalcinosis. Of note, during conventional histological processing most urate crystals are dissolved in ethanol and only the weakly eosinophilic small to medium sized areas surrounded by the histiocytic and giant cells remain, a feature diagnostic for this condition (but not to be confused with small foreign body granulomas after intra-articular injections). Though gout is a condition mostly affecting the peri-articular soft tissues, urate crystal deposits can be occasionally found also in articular cartilage. Epidemiological evidence suggests also that osteoarthritic degenerative joint disease is one potential risk factor for gouty arthritis.32 Pseudogout In contrast to gout, pseudogout (chondrocalcinosis, calcium pyrophosphate dehydrate crystal deposition disease) primarily affects the articular cartilage, in which radiologically calcium deposition can be detected in plain X-rays. Besides the articular cartilage also the synovial membrane/capsule, ligaments, tendons,


165

menisci and within the hip joint the limbi are affected. Usually, the calcium pyrophosphate crystals are deposited without significant (inflammatory) reaction by the surrounding tissue, but in cases of symptomatic pseudogout arthritis a mononuclear infiltrate with accompanying giant cell reaction similar to gout can be observed around the deposits. The incidence of chondrocalcinosis in the elderly is high, reaching 25% in 80-year-olds (for review, see Wise33). In polarized light microscopy the calcium pyrophosphate crystals are very small, hardly birefringent needle-like structures that are difficult to see (Fig. 6.4d). In contrast to the urate crystals, calcium pyrophosphate crystals are less water soluble and, thus, remain mostly within the tissue during histological processing. Overall, the cause and the relevance of chondrocalcinosis remain unclear: clearly, it is well correlated with the degeneration of the articular cartilage and it is probably related to a metabolic disbalance within the cells and the tissue, most likely the articular cartilage and the chondrocytes.

6.4

In vivo cartilage repair

At the margins of joints, in particular in osteoarthritic joint disease, frequently (osteo)cartilaginous outgrowths appear ((chondro-)osteophytes). They are best considered as a process of secondary chondroneogenesis in the adult.34 Osteophytes derive from mesenchymal precursor cells within periosteal or synovial tissue and often merge with or overgrow the original articular cartilage.35,36 Thus, in this process, mesenchymal precursor cells differentiate into chondrocytes. A similar, but less structured process is observed in the areas of the eburnated bone, in which the articular cartilage is completely torn off. Here, mesenchymal multipotential stem cells of the bone marrow undergo also chondrogenic differentiation: metaplastic cartilage in forms of nodules or `tufts' is found either within the bone marrow or at the naked bone surface.37 Osteophytes could be considered as endogenous repair attempts in degenerating joints and might be a physiological response to mechanical overloading by increasing the articulating joint surface. Even if their supportive effect within the joints is doubtful, their chondrogenic potential is of interest, especially having exogenous (therapeutic) repair strategies in mind. Central for the basic understanding of osteophytic tissue is the analysis of the developmental steps during osteophyte formation. Thus, although it is clear that osteophyte development is a continuous process and many osteophytes show different stages in various portions at the same time, one can define basic steps based on the cellular phenotype and the matrix composition of the predominating tissue38 (Table 6.1 and Fig. 6.5). Initially, mesenchymal precursor cells derived either from periosteum or synovium initiate chondrogenic differentiation (stages I and II). This results in fibrocartilage composed of both fibrous and cartilaginous matrix components. In early osteophytes, endochondral ossification is initiated. The deepest cell layer becomes hypertrophic and

166


Table 6.1 Staging of osteophyte development according to Gelse et al.38 Stage 0 (normal)

Normal periosteum

Stage I

· Slight thickening of the periosteum · Incipient formation of fibrocartilage (some round cells, some metachromatic tissue staining of the extracellular matrix) · No/slight active bone formation Molecular markers: · Focal collagen type II expression · No collagen type X

Stage II

· Pronounced thickening of the periosteal layers · Well-established formation of fibrocartilage (many round cells, strong metachromatic tissue staining of the extracellular matrix) · Some/moderate bone formation Molecular markers: · Distinct collagen type II expression · No collagen type X

Stage III

· Pronounced thickening of the periosteal layers · Well-established formation of fibrocartilage (many round cells, strong metachromatic tissue staining of the extracellular matrix, formation of lacunae) · Strong active bone formation Molecular markers: · Distinct collagen type II expression · Collagen type X expression in basal areas

Stage IV

· Significant thickening of the periosteal layer · Apparent formation of fibrocartilage with partial hyalinization of the extracellular matrix (chondrocyte-like cells in lacunae, strong metachromatic tissue staining of the extracellular matrix) · Some active bone formation Molecular markers: · Ubiquitous presence of collagen type II · Collagen type X in basal areas · Collagen type VI within the extracellular matrix accentuated pericellularly

resembles the lowest cells found in the growth plate (stage III).39 Mature osteophytes are characterized by the predominance of a hyaline cartilage-like extracellular matrix (stage IV). At a first glance, mature osteophytes can, macroscopically and histologically, easily be mistaken for original articular cartilage. Indeed, this misconception reflects to some degree the fact that


167

6.5 Osteophyte development can be subdivided into five stages with different structural organization although many osteophytes show different stages simultaneously in different areas. Stage I (early chondrophytes) shows first chondrocytic differentiation of previously undifferentiated mesenchymal precursor cells (b,g,k). Stage II (chondrophytes) shows extensive areas of newly formed cartilage, but no (endochondral) bone formation is observed (c,h,k). Stage III (early osteophytes) shows an arrangement as the fetal growth plate cartilage (d,i,k) whereas stage IV (mature osteophytes) shows a structure most resembling hyaline articular cartilage physiologically covering the joint surfaces (e,j,k). Normal periosteum is shown in a and f. (a±e, hematoxylin eosin staining; f±j, toluidine blue staining).

chondrocytic cells in osteophytes are able to construct an extracellular matrix containing all the typical components of hyaline articular cartilage such as collagen types II, IX and XI as well as aggrecan.34,38 The zonal distribution also resembles that found in adult articular cartilage with collagen type VI concentrated in the pericellular matrix.40 Although hyaline zones in osteophytes resemble articular cartilage in terms of structural composition, there are, never-

168


theless, certain differences such as a more random cellular arrangement, the lack of a distinct tidemark, and a missing linear subchondral bone plate. Furthermore, the proper alignment of the described matrix components, which is obligatory for the cartilage to be able to resist high mechanical forces, has not yet been investigated at the ultrastructural level.

6.5

Grading/scoring systems for cartilage degeneration

Overall, the classification of osteoarthritic cartilage degeneration is rather complex, as all patients present with at least to some extent different histories, symptoms and morphological changes. Common to all of them is some sort of structural joint (cartilage) damage, pain and limitation in joint movement. Fortunately, for most instances an exact classification of the destructive process is of limited clinical and scientific use. Obviously, many other tissues than the articular cartilage are involved in this process, but traditionally, the cartilage has been used in order to score OA severity (at least as long as structural changes are evaluated: in general, the process of joint destruction can be always evaluated for the pathogenesis (`typing'), for its extent (`staging') and for the degree of the most extensive focal damage (`grading'). `Typing' is mostly related to `primary', i.e. idiopathic, and `secondary', i.e. `caused by . . .' OA. Primary OA is most common. Whereas the addition `primary' suggest it to be without any obvious cause, still minor pre-existing conditions also exist in this condition (i.e. `pre-conditions' or `risk factors'). The major causes leading to secondary osteoarthritic joint degeneration are listed in Table 6.2. `Grading' and `staging' have been much more under debate, also Table 6.2 Typing of joint destruction Primary

No causative reason known

Secondary

· Rheumatic disease · Overload causing excessive wear (work, sport, varus or valgus deformity) · Instability (e.g. meniscus lesions) · Trauma · Intra-articular infections · Articular gout · Psoriatic arthritis · Bone infarciation · Endocrine disorders (e.g. hyperparathyroidism) · Neuropathy (e.g. Charcot's joint) · Paget's disease · Haemophilia


169

regarding the basic meaning of both words. This author suggests the use of `grading' for the evaluation of histological changes at one site of analysis, whereas `staging' should refer to the overall disease process (in analogy to `grading' and `staging' in tumour pathology). Both represent an attempt of scoring processes, which is suggested to be used as general term for such activities. The grading system most often used (partly in minor modifications) is that proposed by Mankin and coworkers in 19719 (Table 6.3; Fig. 6.6). Despite repetitive criticisms that the Mankin score shows a high interindividual variability,41 this might be related to the training status of the people doing the scoring. However, clearly some of the subcategories of the MankinÂs score do not belong to primary cartilage degeneration, but describe features observed in secondary cartilage formation (i.e. osteophyte formation: see Table 6.3) and should be excluded in future scoring attempts. A staging system largely used in Germany, but still in principle up to date is that of Otte42 (Table 6.4; Fig. 6.6). Whereas Mankin addresses the piece of cartilage under the microscope, Otte looks at the whole joint surface mostly macroscopically (but if needed the worst lesion can be evaluated histologically). At the site of the highest cartilage damage, grading according to Mankin and staging according to Otte are closely Table 6.3 Grading of osteoarthritis according to Mankin et al.9 Feature

Score

Histological feature

Cartilage structure

0 1 2 3 4 5

Normal Superficial fibrillation Pannus and superficial fibrillation* Fissures to the middle zone Fissures to the deep zone Fissures to the calcified zone

Chondrocytes

0 1 2 3

Normal Diffuse hypercellularity Cell clusters Hypocellularity

Safranin-O staining

0 1 2 3 4 5

Normal Slight reduction Moderate reduction Severe reduction No staining Total disorganization*

Tidemark

0 1

Intact Tidemark penetrated by vessels**

* Should be removed (relates to osteophyte formation). ** Might best be supplemented with: òr duplicated tidemark').

170


6.6 The grading system according to Mankin et al.9 compared with the staging according to Otte.42

Table 6.4 Staging of joint destruction according to Otte42 Grade

Morphology

0 I II

Normal Superficial fibrillation, no cartilage loss Cartilage lesions (without full thickness defects) (deep fibrillation, fissures to middle zone and/or partial cartilage matrix loss) Cartilage lesions (without full thickness defects) (fissures to deep zone and partial cartilage matrix loss) Complete cartilage loss (at least focally)

III IV

correlated. Clearly, Otte is too rough for scientific purposes and a new staging system has recently proposed by Pritzker and colleagues43 (Table 6.5), but in clinical terms this provides in most instances a rough suitable classification of the condition without adding and requiring too much unneeded information. Doubtless, along with new scientific insights and with more extensive and specified medical options we will need more elaborated `grading' and `scoring' systems and this will be a major task in the near future.

6.6

Grading/scoring of cartilage repair

Many approaches (for review see Nesic et al.44) have been followed in order to promote external cartilage repair either by implanting autologous chondrocytes or chondrocyte precursor cells. Whatever therapeutic method is used, the


171

Table 6.5 Grading of osteoarthritis according to Pritzker et al.43 Grade

Histological properties

0

Matrix: surface intact (normal architecture) Cells: intact, appropiate orientation

1

Matrix: superficial zone intact, oedema and/or superficial fibrillation (abrasion), focal superficial matrix condensation Cells: cell death, proliferation (cluster formation), hypertrophy

2

As above: Matrix: discontinuity at superficial zone (deep fibrillation) loss of PG-staining in upper third of cartilage focal perichondral increased PG-stain in middle zone disorientation of chondron collums

3

As above: Matrix: vertical fissures into middle zone and branched fissures loss of PG-staining into lower two-thirds of cartilage new collagen formation Cells: cell death, regeneration, hypertrophy in cartilage domains adjacent to fissures

4

As above: Cartilage matrix loss with delamination of superficial zone Excavation with matrix loss from superficial to middle zone formation of cysts in the middle layer

5

Complete matrix loss with denudation of the sclerotic subchondral bone or fibrocartilage microfracture with repair limited to bone surface

6

Bone remodelling (more than osteophyte formation only) with microfracture, fibrocartilage and osseous repair above the previous surface

important issue, in terms of outcome measurement, is not only clinical symptom evaluation but also the question what tissue is formed in terms of composition and function. Both biochemical composition and biomechanical function are closely related as the newly formed tissue has to fulfil the biomechanical needs of a specific connective tissue, i.e. articular cartilage, which it has to substitute. Thus, although the final functional outcome remains the main criterion for the success of a procedure, the tissue type formed appears to be the major prerequisite for the final success. Basically, three types or levels of repair tissue (besides the complete absence of repair tissue at all) can be distinguished:45 fibrous tissue, fibrocartilage and hyaline-like repair cartilage of varying resemblance to original articular cartilage.46 All these types of tissue resemble the different stages of osteophyte development described above,38 making this in vivo phenomenon so interesting for cartilage repair biology.

172


Fibrous repair tissue can be regarded as a failed repair attempt as it is unlikely to represent a successful long-term restoration of the joint function. Cells produce a matrix with a poor content of glycosaminoglycans and with abundance of type I collagen.47,48 Type II collagen is not present or represents only a very minor fraction. Since this repair tissue lacks the unique properties of articular cartilage, which are required for a successful participation in the articulating processes of joints, fibrous repair tissue fails when exposed to mechanical load.46 Fibrocartilage is the most common form of repair tissue achieved. In terms of structure and composition, fibrocartilage has an intermediate position in between fibrous and hyaline-like cartilaginous tissue. The content of glycosaminoglycans does not reach the abundance found in hyaline articular cartilage, but is increased compared with fibrous tissue. The matrix of fibrocartilage is composed of both type I collagen, typical for fibrous tissue, and type II collagen, typical for hyaline cartilage.46,49,50 Thus, although at first sight the joint surface and the macroscopic integrity of the cartilage appear to be largely restored, fibrocartilage does not possess the biomechanical properties of articular cartilage, which are needed for tolerating long-term constant mechanical loading and movement. Therefore, therapeutic attempts leading to fibrocartilagenous repair tissue are also prone to suffer from long-term deterioration with fibrillation, swelling, loss of cells and finally the loss of the repair tissue itself.47±49 Under certain conditions, a rather complete chondrogenic differentiation and remodelling process was reported that generated repair cartilage which shared great similarity with normal articular cartilage.50±52 Macroscopically, glossy surfaces of the restored defects suggested highly effective repair yielding the restoration of tissue resembling healthy articular cartilage. Histochemical analysis confirmed an abundance of water-binding glycosaminoglycans within the extracellular matrix of this repair tissue. The cells displayed a spherical shape typical for chondrocytes and were embedded in lacunar spaces. The extracellular matrix of these repair tissues was shown to contain types II, IX and XI collagens. The absence of type I collagen indicated a rather complete transformation or differentiation of the implanted cells into a functional chondrocytic phenotype forming hyaline cartilage-like repair tissue.47,51,53,54 Despite the similarities of this repair tissue to normal articular cartilage, which are apparent at first glance, there might still be subtle differences: in normal articular cartilage, the structural organization of the collagen network has a typical zonal pattern. In repair cartilage the fibres appear to be more randomly distributed and the cellular density and cellular arrangement often differ significantly from those of normal articular cartilage.46,55±57 Additionally, repair cartilage often lacks a clear tidemark which separates the upper portions of articular cartilage from the underlying calcified cartilage and bone.52,54 Consensus criteria for the evaluation of tissue repair outcome (on the histopathological level) were issued by the ICRS (International Cartilage Repair


173

Table 6.6 The ICRS visual histological assessment score for assessing repair cartilage quality.45 The observer should evaluate all criteria separately without adding them up at the end I. Surface smooth/continuous discontinuities/irregularities

3 0

II. Matrix hyaline mixture: hyaline/fibrocartilage fibrocartilage fibrous tissue

3 2 1 0

III. Cell distribution columnar mixed/columnar-clusters clusters individual cells/disorganized

3 2 1 0

IV. Cell population viability predominantly viable partially viable

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