Handbook of Tissue Engineering Scaffolds: Volume One [1] 0081025637, 9780081025635

Table of contents :
Cover
Handbook of Tissue Engineering Scaffolds: Volume One
Copyright
List of contributors
Foreword
Preface
Acknowledgment
Part One: An introduction to tissue engineering scaffolds
1 - Introduction to tissue engineering scaffolds
1.1 Introduction
1.1.1 Scaffolding approaches in tissue engineering
1.1.2 Fabrication techniques for tissue engineering application
References
Further reading
2 - The role of scaffolds in tissue engineering
2.1 Introduction
2.1.1 Tissue engineering and scaffolds
2.1.2 Metal-based scaffolds
2.1.3 Ceramic-based scaffolds
2.1.4 Polymer-based scaffolds
2.1.5 Composite-based scaffolds
2.2 Cell–ECM interaction and RGD nanospacing
2.2.1 RGD nanospacing in 2D substrates with different stiffness
2.2.23 D substrates
2.3 Mechanotransduction
2.4 Surface topography–mediated stem cell fate
2.5 Control of cell migration and cancer invasion
2.6 Scaffold for gene delivery
2.7 Scaffold for multimodal drug delivery
2.8 Scaffolds for bone tumor destruction
2.9 Scaffolds for cell separations
2.10 Future direction and conclusions
References
3 - Scaffolds mimicking the native structure of tissues
3.1 Introduction
3.2 Characterization of native tissues
3.2.1 Common chemical components in ECM
3.2.2 Specific characteristics in ECM
3.2.3 Mechanical properties - hard versus soft tissues
3.2.4 Tissue with stratified epithelium (skin, lung, cornea, conjunctiva)
3.2.5 Zonal, layer-specific tissues
3.2.6 Vascularized tissues
3.3 Scaffold designs to mimic the native structure of tissues
3.3.1 Scaffolds for soft tissue
3.3.2 Tissue models with epithelium (coculture + multilayer scaffolds)
3.3.3 Scaffolds with zonal, layered structure
3.3.4 Scaffolds to promote vascularization
3.3.5 Scaffolds from decellularized tissues
3.4 Summary
References
4 - Computational design of tissue engineering scaffolds
4.1 Introduction
4.2 Preprocessing: design of the scaffold
4.2.1 Scaffold structural properties
4.2.2 Mechanical properties
4.2.3 Modeling scaffold degradability
4.2.4 Mass transport
4.3 The fabrication process
4.3.1 Shape fidelity in function of the fabrication process
4.3.2 Biocompatibility of the fabrication process conditions
4.3.3 Biological functionality after the fabrication process
4.4 Postprocessing: bioreactor culture
4.4.1 Incorporating the neotissue domain
4.4.2 Multiphysics models for scaffolds in bioreactors
4.5 Discussion
4.5.1 Multiparametric optimization
4.5.2 Future prospects
Acknowledgments
References
5 - Research progress of scaffold materials
5.1 Introduction
5.1.1 Types of biomaterials
5.1.2 Synthetic biomaterials
5.1.3 Natural biomaterials
5.2 Biomaterials for tissue engineering applications
5.2.1 Biomaterials for hard tissue engineering
5.2.2 Biomaterials for soft tissue engineering
5.3 Research development of tissue engineering biomaterials
5.3.1 First-generation biomaterials
5.3.2 Second-generation biomaterials
5.3.3 Third-generation biomaterials
5.3.4 Fourth-generation biomaterials
5.4 Recent techniques in tissue engineering fabrication
5.4.1 Bioprinting: bioink materials for tissue engineering scaffolds
5.5 State-of-the-art and future perspectives
5.6 Conclusions
List of abbreviations
Acknowledgments
References
Further reading
6 - Fabrication techniques of tissue engineering scaffolds
6.1 Introduction
6.2 Scaffold fabrication techniques
6.2.1 Porous scaffolds
6.2.1.1 Solvent casting and porogen leaching
6.2.1.2 Phase separation
6.2.1.3 Gas foaming
6.2.1.4 Sintering
6.2.1.5 Electrospinning
6.2.1.6 Self-assembly
6.2.1.7 Hybrid scaffolds
6.2.2 Additive manufacturing
6.2.2.1 Powder-bed three-dimensional printing
6.2.2.2 Selective laser sintering
6.2.2.3 Fused deposition modeling
6.2.2.4 Stereolithography
6.2.3 Hydrogels
6.2.4 Tissue/organ decellularization
6.2.5 Tissue and organ bioprinting
6.3 Conclusions
Acknowledgments
References
7 - Scaffolds implanted: what is next?
7.1 Introduction
7.2 Host immune reaction against implanted scaffolds
7.2.1 Surgical procedure induces initial inflammation
7.2.2 Bacterial adhesion
7.2.3 Protein fouling further calls for immune response
7.2.4 Myriad of immune response following protein adsorption
7.3 Wound healing versus scar formation
7.3.1 Scar thickness and its effects
7.4 Recent understandings on immune cells activity against implants
7.4.1 Presence of Th17 helper T cells and recruitment of neutrophils
7.4.2 Macrophages and their polarization
7.5 Immunoengineering scaffolds
7.6 Biointegration of scaffolds with the host body
7.7 Biodegradable scaffolds
7.8 Conclusion and future perspectives
References
8 - Moving from clinical trials to clinical practice
8.1 Introduction
8.2 Clinical applications
8.2.1 Decellularized organs
8.2.1.1 Commercialization of organ decellularization
8.2.2 Clinical applications of scaffolds
8.2.2.1 Characteristics of scaffolds
8.2.2.2 Bone
8.2.2.3 Trachea
8.2.2.4 Cartilage
8.2.2.5 Nerve
8.2.2.6 Skin
8.2.2.7 Urethra
8.3 Conclusion and future research
References
9 - Tissue engineering scaffolds: future perspectives
9.1 Introduction
9.2 Scaffolding approaches for tissue engineering
9.2.13 D scaffolds
9.2.2 Hydrogel-based matrices
9.3 Concluding remarks and future perspectives
Acknowledgments
References
Part Two: Musculoskeletal tissue engineering scaffolds
10 - Scaffold for bone tissue engineering
10.1 Introduction
10.2 Bone structure and properties
10.3 Scaffolds for bone tissue engineering
10.3.1 Biological requirements of bone scaffolds
10.3.2 Structural features of bone scaffolds
10.3.3 Biomaterial composition of bone scaffolds
10.3.3.1 Bioactive ceramics and glasses in bone scaffolds
10.3.3.2 Natural and synthetic polymers and protein templates in bone scaffolds
10.3.3.3 Composites in bone scaffolds
10.3.3.4 Metallic bone scaffolds
10.3.4 Fabrication processes for bone scaffolds
10.3.4.1 Conventional technologies for bone scaffold fabrication
10.3.4.2 Additive manufacturing of bone tissue engineering scaffolds
10.4 FDA-approved bone scaffolds used in humans
10.5 Conclusion
References
11 - Scaffolds for cartilage tissue engineering
11.1 Introduction
11.1.1 Cartilage types and structure
11.1.1.1 Hyaline cartilage, fibrocartilage, and elastic cartilage structural differences
11.1.1.2 Zonal composition of articular cartilage
11.1.2 Clinical techniques
11.1.2.1 Microfracture
11.1.2.2 Autologous chondrocyte implantation
11.1.2.3 Matrix-assisted chondrocyte implantation
11.1.3 What is a scaffold?
11.2 Cartilage scaffolds
11.2.1 Natural materials
11.2.1.1 Collagen
11.2.1.2 Fibrin
11.2.1.3 Hyaluronan
11.2.1.4 Chitosan
11.2.1.5 Agarose and alginate
11.2.1.6 Silk
11.2.1.7 Native cartilage matrix
11.2.2 Synthetic materials
11.2.2.1 Polyglycolic acid
11.2.2.2 Polylactic acid
11.2.2.3 Polylactic-co-glycolic acid
11.2.2.4 Others
11.2.3 Composite scaffolds
11.2.3.1 Chondroinductive approaches
11.2.3.1.1 Growth factors
11.2.3.1.2 Chondroitin sulfate
11.2.3.2 Hybrid scaffolds
11.3 Osteochondral approach
11.3.1 Allografts
11.3.2 OC scaffold configurations
11.3.2.1 Single phase
11.3.2.2 Multiphase
11.3.2.3 Gradient
11.4 Future perspectives
11.5 Conclusions
Acknowledgments
References
12 - Scaffolds for skeletal muscle tissue engineering
12.1 Scaffolds for skeletal muscle engineering
12.1.1 Response of skeletal muscle to injury
12.2 Synthetic scaffolds
12.2.1 Nondegradable synthetic scaffolds
12.2.2 Biodegradable polymeric materials
12.2.3 Biologic scaffolds
12.2.4 Closing remarks
12.3 Cell types for skeletal muscle tissue engineering
12.4 Conclusions and future directions
References
13 - Scaffolds for tendon tissue engineering
13.1 Introduction
13.2 Biomaterial-­based therapies
13.2.1 Electrospinning (ES)
13.2.2 Imprinting
13.2.3 Hydrogels
13.2.4 Extruded microfibers
13.2.5 Lyophilized materials
13.3 Tissue graft–based therapies
13.3.1 Tissue graft fabrication
13.3.2 Tissue grafts from decellularized tendons
13.3.3 Tissue grafts from other tissues
13.4 Scaffold-­free tissue engineering by self-­assembly
13.5 Conclusion and future perspectives
List of abbreviations
Acknowledgments
References
14 - Scaffolds for ligament tissue engineering
14.1 Introduction
14.2 Anatomy, physiology, and function of ligament
14.2.1 Fiber bundle anatomy
14.2.2 Ligament and bone interface
14.2.3 Mechanical properties of the ligament
14.3 Conditions and injuries, diseases, and disorders of ligament tissue
14.4 Ligament healing
14.5 Scaffold design and fabrication techniques
14.6 Biomaterials available for ligament tissue engineering
14.6.1 Natural materials
14.6.2 Synthetic polymers
14.7 Properties of an ideal ligament tissue scaffold
14.8 Current technologies and strategies used in ligament tissue engineering
14.8.1 Biological grafts
14.8.2 Nondegradable grafts
14.8.3 Tissue-engineered biodegradable grafts
14.9 Future research in ligament tissue engineering
References
Further reading
15 - Scaffolds for regeneration of meniscus lesions
15.1 The knee meniscus: structure and function
15.2 Meniscus lesions: available therapeutic options
15.3 Tissue engineering for cartilage and meniscus regeneration
15.3.1 Cells and growth factors
15.3.2 Biopolymer 3D graft
15.3.3 Hydrogel 3D scaffolds and mixed approach
15.4 Conclusions
References
Part Three: Craniomaxillofacial tissue engineering scaffolds
16 - Scaffolds for mandibular reconstruction
16.1 Introduction
16.2 Clinical need of mandibular scaffolds
16.3 Elements of scaffold development
16.3.1 Cells
16.3.2 Growth factors
16.4 Mandibular scaffold options
16.4.1 Scaffolds for small mandibular defects
16.4.2 Scaffolds for critical mandibular defects
16.5 Future requirements in mandibular regeneration
References
17 - Scaffolds for maxillary sinus augmentation
17.1 Introduction
17.2 Maxillary sinus augmentation procedure
17.2.1 Overview of surgical techniques
17.2.2 Lateral window approach
17.2.3 Transalveolar approach
17.3 Scaffolding materials for the maxillary sinus augmentation
17.3.1 Bone grafts
17.3.2 Rigid scaffold
17.3.3 Space maintainers
17.3.4 Biologic agents
17.3.4.1 Bone morphogenetic proteins
17.3.4.2 Recombinant human platelet–derived growth factor-BB
17.3.4.3 Platelet-rich plasma and platelet-rich fibrin
17.3.4.4 Enamel matrix derivate
17.3.4.5 Growth differential factor 5
17.3.5 Bioengineered scaffolds
17.4 Future directions
References
18 - Scaffolds for nasal reconstruction
18.1 Introduction
18.2 Anatomy
18.3 Grafts
18.4 Tissue engineering
18.5 Homografts
18.6 Natural polymers
18.7 Synthetic scaffolds
18.8 Conclusion
References
19 - Scaffolds for the repair of orbital wall defects
19.1 Introduction
19.2 Transplant materials
19.2.1 Autologous bone
19.2.2 Cartilage autografts
19.2.3 Allografts
19.2.4 Xenografts and animal-derived materials
19.3 Synthetic materials for the reconstruction of orbital wall defects
19.3.1 Bioceramics
19.3.1.1 Hydroxyapatite and other calcium phosphates
19.3.1.2 Bioactive glasses
19.3.2 Metals
19.3.2.1 Titanium
19.3.2.2 Cobalt alloys
19.3.3 Polymers
19.4 Composite materials for the repair of orbital wall defects
19.5 Scaffolds for orbital floor reconstruction: challenges and open issues
19.6 Concluding remarks and future perspectives
References
20 - Scaffolds for cleft lip and cleft palate reconstruction
20.1 Introduction on cleft lip and palate reconstruction
20.2 Skin in cleft lip reconstruction
20.2.1 Physiology of the skin/lips
20.2.2 Current surgical treatments
20.2.3 Emerging tissue engineering scaffold technologies
20.3 Oral mucosa in cleft palate reconstruction
20.3.1 Physiology of the oral mucosa
20.3.2 Current surgical treatments
20.3.3 Emerging tissue engineering scaffold technologies
20.4 Muscle in cleft palate reconstruction
20.4.1 Physiology of the muscle
20.4.2 Current surgical treatments
20.4.3 Emerging tissue engineering scaffold technologies
20.5 Bone in cleft palate reconstruction
20.5.1 Physiology of the palate
20.5.2 Current surgical treatments
20.5.3 Emerging tissue engineering scaffold technologies
20.6 Conclusion
20.6.1 Future directions and needs for treatment
References
21 - Scaffolds for temporomandibular joint disc engineering
21.1 Background
21.2 The role of TMJ disc scaffolds
21.3 Scaffolding materials for the TMJ disc
21.3.1 Natural scaffolds
21.3.2 Synthetic scaffolds
21.3.3 Composite scaffolds
21.4 Technologies for scaffolds fabrication
21.4.1 Particulate leaching technologies
21.4.2 Phase separation technologies
21.4.3 Textile technologies
21.4.4 Electrospinning technologies
21.4.5 3D printing and 3D bioprinting techniques
21.5 Biological modifications of scaffolds
21.6 Clinical applications and future directions
References
Part Four: Dental tissue engineering scaffolds
22 - Scaffolds for regeneration of the pulp–dentine complex
22.1 Introduction
22.2 Pulp–dentine biology and response to current treatment therapies
22.3 Role of tissue engineering in regenerative endodontics
22.4 Scaffolds
22.4.1 Definition, ideal requirements, and biomaterial selection
22.4.2 Scaffolds derived from biological sources
22.4.3 Scaffolds of synthetic polymers, bioceramics, and composites
22.4.4 Cell-laden versus cell-free scaffolds
22.4.5 Partial pulp regeneration and complete regeneration of pulp–dentine complex
22.4.6 Advanced scaffolds for pulp–dentine regeneration
22.5 Summary and future perspectives
References
23 - Scaffolds for periodontal tissue engineering
23.1 Introduction
23.2 Periodontal tissue engineering
23.3 Scaffolds in periodontal tissue engineering
23.3.1 Applied biomaterials used in scaffold fabrication for periodontal tissue regeneration
23.3.1.1 Biodegradable natural polymers
23.3.1.2 Biodegradable synthetic polymers
23.3.1.3 Bioceramics
23.3.1.4 Composite biomaterials
23.3.2 Advances in scaffold preparation techniques
23.3.2.1 3D-printed scaffolds
Biphasic scaffolds
Triphasic scaffolds
23.4 Recommendations and future directions
References
24 - Tissue-engineered alloplastic scaffolds for reconstruction of alveolar defects
24.1 Introduction
24.2 Additive manufacturing of synthetic biomaterials for alveolar bone regeneration
24.2.1 Regenerative pharmaceuticals: adenosine receptor stimulation
24.2.1.1 Personalized fabrication of scaffolds
24.3 Integration of tissue engineering principles: translational evidence
24.4 Pediatric alveolar cleft defect regeneration
24.5 Conclusions and future directions
Acknowledgments
Competing financial interests
References
25 - Scaffolds for gingival tissues
25.1 Principles of periodontal treatment
25.2 Guided gingival tissue regeneration
25.3 Nonresorbable gingival membranes
25.4 Resorbable membranes
25.5 Growth factors and cytokines
25.6 Three-dimensional gingival scaffolds
25.7 Gene therapy strategies for gingival tissues
25.8 Conclusion and future perspectives
References
26 - Scaffolds that promote enamel remineralization
26.1 Introduction
26.2 Embryological development of teeth
26.3 Enamel natural genesis
26.3.1 Defining terms
26.3.1.1 Scaffolds
26.3.1.2 Starting cells
26.3.1.3 Processing materials
26.3.2 Enamel structure and ultrastructure
26.4 Technics for enamel rebuilding
26.4.1 Biomimetic methods
26.4.2 Self-­assembling peptide methods
26.4.3 Regeneration of enamel using hydroxyapatite as basement method
26.4.4 Natural or semisynthetic scaffold with stem cell methodology
26.4.5 Synthetic scaffolds
26.5 Biological evaluation of enamel scaffold
26.6 Conclusion
References
Further reading
27 - Scaffolds for dental cementum
27.1 Introduction
27.2 Cementum anatomy, function, and structure
27.3 Cementum formation
27.4 Common problems associated with cementum
27.5 Common resolutions for issues with the cementum
27.6 Cell selection
27.7 Scaffold/structure synthesis
27.7.1 Electrospinning
27.7.2 Self-assembling
27.7.3 Solvent-casting, particulate leaching
27.7.4 Rapid prototyping/3D printing
27.7.5 Supercritical fluid-gas processing
27.7.6 Layered nanocomposites
27.7.7 Thermally induced phase separation
27.7.8 Freeze casting/drying
27.7.9 Gas foaming
27.7.10 Grafts
27.8 Biomaterials for cementum scaffolds
27.8.1 Nonrigid biomaterials
27.8.2 Rigid biomaterials
27.8.3 Other biomaterials
27.9 Summary
References
Further reading
28 - Scaffolds for engineering tooth–ligament interfaces
28.1 Introduction
28.2 Scaffolds for periodontal regeneration
28.2.1 Monophasic scaffolds
28.3 Multiphasic scaffolds
28.3.1 Biphasic
28.3.2 Triphasic scaffolds
28.3.3 Clinical translation and personalized scaffold
28.4 Whole tooth reconstruction
28.5 Conclusion
References
Part Five: Cardiaovascular tissue engineering scaffolds
29 - Whole-heart scaffolds—how to build a heart
29.1 The need for tissue-engineered hearts
29.2 The native human heart: structure and function
29.3 Essential components of an engineered heart
29.3.1 Whole-heart scaffolds
29.3.1.1 Decellularized ECM scaffolds
29.3.1.2 Synthesized scaffolds
29.3.2 Cells
29.3.3 Vasculature
29.4 Building a whole heart in the laboratory
29.4.1 Methods for recellularization
29.4.2 Delivering cells via 3D bioprinting
29.4.3 Perfusion bioreactors
29.5 Moving in vivo
References
30 - Scaffolds for engineering heart valve
30.1 Introduction
30.2 The cardiac cycle
30.3 Heart valves
30.3.1 Aortic valve
30.3.2 Pulmonary heart valve
30.3.3 Mitral heart valve
30.3.4 Tricuspid valve
30.4 Heart valve dysfunction
30.4.1 Aortic regurgitation
30.4.2 Pulmonary atresia
30.5 Current treatment
30.5.1 Mechanical valves
30.5.2 Bioprostethics
30.6 Tissue engineering
30.7 Biomaterials and scaffolds
30.8 Fabrication methods
30.9 Cell sources
30.10 Summary and further directions
References
31 - Scaffolds for blood vessel tissue engineering
31.1 Introduction
31.2 Native blood vessels
31.3 Existing disorders and treatments associated with blood vessels
31.4 Mechanical requirements
31.4.1 Mechanical stretch and burst pressure
31.4.2 Fatigue resistance
31.4.3 Suture retention
31.5 Biomaterial’s requirements
31.5.1 Biodegradability
31.5.2 Biocompatibility
31.5.3 Biomechanical interactivity
31.6 Scaffold fiber diameter and porosity
31.7 Polymers
31.7.1 Natural polymers
31.7.1.1 Collagen
31.7.1.2 Fibronectin
31.7.1.3 Fibrin/fibrinogen
31.7.1.4 Gelatin
31.7.1.5 Chitosan
31.7.2 Synthetic polymers
31.7.2.1 Polylactide
31.7.2.2 Polyglycolide
31.7.2.3 Polylactide-co-glycolide
31.7.2.4 Polycaprolactone
31.7.2.5 Polyurethanes
31.7.3 Natural versus synthetic polymers
31.8 Methods of fabrication
31.8.1 Solvent casting
31.8.2 Freeze-drying
31.8.3 Self-assembly
31.8.4 Electrospinning
31.9 Summary
References
Further reading
32 - Scaffolds for tissue engineering of functional cardiac muscle
32.1 Introduction
32.2 Materials for cardiac tissue engineering
32.3 Scaffolds for improving cell adhesion
32.4 Scaffolds with improved mechanical properties
32.5 Imitating the natural cardiac microenvironment
32.5.1 Controlling the structural and mechanical properties of the scaffold
32.5.2 Controlling the biochemical microenvironment
32.5.3 Covalently linked growth factors
32.5.4 Sustained growth factor release
32.5.5 On-demand growth factor release
32.6 Improving the electrical conductivity of scaffolds
32.6.1 Conductive polymers
32.6.2 Noble metals
32.6.3 Carbon nanoparticles
32.7 Online control and monitoring of tissue function
32.8 Outlook
References
33 - Bioengineered cardiac patch scaffolds
33.1 Introduction
33.2 Cardiovascular anatomy and physiology
33.3 Organogenesis of myocardium
33.4 Common problems and treatment options associated with myocardium
33.4.1 Coronary heart disease
33.4.2 Heart arrhythmia
33.5 Cell selection
33.5.1 Progenitor cells
33.5.2 Pluripotent stem cell
33.5.3 Mesenchymal stem cell
33.6 How to fabricate cardiac patch scaffolds
33.7 Biomaterials for myocardium scaffolds
33.7.1 Natural polymers
33.7.1.1 Collagen
33.7.1.2 Hyaluronic acid
33.7.1.3 Alginate
33.7.2 Synthetic polymers
33.7.2.1 Polyurethanes (PU)
33.7.2.2 Polycaprolactone (PCL)
33.7.2.3 Polyglycerol sebacate (PGS)
33.8 Summary
References
Further reading
Index
A
B
C
D
E
F
G
H
I
J
K
L
M
N
O
P
Q
R
S
T
U
V
W
X
Z
Back Cover

Citation preview

Woodhead Publishing Series in Biomaterials

Handbook of Tissue Engineering Scaffolds: Volume One Edited by

Masoud Mozafari Farshid Sefat Anthony Atala

Woodhead Publishing is an imprint of Elsevier The Officers’ Mess Business Centre, Royston Road, Duxford, CB22 4QH, United Kingdom 50 Hampshire Street, 5th Floor, Cambridge, MA 02139, United States The Boulevard, Langford Lane, Kidlington, OX5 1GB, United Kingdom Copyright © 2019 Elsevier Ltd. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions. This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein). Notices Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. Library of Congress Cataloging-in-Publication Data A catalog record for this book is available from the Library of Congress British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library ISBN (Print): 978-0-08-102563-5 ISBN (Online): 978-0-08-102564-2 For information on all Woodhead Publishing publications visit our website at https://www.elsevier.com/books-and-journals

Publisher: Matthew Deans Acquisition Editor: Gwen Jones Editorial Project Manager: Emma Hayes Production Project Manager: Joy Christel Neumarin Honest Thangiah Cover Designer: Victoria Pearson

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List of contributors

Saiful Ali Department of Biomedical and Electronics Engineering, School of Engineering, University of Bradford, Bradford, United Kingdom Farah Asa’ad Institute of Odontology, Sahlgrenska Academy, University of Gothenburg, Göteborg, Sweden Anthony Atala Wake Forest Institute for Regenerative Medicine, Winston–Salem, NC, United States; Wake Forest University School of Medicine, Winston–Salem, NC, United States Zohya Azhar Department of Biomedical and Electronics Engineering, School of Engineering, University of Bradford, Bradford, United Kingdom Stephen F. Badylak McGowan Institute for Regenerative Medicine, University of Pittsburgh, Pittsburgh, PA, United States; Department of Bioengineering, University of Pittsburgh, Pittsburgh, PA, United States; Department of Surgery, University of Pittsburgh, Pittsburgh, PA, United States Francesco Baino Institute of Materials Physics and Engineering, Applied Science and Technology Department, Politecnico di Torino, Torino, Italy Joseph Bartolacci McGowan Institute for Regenerative Medicine, University of Pittsburgh, Pittsburgh, PA, United States; Department of Bioengineering, University of Pittsburgh, Pittsburgh, PA, United States Yves Bayon Medtronic, Sofradim Production, Trevoux, France Morteza Bazgir Department of Biomedical and Electronics Engineering, School of Engineering, University of Bradford, Bradford, United Kingdom Veerle Bloemen Prometheus, LRD Division of Skeletal Tissue Engineering, KU Leuven, Leuven, Belgium; Materials Technology TC, Campus Group T, KU Leuven, Leuven, Belgium Ciardulli Maria Camilla Department of Medicine, Surgery and Dentistry, University of Salerno, Baronissi, Italy

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List of contributors

Héctor Capella-Monsonís Regenerative, Modular & Developmental Engineering Laboratory (REMODEL), National University of Galway Ireland (NUI Galway), Galway, Ireland; Science Foundation Ireland (SFI) Centre for Research in Medical Devices (CÚRAM), National University of Galway Ireland (NUI Galway), Galway, Ireland Yahya E. Choonara Wits Advanced Drug Delivery Platform Research Unit, Department of Pharmacy and Pharmacology, School of Therapeutic Sciences, Faculty of Health Sciences, University of the Witwatersrand, Johannesburg, South Africa Phil Coates Interdisciplinary Research Centre in Polymer Science and Technology (IRC Polymer), University of Bradford, Bradford, United Kingdom Paulo G. Coelho Department of Biomaterials, New York University College of Dentistry, New York, NY, United States; Hansjörg Wyss Department of Plastic Surgery, New York University School of Medicine, New York, NY, United States; Department of Mechanical Engineering, Tandon School of Engineering, New York University, New York, NY Ricardo Rodriguez Colon Icahn School of Medicine at Mount Sinai, New York, NY, United States Vitor M. Correlo 3B’s Research Group, I3Bs – Research Institute on Biomaterials, Biodegradables and Biomimetics, University of Minho, Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, Guimarães, Portugal; ICVS/3B’s– PT Government Associate Laboratory, Braga/Guimarães, Portugal; The Discoveries Centre for Regenerative and Precision Medicine, Headquarters at University of Minho, Guimarães, Portugal Bruce N. Cronstein Department of Medicine, New York University School of Medicine, New York, NY, United States Eric Dowling Mayo Clinic, Rochester, MN, United States Lisa C. du Toit Wits Advanced Drug Delivery Platform Research Unit, Department of Pharmacy and Pharmacology, School of Therapeutic Sciences, Faculty of Health Sciences, University of the Witwatersrand, Johannesburg, South Africa Monica Talarico Duailibi Universidade Federal de Sao Paulo, Surgery Department – Pos Graduation Program on Translational Surgery of Medical School at Paulista Medicine School, Sao Paulo, Brazil Silvio Eduardo Duailibi Universidade Federal de Sao Paulo, Surgery Department – Pos Graduation Program on Translational Surgery of Medical School at Paulista Medicine School, Sao Paulo, Brazil

List of contributors

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Tal Dvir The School for Molecular Cell Biology and Biotechnology, Faculty of Life Sciences, Tel Aviv University, Tel Aviv, Israel; The Center for Nanoscience and Nanotechnology, Tel Aviv University, Tel Aviv, Israel; Department of Materials Science and Engineering, Faculty of Engineering, Tel Aviv University, Tel Aviv, Israel; Sagol Center for Regenerative Biotechnology, Tel Aviv University, Tel Aviv, Israel Jenna Dziki McGowan Institute for Regenerative Medicine, University of Pittsburgh, Pittsburgh, PA, United States; Department of Surgery, University of Pittsburgh, Pittsburgh, PA, United States Abdelmotagaly Elgalad Regenerative Medicine Research, Texas Heart Institute, Houston, Texas, United States Ron Feiner The School for Molecular Cell Biology and Biotechnology, Faculty of Life Sciences, Tel Aviv University, Tel Aviv, Israel; The Center for Nanoscience and Nanotechnology, Tel Aviv University, Tel Aviv, Israel Ana Marina Ferreira School of Mechanical and Systems Engineering, Newcastle University, Newcastle-upon-Tyne, United Kingdom Roberto L. Flores Hansjörg Wyss Department of Plastic Surgery, New York University School of Medicine, New York, NY, United States R. Gater Institute of Science & Technology in Medicine, University of Keele, Stokeon-Trent, United Kingdom Michael Gelinsky Centre for Translational Bone, Joint and Soft Tissue Research, Faculty of Medicine Carl Gustav Carus, Technische Universität Dresden, Dresden, Germany Piergiorgio Gentile School of Mechanical and Systems Engineering, Newcastle University, Newcastle-upon-Tyne, United Kingdom Liesbet Geris Prometheus, LRD Division of Skeletal Tissue Engineering, KU Leuven, Leuven, Belgium; Biomechanics Research Unit, GIGA - In silico Medicine, Université de Liège, Liège, Belgium; Biomechanics Section, KU Leuven, Leuven, Belgium Della Porta Giovanna Department of Medicine, Surgery and Dentistry, University of Salerno, Baronissi, Italy Mershen Govender Wits Advanced Drug Delivery Platform Research Unit, Department of Pharmacy and Pharmacology, School of Therapeutic Sciences, Faculty of Health Sciences, University of the Witwatersrand, Johannesburg, South Africa

xviii

List of contributors

Salomé Guillaumin Regenerative, Modular & Developmental Engineering Laboratory (REMODEL), National University of Galway Ireland (NUI Galway), Galway, Ireland; Science Foundation Ireland (SFI) Centre for Research in Medical Devices (CÚRAM), National University of Galway Ireland (NUI Galway), Galway, Ireland Grant S. Hamilton III Mayo Clinic, Rochester, MN, United States Zoe Hancox Department of Biomedical and Electronics Engineering, School of Engineering, University of Bradford, Bradford, United Kingdom Nasira Haque Department of Biomedical and Electronics Engineering, School of Engineering, University of Bradford, Bradford, United Kingdom L. Araida Hidalgo-Bastida Centre for Biosciences, School of Healthcare Sciences, Manchester Metropolitan University, Manchester, United Kingdom Katherine R. Hixon Department of Biomedical Engineering, Saint Louis University, St. Louis, MO, United States Camila Hochman-Mendez Regenerative Medicine Research, Texas Heart Institute, Houston, Texas, United States Ming-Yeah Hu Department of Materials Science & Engineering, University of Connecticut, Storrs, CT, United States Sunaina Indermun Wits Advanced Drug Delivery Platform Research Unit, Department of Pharmacy and Pharmacology, School of Therapeutic Sciences, Faculty of Health Sciences, University of the Witwatersrand, Johannesburg, South Africa Sašo Ivanovski The University of Queensland, School of Dentistry, Herston, Qld, Australia Jeffrey Janus Mayo Clinic, Rochester, MN, United States Saeid Kargozar Department of Modern Sciences and Technologies, School of Medicine, Mashhad University of Medical Sciences, Mashhad, Iran Maria Katsikogianni Department of Chemistry and Biosciences, University of Bradford, Bradford, United Kingdom; Interdisciplinary Research Centre in Polymer Science and Technology (IRC Polymer), University of Bradford, Bradford, United Kingdom Zohaib Khurshid Department of Prosthodontics and Implantology, College of Dentistry, King Faisal University, Al-Ahsa, Saudi Arabia

List of contributors

xix

Gahyun Grace Kim Department of Surgery, University of California Irvine, Orange, CA, United States; Sue and Bill Gross Stem Cell Research Center, University of California, Irvine, CA, United States Maryam Koopaie Department of Oral Medicine, School of Dentistry, Tehran University of Medical Sciences, Tehran, Iran Stefanie Korntner Regenerative, Modular & Developmental Engineering Laboratory (REMODEL), National University of Galway Ireland (NUI Galway), Galway, Ireland; Science Foundation Ireland (SFI) Centre for Research in Medical Devices (CÚRAM), National University of Galway Ireland (NUI Galway), Galway, Ireland Pradeep Kumar Wits Advanced Drug Delivery Platform Research Unit, Department of Pharmacy and Pharmacology, School of Therapeutic Sciences, Faculty of Health Sciences, University of the Witwatersrand, Johannesburg, South Africa Jonathan R.T. Lakey Department of Surgery, University of California Irvine, Orange, CA, United States; Biomedical Engineering, University of California Irvine, Irvine, CA, United States; Sue and Bill Gross Stem Cell Research Center, University of California, Irvine, CA, United States Lena Larsson Department of Periodontology, Institute of Odontology, University of Gothenburg, Göteborg, Sweden Hien Lau Biomedical Engineering, University of California Irvine, Irvine, CA, United States Alexander Y. Lin Division of Plastic Surgery, Saint Louis University School of Medicine, St. Louis, MO, United States; St. Louis Cleft-Craniofacial Center, SSM Health Cardinal Glennon Children’s Hospital, St. Louis, MO, United States Han Lin Guanghua School of Stomatology, Hospital of Stomatology, Sun Yat-sen University, Guangzhou, People’s Republic of China Fábio Moyses Lins Dantas INT – National Institute of Technology, Laboratory of Polymeric Materials Technology, Division of Materials Processing and Characterization, Rio de Janeiro, Brazil Yi Lin McGill University, Montreal, QC, Canada; Guanghua School of Stomatology, Hospital of Stomatology, Sun Yat-sen University, Guangzhou, People’s Republic of China Christopher D. Lopez Icahn School of Medicine at Mount Sinai, New York, NY, United States; Hansjörg Wyss Department of Plastic Surgery, New York University School of Medicine, New York, NY, United States

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List of contributors

Robert M. Love School of Oral Health and Dentistry, Griffith University, Gold Coast, Australia Jennifer Cam Luong Biomedical Engineering, University of California Irvine, Irvine, CA, United States James Melville Department of Oral and Maxillofacial Surgery, School of Dentistry, University of Texas Health Science Center in Houston, Houston, TX, United States Peiman Brouki Milan Cellular and Molecular Research Centre, Iran University of Medical Sciences, Tehran, Iran; Department of Tissue Engineering and Regenerative Medicine, Faculty of Advanced Technologies in Medicine, Iran University of Medical Sciences (IUMS), Tehran, Iran M. Rezaa Mohammadi Department of Materials Science and Engineering, University of California, Irvine, CA, United States; Sue and Bill Gross Stem Cell Research Center, University of California, Irvine, CA, United States Alberto Monje Department of Periodontology, Universitat Internacional de Catalunya, Barcelona, Spain Masoud Mozafari Bioengineering Research Group, Nanotechnology and Advanced Materials Department, Materials and Energy Research Center (MERC), Tehran, Iran; Department of Tissue Engineering and Regenerative Medicine, Faculty of Advanced Technologies in Medicine, Iran University of Medical Sciences, Tehran, Iran; Cellular and Molecular Research Centre, Iran University of Medical Sciences, Tehran, Iran Thomas T. Nguyen Department of Oral Medicine, Infection and Immunity, Harvard School of Dental Medicine, Boston, MA, United States Maffulli Nicola Department of Medicine, Surgery and Dentistry, University of Salerno, Baronissi, Italy W. Njoroge Institute of Science & Technology in Medicine, University of Keele, Stoke-on-Trent, United Kingdom Syam Nukavarapu Department of Materials Science & Engineering, University of Connecticut, Storrs, CT, United States; Department of Biomedical Engineering, University of Connecticut, Storrs, CT, United States; Department of Orthopaedic Surgery, University of Connecticut Health, Farmington, CT, United States Claire-Marie Nuttegg Centre for Biosciences, School of Healthcare Sciences, Manchester Metropolitan University, Manchester, United Kingdom

List of contributors

xxi

Masami Okamoto Advanced Polymeric Nanostructured Materials Engineering, Graduate School of Engineering, Toyota Technological Institute, Nagoya, Japan J. Miguel Oliveira 3B’s Research Group, I3Bs – Research Institute on Biomaterials, Biodegradables and Biomimetics, University of Minho, Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, Guimarães, Portugal; ICVS/3B’s– PT Government Associate Laboratory, Braga/Guimarães, Portugal; The Discoveries Centre for Regenerative and Precision Medicine, Headquarters at University of Minho, Guimarães, Portugal Agbabiaka Oluwadamilola Department of Biomedical and Electronics Engineering, School of Engineering, University of Bradford, Bradford, United Kingdom H.A. Owida Institute of Science & Technology in Medicine, University of Keele, Stoke-on-Trent, United Kingdom Olga C. Paiva ISEP - School of Engineering | Polytechnic of Porto, Rua António Bernardino de Almeida, Porto, Portugal Ioannis Papantoniou Prometheus, LRD Division of Skeletal Tissue Engineering, KU Leuven, Leuven, Belgium; Skeletal Biology & Engineering Research Center, KU Leuven, Leuven, Belgium Viness Pillay Wits Advanced Drug Delivery Platform Research Unit, Department of Pharmacy and Pharmacology, School of Therapeutic Sciences, Faculty of Health Sciences, University of the Witwatersrand, Johannesburg, South Africa Sandra Pina 3B’s Research Group, I3Bs – Research Institute on Biomaterials, Biodegradables and Biomimetics, University of Minho, Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, Guimarães, Portugal; ICVS/3B’s– PT Government Associate Laboratory, Braga/Guimarães, Portugal Tehmeena Israr Raja Department of Biomedical and Electronics Engineering, School of Engineering, University of Bradford, Bradford, United Kingdom Murali Ramamoorthi Craniofacial Tissue Engineering and Stem Cells Laboratory, Faculty of Dentistry at McGill University, Montreal, QC, Canada Poornima Ramburrun Wits Advanced Drug Delivery Platform Research Unit, Department of Pharmacy and Pharmacology, School of Therapeutic Sciences, Faculty of Health Sciences, University of the Witwatersrand, Johannesburg, South Africa

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List of contributors

Esther Reina-Romo University of Seville, Department of Mechanical Engineering and Manufacturing, Seville, Spain Rui L. Reis 3B’s Research Group, I3Bs – Research Institute on Biomaterials, Biodegradables and Biomimetics, University of Minho, Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, Guimarães, Portugal; ICVS/3B’s– PT Government Associate Laboratory, Braga/Guimarães, Portugal; The Discoveries Centre for Regenerative and Precision Medicine, Headquarters at University of Minho, Guimarães, Portugal Sofia Ribeiro Regenerative Modular & Developmental Engineering Laboratory (REMODEL), National University of Galway Ireland (NUI Galway), Galway, Ireland; Medtronic, Sofradim Production, Trevoux, France Viviana P. Ribeiro 3B’s Research Group, I3Bs – Research Institute on Biomaterials, Biodegradables and Biomimetics, University of Minho, Headquarters of the European Institute of Excellence on Tissue Engineering and Regenerative Medicine, Parque de Ciência e Tecnologia, Zona Industrial da Gandra, Guimarães, Portugal; ICVS/3B’s–PT Government Associate Laboratory, Braga/Guimarães, Portugal Aicale Rocco Department of Medicine, Surgery and Dentistry, University of Salerno, Baronissi, Italy Luiz C. Sampaio Regenerative Medicine Research, Texas Heart Institute, Houston, Texas, United States Farshid Sefat Department of Biomedical and Electronics Engineering, School of Engineering, University of Bradford, Bradford, United Kingdom; Interdisciplinary Research Centre in Polymer Science and Technology (IRC Polymer), University of Bradford, Bradford, United Kingdom; Department of Materials and Polymer Engineering, Hakim Sabzevari University, Sabzevar, Iran; Department of Biology, Faculty of Sciences, Hakim Sabzevari University, Sabzevar, Iran Scott A. Sell Department of Biomedical Engineering, Saint Louis University, St. Louis, MO, United States Assaf Shapira The School for Molecular Cell Biology and Biotechnology, Faculty of Life Sciences, Tel Aviv University, Tel Aviv, Israel; The Center for Nanoscience and Nanotechnology, Tel Aviv University, Tel Aviv, Israel Lavanya Ajay Sharma School of Oral Health and Dentistry, Griffith University, Gold Coast, Australia

List of contributors

xxiii

Reuben Staples The University of Queensland, School of Dentistry, Herston, Qld, Australia Jean-Philippe St-Pierre Department of Chemical and Biological Engineering, University of Ottawa, Ottawa, ON, Canada John Syrbu Department of Family Dentistry, The University of Iowa College of Dentistry, Iowa City, IA, United States Tara Tariverdian Bioengineering Research Group, Nanotechnology and Advanced Materials Department, Materials and Energy Research Center (MERC), Tehran, Iran Doris A. Taylor Regenerative Medicine Research, Texas Heart Institute, Houston, Texas, United States Andrea Torroni Hansjörg Wyss Department of Plastic Surgery, New York University School of Medicine, New York, NY, United States Simon D. Tran Craniofacial Tissue Engineering and Stem Cells Laboratory, Faculty of Dentistry at McGill University, Montreal, QC, Canada Cristina Tuinea-Bobe Interdisciplinary Research Centre in Polymer Science and Technology (IRC Polymer), University of Bradford, Bradford, United Kingdom Peter Twigg Department of Biomedical and Electronics Engineering, School of Engineering, University of Bradford, Bradford, United Kingdom Cedryck Vaquette The University of Queensland, School of Dentistry, Herston, Qld, Australia Denis Fabricio Viera Rey Department of Chemical and Biological Engineering, University of Ottawa, Ottawa, ON, Canada Maxime M. Wang Department of Biomaterials, New York University College of Dentistry, New York, NY, United States; Hansjörg Wyss Department of Plastic Surgery, New York University School of Medicine, New York, NY, United States Lukasz Witek Department of Biomaterials, New York University College of Dentistry, New York, NY, United States David T. Wu Craniofacial Tissue Engineering and Stem Cells Laboratory, Faculty of Dentistry at McGill University, Montreal, QC, Canada Y. Yang Institute of Science & Technology in Medicine, University of Keele, Stokeon-Trent, United Kingdom

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Pamela Crotty Yelick Tufts University, Department of Oral and Maxillofacial Pathology, Boston, MA, United States Simon Young Department of Oral and Maxillofacial Surgery, School of Dentistry, University of Texas Health Science Center in Houston, Houston, TX, United States Safiyya Yousaf Department of Biomedical and Electronics Engineering, School of Engineering, University of Bradford, Bradford, United Kingdom Mansour Youseffi Department of Biomedical and Electronics Engineering, School of Engineering, University of Bradford, Bradford, United Kingdom Muhammad Sohail Zafar Department of Dental Materials, College of Dentistry, Taibah University, Medina Munawwarah, Saudi Arabia; Department of Dental Materials, Islamic International Dental College, Riphah International University, Islamabad, Pakistan Mahbubeh Zare Department of Biomedical and Electronics Engineering, School of Engineering, University of Bradford, Bradford, United Kingdom Dimitrios I. Zeugolis Regenerative, Modular & Developmental Engineering Laboratory (REMODEL), National University of Galway Ireland (NUI Galway), Galway, Ireland; Science Foundation Ireland (SFI) Centre for Research in Medical Devices (CÚRAM), National University of Galway Ireland (NUI Galway), Galway, Ireland Wei Zhang State Key Laboratory of Polymer Materials Engineering, Polymer Research Institute at Sichuan University, Chengdu, China Zhiguang Zhang Guanghua School of Stomatology, Hospital of Stomatology, Sun Yat-sen University, Guangzhou, People’s Republic of China

Foreword

Beginning in the late 1980s, tissue engineering began to emerge at the intersection of numerous disciplines, combining bioengineering, life sciences, and clinical sciences, to meet a global clinical need for technologies to promote the regeneration of functional living tissues and organs. Scaffolds are essential tools for scientists in the field of tissue engineering. Advances in biomaterials science combined with increasing knowledge of extracellular matrix biology and the role of environmental factors in tissue formation have led to the development of scaffolds tailored to provide appropriate structural support and, in some cases, biological and mechanical cues to promote tissue regeneration. It is impressive to look back over the last 30 years of this field and see the many advances that have been made to harness nature and the human body’s innate ability to regenerate. It is therefore timely to introduce this comprehensive and authoritative textbook on biomaterials. The coeditors, Drs. Masoud Mozafari and Farshid Sefat, who originated the concept for this book, and the chapter authors, all leaders in their field of expertise, have provided an in-­depth content that is dedicated to the topic of composite scaffolds, bringing together all the information related to recent advancements in their application and use in tissue engineering. The combined research interests of the authors in the areas of biomaterials, regenerative medicine, biomedical sciences, and tissue engineering make them a formidable group of experts who have come together to deliver a comprehensive handbook covering all the relevant aspects related to scaffolds from materials, innovative techniques, and approaches, as well as challenges faced and future perspectives. Students, graduates, researchers, clinicians, and individuals from academia, industry, and government will find this textbook to be a go-­to resource on the topic of tissue engineering scaffolds. Collectively, significant advancements in scaffold development have propelled the science forward and closer to the ultimate goal—to provide healing therapies and technologies to patients. There are still challenges to overcome, but knowledge and best practices are constantly changing to propel us forward. We hope that readers use this vast information to lead the field toward future advancements and successes. On behalf of the editors, Anthony Atala, MD Wake Forest Institute for Regenerative Medicine Wake Forest University Winston Salem, NC United States

Preface

Over the last few decades, an increasing number of new tissue engineering scaffolds have been reported, with respect to the repair and regeneration of different tissues and organs. This is mainly due to the multitude of advantages that these porous materials have over more conventional ways of tissue repair. Although there are several publications on the advantages of using scaffolds for tissue repair, the literature has always suffered from lack of an integrated and comprehensive source to adequately address this subject area. Handbook of Tissue Engineering Scaffolds (two volumes) aims to fill this gap by providing a comprehensive and authoritative overview on the recent advancements in the application and use of tissue engineering scaffolds. This handbook deals with the basic principles of scaffolds and then goes to the specific applications for different tissues and organs based on body systems. It contains 13 different parts and 66 chapters presenting almost all the theoretical and practical information needed to design tissue engineering scaffolds. The editors attempted to provide a comprehensive handbook that, on the one hand, strikes a balance among the multiplicity of themes that are related to the design of scaffolds, including materials science, engineering, biology, chemistry, immunology, and transplantation, and, on the other hand, emphasizes on the specific considerations when designing scaffolds for particular tissues and organs. The global market for tissue engineering products is estimated to grow from $7.06 billion in 2016 to reach $16.82 billion by 2023 with an annual growth rate of 13.2%. It is known that scaffolds are among the most important elements of this fast-­growing huge market. Therefore, technological advancements are necessary to overcome the limitations and sustain the development of this market growth. The potential applications of scaffolds are wide among almost all tissues and organs in different body systems including musculoskeletal, craniomaxillofacial, dental, cardiovascular, neural, skin, reproductive, respiratory, urinary, digestive, ocular, endocrinology, and metabolism systems. Handbook of Tissue Engineering Scaffolds is intended to bring all the information together in one major reference, providing an in-­depth understanding of the role of scaffolds and their application and use in different body systems. It is not only a textbook for students in cell biology, biotechnology, and medical courses at advanced undergraduate and graduate levels but also a reference tool for research and clinical laboratories. This handbook represents the combined intellect of many active scientists whose pioneering work has been instrumental to ushering in the field of tissue engineering. We believe that the knowledge of the contributors added an extra value to this handbook toward an in-­depth source for tissue engineering scaffolds. Masoud Mozafari Farshid Sefat Anthony Atala

Acknowledgment

We are grateful to all of those with whom we have had the pleasure to work during this project. We would like to extend our special appreciation to editorial assistants Natasha Welford, Emma Hayes, Tara Tariverdian, Zoe Hancox, Safiyya Yousaf, and Maryam Rahmati for leading a team who did most of the work and deserve the props. Without them, this would not happen.

Introduction to tissue engineering scaffolds

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Farshid Sefat1,2,8,9, Masoud Mozafari3,4,7, Anthony Atala5,6 1Department of Biomedical and Electronics Engineering, School of Engineering, University of Bradford, Bradford, United Kingdom; 2Interdisciplinary Research Centre in Polymer Science and Technology (IRC Polymer), University of Bradford, Bradford, United Kingdom; 3Bioengineering Research Group, Nanotechnology and Advanced Materials Department, Materials and Energy Research Center (MERC), Tehran, Iran; 4Department of Tissue Engineering and Regenerative Medicine, Faculty of Advanced Technologies in Medicine, Iran University of Medical Sciences, Tehran, Iran; 5Wake Forest Institute for Regenerative Medicine, Winston–Salem, NC, United States; 6Wake Forest University School of Medicine, Winston–Salem, NC, United States; 7Cellular and Molecular Research Centre, Iran University of Medical Sciences, Tehran, Iran; 8Department of Materials and Polymer Engineering, Hakim Sabzevari University, Sabzevar, Iran; 9Department of Biology, Faculty of Sciences, Hakim Sabzevari University, Sabzevar, Iran

1.1  Introduction Tissue engineering and regenerative medicine are fast developing approaches in the production of new organs and body tissues. On the other hand, it is a field that seeks to replace/repair or enhance the biological function of a tissue or an organ by manipulating cells via their extracellular environment [1–5]. The concept of directly engineering tissue was articulated in detail in 1985 [6]. The term “tissue engineering” was first used during a meeting sponsored by the National Science Foundation (NSF) in 1987. The first true tissue engineering symposium was held in 1988, where a working definition was proposed [4,5,7] in which tissue engineering was defined as “the application of the principles and methods of engineering and life sciences toward the fundamental understanding of structure–function relationships in normal and pathological mammalian tissue and the development of biological substitutes to restore, maintain, or improve tissue function” [4,5,7]. Even though, everyone believe that the field of tissue engineering may be relatively new, the idea of replacing tissue with another goes as far back as the 16th century [8]. During 1546–99, Gasparo Tagliacozzi initiated a nose replacement that he had constructed. Over the past few decades, there have been a wide range of research studies that have been conducted on the provision of tissue-engineered and regenerative medicine, which lead to a significant improvement in production of scaffolds with similar characteristics to a natural tissue/organ [4,5]. These scaffolds are needed, because of trauma/injury, genetic disorders, and diseases, which can lead to damage and degeneration of tissues in the human body, which necessitates treatments to facilitate their repair, replacement, or regeneration [8]. Handbook of Tissue Engineering Scaffolds: Volume One. https://doi.org/10.1016/B978-0-08-102563-5.00001-0 Copyright © 2019 Elsevier Ltd. All rights reserved.

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Handbook of Tissue Engineering Scaffolds: Volume One

In reality, tissue engineering has a multidisciplinary approach consisting of cell engineering, molecular biology, biomaterial engineering, design, imaging, etc., to develop materials to replace/repair diseased or damaged tissue and restore and improve their function [9]. For instance, bone and joint diseases cause many people to suffer for years with crippling effects. With the progressive aging of the population, the need for functional tissue substitutes is increasing [10]. According to previous research studies, the cost of fractures in the United Kingdom is above £5.1 billion each year [11]. Large bone defects resulting from trauma, tumors, infections, or congenital abnormalities often require reconstructive surgery to restore function [12,13]. Organ transplantation and mechanical devices have revolutionized medical practice but have limitations. New bone tissue engineering strategies have been proposed that promise greater bone restoration without many of the limitations of the current therapies [4,5,14]. Bone tissue engineering is a different method of treatment as compared with drug therapy, gene therapy, or permanent implants because engineered bone becomes incorporated within the patient, giving rise hopefully to a permanent cure for the patients suffering from various bone disorders. Bone cells (osteoblast, osteocyte, osteoclast), osteoconductive factor (three-dimensional matrices or scaffold), and osteoinductive factor (recombinant signaling molecules or growth factors) are the three main key elements (see Fig. 1.1) in the tissue engineering of bone [4,5,16]. This combination of cells, signals, and scaffold is often referred to as a tissue engineering triad [8]. Regenerative medicine for the past two decades has captured the attention of the scientific community mainly in the field of medicine that is expected to be used in near future instead of traditional therapies, which cause enormous side effects. Biomaterial scaffold is one of the main elements in this field, which work parallel to cells, environmental factors, and signaling molecules, which plays an important role in successful functional tissue engineering. Successful research studies were conducted for the past few years, and significant improvement and progress have been reported in reconstruction of various human tissues replacements and prosthesis including lung [17], kidney [18], bladder [19,20], intestine [21], bone [15,22], cartilage [19,20,23–26], skin [27,28], oral tissues [29,30], dental [31], cornea [32,33], blood vessels [34,35], trachea [25], nerve [36,37], and adipose tissue [38]. Regardless of the tissue type, a number of key considerations are important during material section when designing or determining the suitability of a scaffold for use in tissue engineering, including [8]: 1. Biocompatibility:

The first question by scientist who aims to use a biomaterial in the field of regenerative medicine is to do with biocompatibility of that particular scaffold, which has to be Bone tissue engineering

Bone cells

Osteoconductive factors

Osteoinductive factors

Figure 1.1  Three main elements of bone tissue engineering [4,5,15].

Introduction to tissue engineering scaffolds

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implanted within human body. Therefore, good biocompatibility will provide adequate cell attachment, which makes cells to function normally with successful cell migration to the site of injury/implant and finally enable proper proliferation of cells before new matrix establishment. Minimum or no immune reaction without any inflammatory response and rejection is an excellent achievement for any biomaterials within human body. 2. Biodegradability:

The main principle of tissue engineering is to allow cells within human body to release enough enzymes to digest biomaterials and enable cells to produce their own extracellular matrix (ECM) within the site of injury, and at the same time, biomaterials will be faded away and leave cells within own matrix to grow and provide suitable structure for that specific tissue. Therefore, implanted biomaterials do to stay within human body except some metallic implants such as knee or hip metal implants. ECM plays an important role in tissue engineering as cells are capable to attach to these natural materials within human body. Table 1.1 shows the functions of ECM in native tissue and of scaffolds in engineered tissue [39]. On the other hand, the waste materials released from biomaterials should not be toxic and do not harm other tissue/organs. Immune systems such as macrophages provide an excellent support for this kind of degradation within human body [8] 3. Mechanical properties:

Preferably, each body tissue/organ has its own mechanical stability and this is crucial to select a suitable scaffold, which functions well without any failure; therefore, mechanical properties are consistent with the anatomy of each tissue/organ. Scaffold handling is another issue during implantation and surgery; therefore, researchers usually carry out enough investigation to ensure strong biomaterials selected to avoid any disappointment during and after surgery. A serious challenge is to do with orthopedic scaffolds (implants) as too much load from different range of patients could cause severe complications. Another challenge within health care is to do with elderly as the rate of repair slows down. Porosity of implanted materials usually is another challenge because of the problem associated with the balance between mechanical properties and enough porous architecture to allow adequate cell infiltration and vascularization [9,40]. 4. Scaffold architecture:

The architecture of scaffolds is an important factor in regenerating new tissue or organ. For instance, porous titanium construct cup-in-hip implants provide enough porosity to ensure cellular penetration and adequate diffusion of nutrients to cells within the construct and to the ECM. The porosity also allows diffusion of waste products out of the scaffold. Fig. 1.2 shows SEM pictures of the MG63 cells attached on the PC membrane surfaces with different micropore sizes [41]. On the other hand, infiltration of cells and cell attachment are directly related to the mean pore size of the scaffold. Also, the morphology and architecture of cells are determined by a sequence of dynamic interactions that are mediated by a heterogeneous population of transmembrane adhesion molecules [42,43]. Besides mediating cell–cell and cell–extracellular–­ matrix attachment, cell–adhesion molecules facilitate cytoskeletal–membrane

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Handbook of Tissue Engineering Scaffolds: Volume One

Table 1.1  Functions of extracellular matrix (ECM) in native tissues and of scaffolds in engineered tissue. Functions of ECM in native tissues 1.

Provides structural support for cells to reside

2.

Contributes to the mechanical properties of tissues

3.

Provides bioactive cues for cells to respond to their microenvironment

4.

Acts as the reservoirs of growth factors and potentiates their actions

5.

Provides a flexible physical environment to allow remodeling in response to tissue dynamic processes such as wound healing

Analogous functions of scaffolds in engineered tissues

Architectural, biological, and mechanical features of scaffolds

Provides structural support for exogenously applied cells to attach, grow, migrate, and differentiate in vitro and in vivo Provides the shape and mechanical stability to the tissue defect and gives the rigidity and stiffness to the engineered tissues Interacts with cells actively to facilitate activities such as proliferation and differentiation Serves as delivery vehicle and reservoir for exogenously applied growth-­ stimulating factors Provides a void volume for vascularization and new tissue formation during remodeling

Biomaterials with binding sites for cells; porous structure with interconnectivity for cell migration and for nutrients diffusion; temporary resistance to biodegradation upon implantation Biomaterials with sufficient mechanical properties filling up the void space of the defect and simulating that of the native tissue Biological cues such as celladhesive binding sites; physical cues such as surface topography Microstructures and other matrix factors retaining bioactive agents in scaffold

Porous microstructures for nutrients and metabolites diffusion; matrix design with controllable degradation mechanisms and rates; biomaterials and their degraded products with acceptable tissue compatibility

Adapted with permission from B.P. Chan, K.W. Leong, Scaffolding in tissue engineering: general approaches and ­tissue-specific considerations, Eur Spine J 17 (Suppl. 4) (2008) S467–S479.

interaction and signal transduction processes, which control some cell’s function including, cytoskeletal organization, cell motility, cell viability, and receptor activation. Cell adhesion proteins such as selectins and cadherins are often transmembrane receptors. Transmembrane cell adhesion proteins expand across the cell surface membrane and have domains that extend into both the extracellular space and the intracellular space. The extracellular domain of a cell adhesion protein can attach to other molecules that might be either on the surface of neighboring cells (cell-to-cell adhesion) or part of the ECM (cell-to-ECM adhesion) [42].

Introduction to tissue engineering scaffolds

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Figure 1.2  SEM pictures of the MG63 cells attached on the PC membrane surfaces with different micropore sizes: (a) 0.2, (b) 0.4, (c) 1.0, (d) 3.0, (e) 5.0, and (f) 8.0 mm [41].

Cell adhesion proteins bind to specific ligands. There are families of cell adhesion proteins that can be categorized in terms of the structure of the adhesion proteins and their ligands. Homophilic binding is an adhesion between similar adhesion proteins, whereas heterophilic binding is an adhesion between an adhesion protein and some other molecule. For example, scaffolds synthesized from natural ECM such as collagen or fibronectin naturally possess these ligands in the form of Arg-Gly-Asp (RGD) binding sequences (Fig. 1.3), whereas scaffolds made from synthetic materials may require deliberate incorporation of these ligands through, for example, protein adsorption [8] 5. Manufacturing technology:

For commercializing a medical product such as an implant or tissue-engineered construct, cost is an important factor because an expensive product rarely goes through the production line and NHS and even private sectors are not willing to pay significant amount for such products. On the other hand, the development of scalable manufacturing processes to good manufacturing practice (GMP) standard is so crucial. Packaging, storage, sterilization, and shipping are other important factors, which need to be considered. For the past few years, the idea of having off-the-shelf product becomes popular between clinicians and scientists, which of course is an ideal solution for medical implantable applications. The interaction of cells with either natural extracellular matrix or biomaterial (artificial/natural scaffold) is of major importance in tissue development and repair [40,44]. For instance, the interaction and adhesion of bone cells with their surrounding ECM environment influence physiological functions and pathological processes [45]. These physiological functions are normal skeletal development and bone matrix

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Handbook of Tissue Engineering Scaffolds: Volume One

Figure 1.3  Confocal micrograph showing osteoblast cells (green) attached to a highly porous collagen–GAG scaffold (red). The mechanism by which cells attach to biomaterials and scaffolds for tissue engineering is critically important for successful tissue regeneration. Used by permission from F.J. O’Brie, Biomaterials and scaffolds for tissue engineering, Mater Today 14 (3) (2011) 88–95.

production. The interaction of bone cells with ECM is mediated by integrins, a family of cell surface receptors [46,47]. Integrins are a group of membrane spanning receptor proteins, which bind to components of the extracellular matrix [48], and they play key roles in the assembly of the actin cytoskeleton as well as in modulating signal transduction pathways [49] that control biological and cellular functions including cell adhesion, migration, proliferation, cell differentiation, and apoptosis [47,48,50] (Fig. 1.4). Integrins exist as two noncovalently bound α and β subunits [49,51,52]. There are 18α and 8β known subunits, which combine to form at least 24 distinct integrin heterodimers [53]. When integrins are inserted into the leading edge of the cell, then cell surface coupling will predominate in this region, which prevents membrane retraction and provides adhesive traction for cell movement.

1.1.1  Scaffolding approaches in tissue engineering For the past two decades, four major scaffolding approaches for tissue engineering have evolved [39] (Fig. 1.5). Table 1.2 represents the principles and the characteristics of these four approaches [39]. 1. Premade porous scaffolds for cell seeding 2. Decellularized ECM from allogenic or xenogenic tissues for cell seeding 3. Cell sheets with self-secreted ECM 4. Cell encapsulation in self-assembled hydrogel matrix 1. Premade porous scaffolds for cell seeding   

Porosity of scaffolds is a successful approach, which has been used for the past few years extensively. This is mainly for seeding cells, which have been treated in vitro and

Introduction to tissue engineering scaffolds

αIIb

β3

β4

RGD receptors

β8

Laminin receptors

α7

α5

αv

β5

9

α6

β6

α3

α8

β1 αX

αL α1 Collagen receptors

α2

α9

α10

α11

α4 αE

β7

β2 αM

αD

Leukocyte-specific receptors

Figure 1.4  Classification of integrin family of heterodimers. Image adapted from S.V. Hudson, C.E. Dolin, L.G. Poole, V.L. Massey, D. Wilkey, J.I. Beier, M.L. Merchant, H.B. Frieboes, G.E. Arteel, Modeling the Kinetics of integrin receptor binding to Hepatic extracellular matrix proteins, Sci Rep 7 (2017) 12444, https://doi.org/10.1038/ s41598-017-12691-y.

are ready to be injected within the pores of various biomaterials for treatment of different disorders/diseases. Both natural and synthetic scaffolds have been used widely for many applications; however, natural biomaterials showed some limitations such as mechanical stability. However, some researchers developed stronger new scaffolds with new shape and architecture with natural biomaterials once mixed with other synthetic materials, which resulted in a useful composite [54] and cross-linking [55,56]. Synthetic biomaterials can be categorized into inorganic (bioglasses) and organic (synthetic polymers). There are various techniques exist and described in many articles, which enable researchers to produce different types of scaffolds including solvent casting, particulate leaching, gas foaming, freeze drying, and phase separation [39,57]. For porous biomaterials, specific techniques such as selective laser sintering, stereolithography, and 3D printing have been used [39]. In the case of woven and nonwoven fiber structures, scaffolds can be produced using techniques include fiber bonding [57] or electrospinning [58,59]. 2. Decellularized ECM from allogenic or xenogenic tissues for cell seeding

For the past few years, special attention goes toward acellular ECM processed from allogenic or xenogenic tissues, which have been used in some tissue engineering applications such as heart valves [60], vessels [61], nerves [62], tendon, and ligament [63]. Scaffolds fabricated from this approach are well tolerated immunologically as the allogenic or xenogenic cellular antigens have been removed from the tissues as the scaffolds come from a similar source, and it is expected to see minimum or no immune rejection. The main advantage of this approach is lack of cellular components within the natural ECM, which could be achieved using enzymes or detergents [64,65].

1. Pre-made porous scaffolds

3. Cell sheets with secreted extracellular matrix

Native tissues

Raw materials

Confluent cells ECM secretion

Decellularization

Porous scaffolds Cell seeding

Porous scaffolds

Cell sheet

4. Cell encapsulated in selfassembled hydrogel

10

Fabrication technologies

2. Decellularized extracellular matrix (ECM)

Monomer solution Cell Initiation of mixing self-assembly

Cell encapsulated in hydrogel

Lamination

Cell seeding

Cell-seeded scaffolds

Multiple cell sheets

Implantation

Implantation

Implantation

Injection

Defective tissues

Figure 1.5  Schematic diagram showing different scaffolding approaches in tissue engineering. Reused with permission from B.P. Chan, K.W. Leong, Scaffolding in tissue engineering: general approaches and tissue-specific considerations, Eur Spine J 17 (Suppl. 4) (2008) S467–S479.

Handbook of Tissue Engineering Scaffolds: Volume One

Cell-seeded scaffolds

Table 1.2  Characteristics of different scaffolding approaches in tissue engineering. (3) Confluent cells with secreted extracellular matrix

Synthetic or natural biomaterials

Allogenic or xenogenic tissues

Cells

Incorporation of porogens in solid materials; solid free-form fabrication technologies; techniques using woven or nonwoven fibers Seeding

Decellularization technologies

Secretion of extracellular matrix by confluent cells

Seeding

Cells present before self-assembly

Implantation

Implantation

Cells present before extracellular matrix secretion Implantation

Most diversified choices for materials; precise design for microstructure and architecture Time-consuming cell seeding procedure; inhomogeneous distribution of cells

Most nature-simulating scaffolds in terms of composition and mechanical properties

Cell-secreted extracellular matrix is biocompatible

Inhomogeneous distribution of cells, difficulty in retaining all extracellular matrix, immunogenicity upon incomplete decellularization Tissues with high ECM content; load-bearing tissues

Need multiple laminations

Injectable, fast, and simple one-step procedure; intimate cell and material interactions Soft structures

(1) Premade porous scaffolds for cell seeding

Raw materials

Processing or fabricating technology

Strategy to combine with cells Strategy to transfer to host tissues Advantages

Disadvantages

Preferred applications

Both soil and hard tissues; load-bearing tissues

Tissues with high cellularity, epithelial tissues, endothelial tissues, thin layer tissues

(4) Cell encapsulated in self-assembled hydrogel Synthetic or natural biomaterials able to self-­ assemble into hydrogels Initiation of self-assembly process by parameters such as pH and temperature

Introduction to tissue engineering scaffolds

(2) Decellularized extracellular matrix for cell seeding

Scaffolding approach

Injection

Soft tissues

11

Adapted with permission from B.P. Chan, K.W. Leong, Scaffolding in tissue engineering: general approaches and tissue-specific considerations, Eur Spine J 17 (Suppl. 4) (2008) S467–S479.

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Handbook of Tissue Engineering Scaffolds: Volume One

Mechanical and biological properties of the decellularized ECM are the optimum conditions for similar applications within human body. 3. Cell sheets with self-secreted ECM

In normal cell culture, environment cells divide until reaching confluency and focal inhibition in which cell stops dividing further. Cells such as cornea epithelium, endothelium, or vascular endothelium go under multiple divisions and produce a single sheet on cells, and upon confluency, there is a possibility to produce another sheet on top of the first layer. This method known as “cell sheet engineering” or “cell sheet approach” in which cells secret their own ECM upon confluency and enzymatic reaction is not required for cell detachment [15]. The best example for this approach is culturing cells on thermoresponsive polymer in which cells detached by temperature change. This technique introduced by a Japanese research team in which many single cells were created on top of each other [66–68]. Cornea [69], myocardium [70], blood vessel, and trachea are the best examples of cell/tissues for this approach, and research studies were carried out recently using the same technique [71,72] as cell growth in a high density to form tight junctions with other cells and secrete ECM of their own [39]. The main issue with this approach is difficult to construct thick tissues [73,74], which make it less clinically feasible as well as difficult to secret enough ECM with cells such as bone and cartilage. 4. Cell encapsulation in self-assembled hydrogel matrix

Encapsulation techniques were used in many applications in science, and various cells were entrapped within a membrane for specific purposes [75–77]. Hydrogels are the best candidate for encapsulation, which are formed by covalent or ionic cross-linking of water-soluble polymers [39]. Various types of materials were used previously for encapsulating cells such as agarose [78], chitosan [79], poly(ethylene glycol) (PEG) [80], and polyvinyl alcohol (PVA) [81]. The main key point in encapsulation of cell is to use correct cross-linker to enable membrane to be impenetrable to cells, antibodies, and cellular antigens and permeable to some nutrients (glucose). This approach sometimes known as injectable scaffolding or in situ tissue engineering [39] is minimally invasive and useful for defect with abnormal shape; however, the weakness of this approach is its poor mechanical properties (Table 1.3).

1.1.2  Fabrication techniques for tissue engineering application Biomaterials were fabricated with various techniques such as [82]: 1. Biodegradable porous scaffold fabrication a. Solvent casting b. Ice particle leaching method c. Gas foaming/salt leaching method 2. Microsphere fabrication a. Solvent evaporation technique b. Particle aggregated scaffold c. Freeze drying method d. Thermally induced phase separation

Introduction to tissue engineering scaffolds

13

Table 1.3  Scaffolds’ fabrication techniques in tissue engineering applications. Method

Polymers

Unique factors

Application

Biodegradable porous scaffold fabrication Solvent casting/ salt leaching method Ice particle leaching method Gas foaming/ salt leaching method

Absorbable polymer (PLLA, PLGA, collagen, etc.) PLLA and PLGA

PLLA, PLGA, and PDLLA

Biodegradable controlled porous scaffolds

Bone and cartilage tissue engineering

Control of pore structure and production of thicker scaffolds Controlled porosity and pore structure sponge

Porous 3D scaffolds for bone tissue engineering Drug delivery and tissue engineering

High-density cell culture, due to the extended surface area High mechanical stability

Bone repair

Microsphere fabrication Solvent evaporation technique

PLGA, PLAGA

Particle aggregated scaffold

Chitosan, HAP

Freeze drying method

PLGA, PLLA, PGA, PLGA/ PPF, Collagen, and Chitosan PEG, PLLA

Thermally induced phase separation

3D porous sponge structure, durable, and flexible

Bone, cartilage, or osteochondral tissue engineering Tissue engineering scaffolds

Highly porous scaffold for cellular transplantation

Complicated shapes for tissue engineering applications

Porosity and bioresorbability

Cartilage tissue engineering

Biomimetically exhibit biocompatibility and cause minimal inflammatory responses, thrombosis, and tissue damage

Cartilage, bone tissue engineering, and drug delivery

Injectable gel scaffold fabrication Ceramic-based injectable scaffolds Hydrogel-based injectable scaffolds

CP ceramics, HAp, TCP, BCP, and BG Hydrophilic/ hydrophobic diblock and triblock copolymer combinations of PLA, PGA, PLGA, and PEG. Copolymers of PEO and PPO and polyoxamer. Alginates, collagen, chitosan, HA, and fibroin

Continued

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Handbook of Tissue Engineering Scaffolds: Volume One

Table 1.3  Scaffolds’ fabrication techniques in tissue engineering applications.––cont’d Method

Polymers

Unique factors

Application

Hydrogel scaffold fabrication Micromolding

Alginate, PMMA, HA, PEG

Photolithography

Chitosan, fibronectin, HA, PEG, PNIAAm, PAA, PMMA, PAam, and PDMAEM PGS, PEG, calcium alginate, silicon, and PDMS

Microfluidics

Emulsification

Gelatin, HA, and collagen

Microgels, biologically degradable, mechanical, and physical complexity Microwells, microarrays, controlled size, and shape

Microbeads, microrods, valves, and pumps

Microgels, microsensors, cell-based diagnostics

Insulin delivery, gene therapy, bioreactor, and immunoisolation Microdevices, biosensors, growth factors, matrix components, forces, and cell–cell interactions Sensing, cell separation, cell-based microreactors, and controlled microreactors Sustainable and controllable drug delivery therapies

Acellular scaffold fabrication Decellularization process

Biological tissues

Retain anatomical structure, native ECM, and similar biomechanical properties

Tissue engineering

Biocompatibility

Drug delivery, wound healing, soft tissue augmentation, synthetic skin, coatings for implants, and scaffolds for tissue engineering

High surface area, biomechanical, and biocompatibility

Drug delivery, wound healing, soft tissue synthetic skin, and scaffolds for tissue engineering Solar sails, reinforcement, vascular grafts, nonwetting textile surfaces, and scaffolds for tissue

Keratin scaffold fabrication Self-assembled process

Keratin

Fibrous scaffold fabrication Nanofiber electrospinning process

Microfiber wet-spinning process

PGA, PLA, PLGA, PCL copolymers, collagen, elastin, and so forth PLGA, PLA, chitosan, and PCL

Biocompatible fibers with good mechanical properties

Introduction to tissue engineering scaffolds

15

Table 1.3  Scaffolds’ fabrication techniques in tissue engineering applications.––cont’d Method

Polymers

Unique factors

Application

Nonwoven fiber by melt-blown process

Polyesters, PGA, and PDO

Submicron fiber size, highly porous scaffold

Filtration, membrane separation, protective military clothing, biosensors, wound dressings, and scaffolds for tissue engineering

Membranes, hydrogels, foams, microsphere, and particles

Angiogenesis, bone regeneration, and wound healing

Interconnected porous ceramic scaffolds

Bone tissue engineering

Improve biocompatibility or enhance the bioreactivity

Orthopedic application

Functional scaffold fabrication Growth factor’s release process

Collagen, gelatin, alginate, chitosan, fibrin, PLGA, PLA, and PEG

Ceramic scaffold fabrication Sponge replication method Simple calcium phosphate coating method

PU sponge, PVA, TCP, BCP, or calcium sulfate Coating on metals, glasses, inorganic ceramics and organic polymers (PLGA, PS, PP, silicone, and PTFE), collagens, fibers of silk, and hairs

Automation and direct organ fabrication Inkjet printing process

Sodium alginate

To build complex tissues composed of multiple cell types (hydrogel scaffold)

Melt-based rapid prototyping process Computer-aided design (CAD) data manipulation techniques

Biodegradable polymers or blends

Complex 3D solid object, good mechanical strength Design and fabrication of patient-specific scaffolds and automated scaffold assembly algorithm

Biosensor development, microdeposition of active proteins on cellulose, biochips, and acellular polymeric scaffolds Honey comb structure scaffold, hard-tissue scaffolds Develop a program algorithm that can be used to design scaffold internal architectures Continued

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Handbook of Tissue Engineering Scaffolds: Volume One

Table 1.3  Scaffolds’ fabrication techniques in tissue engineering applications.––cont’d Method

Polymers

Unique factors

Application

Organ printing

Tubular collagen gel

Layer-by-layer deposition of cells or matrix

To print complex 3D organs with computer-controlled

For degradable polymers and porous scaffolds, high penetration ability, and compatibility Proven process is safe, reliable, and highly effective at treating single-use medical devices Compatibility, low penetration, in-line sterilization of thin products

Absolute freedom from biological contamination in scaffolds

Scaffold sterilization Ethylene oxide gas (EOG)

Gamma-radiation sterilization

Electron beam radiation

Dry-heat sterilization

Steam sterilization

Surgical disposables: Surgical sutures, bandages, dressings, gauge pads, implants

Commercially successful technology for sterilizing a variety of disposable medical devices with a wide range of densities Efficacy, speed, Heat is absorbed by process simplicity, the exterior surface and lack of toxic of scaffold and then residues passed inward to the next layer Removal of all conPorous scaffold tamination, and scaffor living cell fold can be reused immobilization

Adapted from B. Dhandayuthapani, Y. Yoshida, T. Maekawa, D.S. Kumar, Polymeric scaffolds in tissue engineering application: a review, Int J Polym Sci 2011 (2011) 19 Article ID 290602.

3. Injectable gel scaffold fabrication a. Ceramic-based injectable scaffolds b. Hydrogel-based injectable scaffolds 4. Hydrogel scaffold fabrication a. Micromolding b. Photolithography c. Microfluidics d. Emulsification 5. Acellular scaffold fabrication a. Decellularization process 6. Keratin scaffold fabrication a. Self-assembly process

Introduction to tissue engineering scaffolds

17

7. Fibrous scaffold fabrication a. Nanofiber electrospinning process b. Microfiber wet-spinning process c. Nonwoven fiber by melt-blown process 8. Functional scaffold fabrication a. Growth factor’s release process 9. Ceramic scaffold fabrication a. Sponge replication method b. Simple calcium phosphate coating method 10. Automation and direct organ fabrication a. Inkjet printing process b. Melt-based rapid prototyping process c. Computer-aided design (CAD) data manipulation techniques d. Organ printing

This handbook covers both the basic and clinical sciences of various tissue engineering biomaterials with a view to meet the researchers’ and practitioners’ need. In this book, authors look at various biomaterials’ properties and characterization techniques for specific applications within human body. This handbook provides a comprehensive and authoritative review on recent advancements in the application and use of various types of scaffolds in tissue engineering. Chapters focus on specific tissue/ organ (mostly on the structure and anatomy), the materials used for treatment, natural composite scaffolds, synthetic composite scaffolds, fabrication techniques, innovative materials and approaches for scaffolds preparation, host response to the scaffolds, challenges and future perspectives, and more. Bringing all the information together in one major reference provides an in-depth understanding of scaffold use in different body systems.

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[10] G. Vunjak-Novakovic, I.R. Freshney, Culture of Cells for Tissue Engineering, Wiley, Hoboken, NJ, 2006. [11] O. Strom, The burden of fractures in France, Germany, Italy, Spain, Sweden, and the UK, Osteoporos Int 22 (2011). [12] G.M. Crana, S.L. Ishaug, A.G. Mikos, Bone tissue engineering, Nat Med 1 (1995) 1322–1324. [13] V.I. Sikavitsas, G.N. Bancroft, A.G. Mikos, Formation of three-dimensional cell/polymer constructs for bone tissue engineering in a spinner flask and a rotating wall vessel bioreactor, J Biomed Mater Res 62 (2002) 36–48. [14] D.J. Mooney, A.G. Mikos, Growing new organs, Sci Am 280 (1999) 60–65. [15] F. Sefat, M. Youseffi, M.C.T. Denyer, Imaging via widefield surface plasmon resonance microscope for studying bone cell interactions with micro-patterned ECM proteins, J Microsc 241 (3) (2010) 282–290. [16] N.L. Ramoshebi, N.T. Matsaba, J. Teare, L. Renton, J. Patton, U. Ripamonti, Tissue Engineering: TGF-[beta] Superfamily Members and Delivery Systems in Bone Regeneration, 2002. [17]  S. Moztarzadeh, K. Mottaghy, F. Sefat, A. Samadikuchaksaraei, M. Mozafari, Nanoengineered biomaterials for lung regeneration, Nanoeng Biomater Regen Med (2018) 305–323. [18] T. Tariverdian, P. Zarintaj, P. Milan, M. Saeb, S. Kargozar, F. Sefat, A. Samadikuchaksaraei, M. Mozafari, Nanoengineered biomaterials for kidney regeneration, Nanoeng Biomater Regen Med (2018) 325–344. [19] F. Sefat, T. Raja, M. Zafar, Z. Khurshid, S. Najeeb, S. Zohaib, E. Ahmadi, M. Rahmati, M. Mozafari, Nanoengineered biomaterials for cartilage repair, Nanoeng Biomater Regen Med (2018) 39–71. [20]  F. Sefat, T. Raja, Z. Moghadam, P. Milan, A. Samadikuchaksaraei, M. Mozafari, Nanoengineered biomaterials for bladder regeneration, Nanoeng Biomater Regen Med (2018) 459–474. [21] A. Urbanska, F. Sefat, S. Yousaf, S. Kargozar, P. Milan, M. Mozafari, Nanoengineered biomaterials for intestine regeneration, Nanoengineered Biomaterials for Regenerative Medicine (2018) 363–378. [22] F. Sefat, M.C.T. Denyer, M. Youseffi, Effects of different transforming growth factor beta (TGF-β) isomers on wound closure of bone cell monolayers, Cytokines 64 (2014) 75–86. [23] E. Daghigh Ahmadi, T.I. Raja, S.A. Khaghani, C.F. Soon, M. Mozafari, M. Youseffi, F. Sefat, The role of photonics and natural curing agents of TGF-β1 in treatment of osteoarthritis, Mater Today Procedia 5 (7 Part 3) (2018) 15540–15549. [24] P. Kaur, S.A. Khaghani, Z. Khurshid, M.S. Zafar, M. Mozafari, F. Sefat, Fabrication and Characterisations of Hydrogels for Cartilage Repair Advances in Tissue Engineering and Regenerative Medicine, 2017. [25] T. Raja, M. Mozafari, P. Milan, A. Samadikuchaksaraei, F. Sefat, Nanoengineered biomaterials for tracheal replacement, Nanoeng Biomater Regen Med (2018) 285–303. [26] T.I. Raja, S.A. Khaghani, M.S. Zafar, Z. Khurshid, M. Mozafari, M. Youseffi, F. Sefat, Effect of TGF-β1 on water retention properties of healthy and osteoarthritic chondrocytes, Materia Today Proc 5 (7 Part 3) (2018) 15717–15725. [27] F.J. Bye, A.J. Bullock, R. Singh, F. Sefat, S. Roman, S. Macneil, Development of a basement membrane substitute incorporated into an electrospun scaffold for 3D skin tissue engineering, J Biomater Tissue Eng 4 (2014) 1–7. [28] S.B. Mahjour, X. Fu, X. Yang, J. Fong, F. Sefat, H. Wang, Rapid creation of skin substitutes from human skin cells and biomimetic nanofibers for acute full-thickness wound repair, Burns 41 (8) (2015) 1764–1774.

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[29]  S. Najeeb, Z. Khurshid, M.A. Agwan, S.A. Ansari, M.S. Zafar, J.P. Matinlinna, Regenerative potential of Platelet Rich Fibrin (PRF) for curing intrabony periodontal defects: a systematic review of clinical studies, Tissue Eng Regen Med 1 (2017) 1–8. [30] T. Nejatian, Z. Khurshid, M.S. Zafar, S. Najeeb, S. Zohaib, M. Mozafari, L. Hopkinson, F. Sefat, Dental biocomposites, in: Biomaterials for Oral and Dental Tissue Engineering, Elsevier, 2017, pp. 65–83 (Chapter 5). [31] Z. Khurshid, S. Najeeb, M. Mali, S.F. Moin, S.Q. Raza, S. Zohaib, F. Sefat, M.S. Zafar, Histatin peptides: pharmacological functions and their applications in dentistry, Saudi Pharmaceut J 25 (1) (2017) 25–31 1. [32] P. Deshpande, F. Sefat, C. Ramchadaran, I. Mariappan, C. Johnson, R. Mckean, M. Hannah, V. Sangwan, F. Claeyssens, A.J. Ryan, S. Macneil, Simplifying corneal surface regeneration using a biodegradable synthetic membrane and limbal tissue explants, Biomaterials 34 (21) (2013) 5088–5106. [33] I. Ortega, F. Sefat, T. Paterson, P. Deshpande, C. Ramchadaran, F. Claeyssens, V.S. Sangwan, A.J. Ryan, S. Macneil, Combination of microstereolithography and electrospinning to produce membranes equipped with niches for corneal regeneration, J Vis Exp (2014) e51826. [34] S. Cheng, Y. Jin, N. Wang, F. Cao, W. Zhang, W. Bai, W. Zheng, X. Jiang, Self-adjusting, polymeric multilayered roll that can keep the shapes of the blood vessel scaffolds during biodegradation, Adv Mater 29 (28) (2017) 1700171. [35] K. Sakaguchi, T. Shimizu, T. Okano, Construction of three-dimensional vascularized cardiac tissue with cell sheet engineering, J Control Release 205 (2015) 83–88. [36] A. Mohammadi, A. Maleki-Jamshid, D. Sanooghi, P. Brouki Milan, A. Rahmani, F. Sefat, K. Shahpasand, M. Soleimani, M. Bakhtiari, R. Belali, F. Faghihi, M.T. Joghataei, G. Perry, M. Mozafari, Transplantation of human chorion-derived cholinergic progenitor cells: a novel treatment for neurological disorders, Mol Neurobiol 56 (1) (2019, Jan) 307–318. [37] F. Mohamadi, S. Ebrahimi, M.R. Nourani, K. Mansoori, A.A. Alizadeh, S.M. Tavangar, M. Salehi, F. Sefat, J. Ai, Enhanced Sciatic nerve regeneration by human endometrial Stem Cells in an electrospun poly (ε-caprolactone)/collagen/NBG nerve conduit in a rat, Artif Cells Nanomed Biotechnol 46 (8) (2018, Dec) 1731–1743. [38] N. Amini, N. Vousooghi, A. Alizade, S. Ramezani, M.T. Joghataei, P. Brouki Milan, S. Mehrabi, S. Ababzadeh, F. Sefat, M. Mozafari, Transplantation of Adipose Tissue-Derived Stem Cells into Brain through Cerebrospinal Fluid in Rat Models: Protocol Development and Initial Outcome Data, Current Stem Cell Research & Therapy, 2018. [39] B.P. Chan, K.W. Leong, Scaffolding in tissue engineering: general approaches and tissue-specific considerations, Eur Spine J 17 (Suppl. 4) (2008) S467–S479. [40] T. Tosounidisa, G. Kontakisa, V. Nikolaoub, A. Papathanassopoulosb, P.V. Giannoudisb, Fracture healing and bone repair: an update, Trauma (2009) 1–12. [41] S.J. Lee, J.S. Choi, K.S. Park, G. Khang, Y.M. Lee, H.B. Lee, Response of MG63 osteoblast-like cells onto polycarbonate membrane surfaces with different micropore sizes, Biomaterials 25 (2004) 4699–4707. [42] H. Lodish, A. Berk, S.L. Zipursky, P. Matsudaira, D. Baltimore, J. Darnell, Molecular Cell Biology, W. H. Freeman, New York, 2000. [43] C.Y. Yang, L.Y. Huang, T.L. Shen, J.A. Yeh, Cell adhesion, morphology and biochemistry on nanotopographic oxidized silicon surfaces, Eur Cell Mater 20 (2010) 415–430. [44] C.T. Brighton, S.M. Albelda, Identification of integrin cell-substratum adhesion receptors on cultured rat bone cells, J Orthop Res 10 (6) (1992) 766–773. [45] S.M. Albelda, C.A. Buck, Integrin and other cell adhesion molecules, FASEB J 4 (1990) 2868–2880.

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[46] S.K. Akiyama, K. Nagata, K.M. Yamada, Cell surface receptors for extracellular matrix components, Biochim Biophys Acta 1031 (1990) 91–110. [47] F. Aoudjit, K. Vuori, Integrin signalling in cancer cell survival and chemoresistance, Chemother Res Pract 212 (2012). [48] R.O. Hynes, Integrins: versatility, modulation, and signalling in cell adhesion, Cell 69 (1992) 11–25. [49] M.B. Srichai, R. Zent, Cell-Extracellular Matrix Interactions in Cancer, Integrin Structure and Function, vol. 2, Springer Science, 2010, pp. 19–41. [50] J.J. Worthington, J.E. Klementowicz, M.A. Travis, TGFb: a sleeping giant awoken by integrins, Trends Biochem Sci 36 (2011) 47–54. [51] S.V. Hudson, C.E. Dolin, L.G. Poole, V.L. Massey, D. Wilkey, J.I. Beier, M.L. Merchant, H.B. Frieboes, G.E. Arteel, Modeling the kinetics of integrin receptor binding to hepatic extracellular matrix proteins, Sci Rep 7 (2017) 12444, https://doi.org/10.1038/ s41598-017-12691-y. [52] T. Saito, S.M. Albelda, C.T. Brighton, Identification of integrin receptors on cultured human bone cells, Bone and Joint Surgery Inc 12 (2004) 384–394. [53] R.O. Hynes, Integrins: bidirectional, allosteric signaling machines, Cell Adhes Migrat 110 (2002) 673–687. [54] A.R. Boccaccini, J.J. Blaker, Bioactive composite materials for tissue engineering scaffolds, Expert Rev Med Devices 2 (3) (2005) 303–317. [55] B.P. Chan, K.F. So, Photochemical crosslinking improves the physicochemical properties of collagen scaffolds, J Biomed Mater Res A 75 (3) (2005) 689–701. [56] J.L. Ifkovits, J.A. Burdick, Review: photopolymerizable and degradable biomaterials for tissue engineering applications, Tissue Eng 13 (10) (2007) 2369–2385. [57] S. Yang, K.F. Leong, Z. Du, C.K. Chua, The design of scaffolds for use in tissue engineering: Part II. Rapid prototyping techniques, Tissue Eng 8 (1) (2002) 1–11. [58] Q.P. Pham, U. Sharma, A.G. Mikos, Electrospinning of polymeric nanofibers for tissue engineering applications: a review, Tissue Eng 12 (5) (2006) 1197–1211. [59] T.J. Sill, H.A. von Recum, Electrospinning: applications in drug delivery and tissue engineering, Biomaterials 29 (13) (2008) 1989–2006. [60] R.L. Knight, H.E. Wilcox, S.A. Korossis, J. Fisher, E. Ingham, The use of acellular matrices for the tissue engineering of cardiac valves, Proc Inst Mech Eng H 222 (1) (2008) 129–143. [61] G.H. Borschel, Y.C. Huang, S. Calve, E.M. Arruda, J.B. Lynch, D.E. Dow, W.M. Kuzon, R.G. Dennis, D.L. Brown, Tissue engineering of recellularized small-diameter vascular grafts, Tissue Eng 11 (5–6) (2005) 778–786. [62] S. Hall, Axonal regeneration through acellular muscle grafts, J Anat 190 (1) (1997) 57–71. [63] J.H. Ingram, S. Korossis, G. Howling, J. Fisher, E. Ingham, The use of ultrasonication to aid recellularization of acellular natural tissue scaffolds for use in anterior cruciate ligament reconstruction, Tissue Eng 13 (7) (2007) 1561–1572. [64] S.F. Badylak, Xenogeneic extracellular matrix as a scaffold for tissue reconstruction, Transpl Immunol 12 (3–4) (2004) 367–377. [65]  T.W. Gilbert, T.L. Sellaro, S.F. Badylak, Decellularization of tissues and organs, Biomaterials 27 (19) (2006) 3675–3683. [66] T. Okano, N. Yamada, M. Okuhara, H. Sakai, Y. Sakurai, Mechanism of cell detachment from temperature-modulated, hydrophilic–hydrophobic polymer surfaces, Biomaterials 16 (4) (1995) 297–303. [67] T. Okano, N. Yamada, H. Sakai, Y. Sakurai, A novel recovery system for cultured cells using plasma-treated polystyrene dishes grafted with poly(N-isopropylacrylamide), J Biomed Mater Res 27 (10) (1993) 1243–1251.

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[68] T. Takezawa, Y. Mori, K. Yoshizato, Cell culture on a thermo-responsive polymer surface, Biotechnology 8 (9) (1990) 854–856. [69] K. Nishida, M. Yamato, Y. Hayashida, K. Watanabe, K. Yamamoto, E. Adachi, S. Nagai, A. Kikuchi, N. Maeda, H. Watanabe, T. Okano, Y. Tano, Corneal reconstruction with tissueengineered cell sheets composed of autologous oral mucosal epithelium, N Engl J Med 351 (12) (2004) 1187–1196. [70] T. Shimizu, M. Yamato, A. Kikuchi, T. Okano, Cell sheet engineering for myocardial tissue reconstruction, Biomaterials 24 (13) (2003) 2309–2316. [71] Y. Tsuda, T. Shimizu, M. Yamato, A. Kikuchi, T. Sasagawa, S. Sekiya, J. Kobayashi, G. Chen, T. Okano, Cellular control of tissue architectures using a three-dimensional tissue fabrication technique, Biomaterials 28 (33) (2007) 4939–4946. [72] J. Yang, M. Yamato, T. Shimizu, H. Sekine, K. Ohashi, M. Kanzaki, T. Ohki, K. Nishida, T. Okano, Reconstruction of functional tissues with cell sheet engineering, Biomaterials 28 (34) (2007) 5033–5043. [73] S. Sekiya, T. Shimizu, M. Yamato, A. Kikuchi, T. Okano, Bioengineered cardiac cell sheet grafts have intrinsic angiogenic potential, Biochem Biophys Res Commun 341 (2) (2006) 573–582. [74] T. Shimizu, H. Sekine, J. Yang, Y. Isoi, M. Yamato, A. Kikuchi, E. Kobayashi, T. Okano, Polysurgery of cell sheet grafts overcomes diffusion limits to produce thick, vascularized myocardial tissues, FASEB J 20 (6) (2006) 708–710. [75] R.P. Lanza, J.L. Hayes, W.L. Chick, Encapsulated cell technology, Nat Biotechnol 14 (9) (1996) 1107–1111. [76] G. Orive, R.M. Hernández, A.R. Gascón, R. Calafiore, T.M. Chang, P. De Vos, G. Hortelano, D. Hunkeler, I. Lacík, A.M. Shapiro, J.L. Pedraz, Cell encapsulation: promise and progress, Nat Med 9 (1) (2003) 104–107. [77]  G. Orive, R.M. Hernández, A.R. Gascón, R. Calafiore, T.M. Chang, P. de Vos, G. Hortelano, D. Hunkeler, I. Lacík, J.L. Pedraz, History, challenges and perspectives of cell microencapsulation, Trends Biotechnol 22 (2) (2004) 87–92. [78] A. Batorsky, J. Liao, A.W. Lund, G.E. Plopper, J.P. Stegemann, Encapsulation of adult human mesenchymal stem cells within collagen-agarose microenvironments, Biotechnol Bioeng 92 (4) (2005) 492–500. [79] B.A. Zielinski, P. Aebischer, Chitosan as a matrix for mammalian cell encapsulation, Biomaterials 15 (13) (1994) 1049–1056. [80] C.R. Nuttelman, M.C. Tripodi, K.S. Anseth, Synthetic hydrogel niches that promote hMSC viability, Matrix Biol 24 (3) (2005) 208–218. [81] H. Iwata, H. Amemiya, R. Hayashi, S. Fujii, T. Akutsu, The use of photocrosslinkable polyvinyl alcohol in the immunoisolation of pancreatic islets, Transplant Proc 22 (2) (1990) 797–799. [82] B. Dhandayuthapani, Y. Yoshida, T. Maekawa, D.S. Kumar, Polymeric scaffolds in tissue engineering application: a review, Int J Polym Sci 2011 (2011) 19 Article ID 290602.

Further reading [1] C.T. Brighton, R.M. Hunt, Early histologic and ultrastructural changes in microvessels of periosteal callus, J Orthop Trauma 11 (1997) 244–253. [2]  B.P. Chan, T.Y. Hui, O.C. Chan, K.F. So, W. Lu, K.M. Cheung, E. Salomatina, A. Yaroslavsky, Photochemical cross-linking for collagen-based scaffolds: a study on optical properties, mechanical properties, stability, and hematocompatibility, Tissue Eng 13 (1) (2007) 73–85.

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[3] H. Chang, Y. Wang, Cell responses to surface and architecture of tissue engineering scaffolds, Regen Med Tissue Eng Cells Biomater (2011), https://doi.org/10.5772/21983. [4]  P. Gentile, K. Mccolgan-Bannon, N. Ceretto, F. Sefat, K. Dalgarno, A.M. Ferreira, Biosynthetic PCL-graft-collagen bulk material for tissue engineering applications, Materials (Basel) 10 (7) (2017 Jun 23) pii: E693. [5] S.B. Mahjour, F. Sefat, Y. Polunin, L. Wang, H. Wang, Improved cell infiltration of electrospun nanofiber mats for layered tissue constructs, J Biomed Mater Res A 104 (6) (2016) 1479–1488. [6] T. Nejatian, F. Sefat, T. Johnson, Impact of packing and processing technique on mechanical properties of acrylic denture base materials, Materials 8 (5) (2015) 2093–2109. ISSN 1996-1944.

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Masami Okamoto Advanced Polymeric Nanostructured Materials Engineering, Graduate School of Engineering, Toyota Technological Institute, Nagoya, Japan 

2.1   Introduction 2.1.1  Tissue engineering and scaffolds Natural living tissues have an adaptive potential to their physiological environment because of a genetically programmed self-repair capacity. In contrast, artificial materials are not capable of self-repair or adaption. In this direction, to fabricate artificial constructs for new tissue regeneration, tissue engineering requires applying methods from materials engineering and molecular biology [1,2]. Using tissue engineering, we can fabricate biological substitutes to repair the failed organs or tissues. The growing cells on scaffolds highly engineered structures act as temporary artificial matrix as a “home” for living cell seeding of the target tissues, maintaining the three-dimensional (3D) stable structure (Fig. 2.1). To enhance the repair of the tissue, the scaffold can locally release growth factors (bioactive proteins), enzymes, drugs, and nonviral genes [3–6]. Generally, scaffolds have certain specifications, i.e., high porosity, proper pore size, biocompatibility, biodegradability, and proper degradation rate [7]. During in vitro or in vivo regeneration, the scaffold needs to provide sufficient mechanical support to maintain stresses and loadings generated. For tissue engineering, suitable materials and proper fabricating techniques are two serious issues. In the last two decades, various materials, such as metals, ceramics, and polymers, have been studied to fabricate scaffolds, and a multiple of techniques have been investigated in this area, including a top-down approach that uses a biodegradable polymeric scaffolds [1], 3D printing [8], a bottom-up approach by using cell sheets [9], and layer-by-layer cell assembly [10].

2.1.2   Metal-based scaffolds So far, metals and their alloys were used as materials in orthopedics because of their superior mechanical properties, such as ductility, stiffness, wear and corrosion resistance, and electrical and thermal conductivity [11]. The advantage of metals is that it is easy to make finished materials from raw material due to their bioinertness. In this regard, they have a minimal interaction with living tissue. Metallic-based scaffolds such as titanium, iron oxide, gold [12], silver [13], and magnesium [14] are used in hard tissue engineering in the form of rods, spheres, wires, shells, particles, or fibers [11]. Metals have properties that allow them to easily obtain Handbook of Tissue Engineering Scaffolds: Volume One. https://doi.org/10.1016/B978-0-08-102563-5.00002-2 Copyright © 2019 Elsevier Ltd. All rights reserved.

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Handbook of Tissue Engineering Scaffolds: Volume One

Original scaffold

Cell

ECMprotein

Polymeric scaffold (2, 3)

Implant scaffold Damaged region

Genes (1)

Enhance cellular adhesion and proliferation

1. Bioactive, 2. biocompatible, 3. biodegradable

Figure 2.1  Schematic representation showing the concept of tissue engineering scaffold. Scaffold material (temporary synthetic extracellular matrix (EMC)) is designed as a three-dimensional mirror image, on which cell growth and regenerate the needed tissue. The scaffold can locally release growth factors (bioactive proteins, enzymes, drugs and non-viral genes (DNAs, RNAs)) or antibiotics and enhance ingrowth to treat defects and even support wound healing. The composite systems combining advantages of polymers and ceramics seem to be a promising choice for bone tissue engineering.

forms, dimensions, and desired compositions, and their chemical surface allows them to be functionalized with antibodies, nucleic acids, peptides, and polymers [15]. Porous metals are used to treat bone, as they improve the integration in the human body and the stability of the bone grafts. For preparation of porous implants with dental or orthopedic applications are anodization, sintering, thermal decomposition, plasma spraying [16] with different powder particles, and blasted with stiff particles, laser micromachining technique, electrical discharge compaction, one-step microwave technique, and biomimetic methods [17]. New discoveries made in the field of porous metals support the osseointegration of metallic implants and the cure of bones [11]. Titanium is one of the materials with the best resistance to corrosion and with the highest biocompatibility. By alloying it with aluminum, molybdenum, vanadium, niobium, tantalum, manganese, cobalt, or nickel, its toughness can be improved and its limit of transition temperature is changed [18]. Ti6Al4V, most common titanium alloy, is used for the reconstruction of bone because they improve the adhesion of osteoblasts, as compared with the conventional material. This improvement in adhesion can lead to increased calcium deposits that are very important to bone reconstruction [18]. A layer of porous TiO2 nanotubes can be obtained on the surface of titanium by anodization [19]. By regulating this mechanism, these nanotubes can be created on dental implants realized from titanium [18]. The activity of cells on TiO2 nanotubes in vitro has shown that on the smallest diameters of the nanotubes, adhesion, proliferation, and differentiation of cultured cells are significantly improved for any type of tested cell [20].

2.1.3  Ceramic-based scaffolds More than 40 years have passed since bioceramic materials have been adopted for hard tissue repair. The first publication with respect to the applications of calcium phosphate (CaP) in hard tissue therapy appeared in 1920, whereas the contribution of sintered hydroxyapatite (HA) powder to this field was mentioned exclusively in 1970 [21].

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Usually, bioactive glass materials and CaPs are adopted for reconstruction therapy owing to their performance to support bone formation. The nanoceramics based on CaP may be fabricated through different techniques such as sol-gel, crystallization of aqueous solution under high temperature, wet chemical methods, and biomimetic deposition [22]. The bioceramics can interact with the human body in three distinctive ways [23]: (1) they do not have any contact with the physiological medium, which indicates that they are bioinert, (2) they can suffer chemical reactions o realize an important connection with the surrounding tissue and that denotes that they are bioactive, (3) they can be dissolved in the organism so as to supply the necessary components for tissue repair and that suggests that they are bioresorbable. CaP nanoparticles have the ability to enrich hard tissue formation after being injected into a bone injury. The positive effect of CaP as nanocarrier of growth factors is responsible for osteogenesis (such as vascular endothelial growth factor [VEGF]). Other advantages such as the reproducibility and low cost of the synthesis method, the diminishing immunogenicity, the bioresorbable properties, and injectable CaP nanoparticles offer great opportunities for hard tissue reconstruction [24]. In recent studies, 3D printing is based on customized parts that are typically made to order in unique configurations and in very small quantities. The use of the Internet enhances the possibilities of design sharing and modifications; thus, the porous ceramics can be printed anywhere. Several studies have investigated the application of 3D printing for scaffold fabrication for tissue engineering [25,26]. Zhange et al. explored the feasibility of using a 3D printing process for a high accuracy fabrication of dental ceramic porcelain structures [27]. However, the 3D printing process has a number of control parameters that can significantly affect the fabrication quality. In many cases, materials are supplied as powders and their characteristics such as particle size, shape, and distribution will significantly influence the resulting structure and thus impact on the properties of the porous ceramic [28].

2.1.4   Polymer-based scaffolds Polymer-based scaffolds play an important role in tissue engineering via cell adhesion, proliferation, and formation of new tissue in 3D, exhibiting great potential in a variety of tissues. Pore size, porosity, and surface area are well recognized as immense parameters for the scaffold. Other architectural points, i.e., pore shape, pore wall morphology, and interconnectivity between pores, are also key issues for cell seeding, migration, growth, mass transport, and the tissue formation [1,29]. The natural scaffolds made from collagen are fast replaced with ultraporous scaffolds from biodegradable polymers. Biodegradable polymers were attractive candidates for scaffold materials because they degrade as the new tissues and are formed, eventually leaving nothing foreign in the body in the end. The major challenges of the scaffold manufacture depend on the design and fabrication of customizable biodegradable constructs with desirable properties for promoting cell adhesion and cell porosity along with sufficient mechanical properties, which well match the host tissue having predictable degradation rate and cytocompatibility [1,2].

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The cytocompatibility of the scaffold materials is imperative, i.e., the substrate materials do not exhibit inflammatory response and immunogenicity of cytotoxicity. In bone tissue engineering, a typical porosity of 90% with a pore diameter of ca 300 μm is required for cell penetration and a proper vascularization of the ingrown tissues [30]. Synthetic biopolymers have been used for the fabrication of porous scaffolds by different techniques set up for the purpose. The main limitation for the use of synthetic biopolymers is their low hydrophilicity that causes a low affinity for the cells as compared with the biological polymers. For this reason, the addition of biological components to a synthetic biopolymer demonstrates an appropriate way to produce a bioactive scaffold that can be considered as a system showing both high cell affinity and adequate mechanical stability [31–35]. Gelatin, collagen, laminin, and fibronectin are a variety of extracellular matrix (ECM) protein components that could be immobilized onto the plasma-treated surface of the synthetic biopolymer to enhance cellular adhesion and proliferation. Table 2.1 shows an overview of the discussed biopolymers and their physical properties [36]. The most often utilized synthetic biopolymers for 3D scaffolds in tissue engineering are saturated poly(α-hydroxy esters), including poly(lactic acid) (PLA), racemic mixture of D,L-PLA (PDLLA), and poly(glycolic acid) (PGA), as well as poly(lactic acid-co-glycolic acid) (PLGA) [37–39]. The mechanism of aliphatic polyester biodegradation is the bioerosion of the material mainly determined by the surface hydrolysis of the polymer. Once degraded, natural pathways remove the monomeric components of each polymer. The body already contains highly regulated mechanisms for completely removing monomeric components of lactic and glycolic acids [36]. PLA is cleared via the tricarboxylic acid cycle. PGA is converted to metabolites or eliminated by other mechanisms. Their mechanical properties and degradation rates are controlled by the changing molecular weights and copolymer composition. Poly(ε-caprolactone) (PCL) degrades at a significantly slower rate than PLA, PGA, and PLGA. The slow degradation makes PCL less attractive for biomedical Table 2.1  Physical properties of synthetic biopolymers used as scaffold materials. Thermal properties Biopolymers PLLA PDLLA PGA PLGA (50/50) PCL aMelting

Tm (°C)a 175 – 226 – 60

Tg (°C)b

Tensile modulus (GPa)

Biodegradation time (months)

65 55 40 55 −60

1.2–3.0 1.9–2.4 5.0–7.0 1.0–2.0 0.1–0.7

>24 12–16 6–12 Adjustable: 1–12 >24

temperature. transition temperature. Reproduced with permission from K. Rezwan, Q.Z. Chen, J.J. Blaker, A.R. Boccaccini, Biodegradable and bioactive porous polymer/inorganic composite scaffolds for bone tissue engineering, Biomaterials 27 (2006) 3413–3431. Copyright 2006 Elsevier. bGlass

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applications but more attractive for long-term implants and controlled release application [1,36]. PCL has been used as a candidate polymer for bone tissue engineering, where scaffolds should maintain physical and mechanical properties for at least 6 months [40]. Of particular significance for application in tissue engineering are debris and crystalline by-products, as well as particularly acidic degradation products of PLA, PGA, PCL, and their copolymers [41]. The interactions between stem cell and their environment are very complex and not fully clarified. In previous work, cells respond to the mechanical properties of the scaffolds on which they are growing (see Section 2.2). Rohman et al. [42] reported that PLGA and PCL are biocompatible for the growth of normal human urothelial and human bladder smooth muscle cells. Their analysis of the potential mechanism has indicated that differences in degradation behavior between polymers are not significant. However, the elastic modulus is a critical parameter, relevant to biology at cellular level, and may also have an impact at tissue/organ scales. The elastic modulus is a property that should be considered in the development and optimization of synthetic biopolymers for tissue engineering [42]. As mentioned above, mesenchymal stem cells (MSCs) provided evidence that ECM elasticity influences differentiation. Indeed, multipotent cells are able to start a transdifferentiation process toward very soft tissues, such as nervous tissue, when the substrate elastic modulus (E) is about 0.5 kPa. Intermediate stiffness (∼10 kPa) addresses cells toward a muscle phenotype and harder E (≥30 kPa) to cartilage/bone [43]. This should address an intelligent design of new synthetic biopolymer intended for specific applications [44]. In tissue engineering, synthetic biopolymers presently used are extremely stiff. PLA has a bulk elasticity of E ∼ 1 GPa, which is ten-thousand times stiffer than most soft tissues. Thus engineering of soft tissue replacements needs to explore biopolymers softer than those presently available. In biochemical and biomechanical factors, mismatch can lead to deterioration of tissue regeneration and eventual failure. Because the cell adhesive activity is correlated with signaling and gene expression of the cells which underlie on the ECM (see Section 2.4).

2.1.5   Composite-based scaffolds Bioactive ceramics such as hydroxyapatite (HA) and CaPs for bone repair [45] (as mentioned in Section 2.1.3) showed appropriate osteoconductivity and cytocompatibility because of their chemical and structural similarity to the mineral phase to native bone but are inherent brittleness and have poor shape ability. For this reason, polymer/ bioactive ceramic composite scaffolds have been developed in application for the bone tissue engineering. They exhibit good bioactivity, manipulation, and control microstructure in shaping to fit bone defects [46]. The addition of HA or CaP increases the surface roughness and chemistry, thereby improving cell adhesion and promoting a well-spread morphology. A well-spread morphology is associated with osteogenic differentiation. Further research should focus on optimizing the diffusion of oxygen and nutrients to the inner regions of a bone substitute to promote osteogenic differentiation and the formation of blood vessels.

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Bioactive glass–incorporated hypoxia-mimicking cobalt ions into collagen–­ glycosaminoglycan (CG) scaffolds have been developed and optimized for bone ­tissue regeneration with a view to improving the mechanical and structural properties of the CG scaffold and enhancing the initial angiogenic step vital for bone regeneration [47]. Hypoxia-mimicking bioactive glass composite scaffolds show for the first time promoting angiogenesis and supporting osteogenesis and overcome the problem of inadequate vascularization of grafts commonly seen in the field of tissue engineering. Further development of composite scaffolds seems to be represented their excellent properties against cell activity and new tissue regeneration.

2.2  Cell–ECM interaction and RGD nanospacing Arginine-glycine-aspartate (Arg-Gly-Asp) (RGD) is one of the most effective and often-employed peptide sequences for stimulating cell adhesion on artificial surfaces [48–53]. The RGD adhesion sequence was discovered in fibronectin [53]. The amino acid sequence of RGD is the minimum unit of a cell adhesive activity domain in adherent proteins (ligands of integrins), e.g., laminin, fibronectin, fibrin, vitronectin, and osteopontin. For instance, the aggregation of α2β1 and ανβ3 integrins has been shown to occur specifically during osteogenesis [54,55]. The RGD domains of vitronectin and fibronectin have major roles to play in the spreading of osteoblasts on HA surfaces and to contribute to osteoconductivity [56]. The regulation patterns of RGD domains are useful for fundamental investigation of cell–material interactions [57]. Owing to the size of transmembrane proteins with 8–12 nm in diameter, the RGD amino acid sequence with length of around 10 nm regulates the distribution of the integrin across the cell membrane. A critical RGD spacing was reported in 2004. The cells are extremely sensitive to interligand spacing, possessing different adhesion behavior, i.e., the nanospacing below about 70 nm revealed the promoted cell adhesion and spreading, whereas above 70 nm a clear cytoskeleton and focal adhesion (FA) is not developed [58]. This kind of behavior has been demonstrated in several preosteoblast (MC3T3-E1) cell line and MSCs [48]. The formation of the aggregated protein (FA complex) might depend on the cross-linking of filamentous actins (F-actins) in lateral direction by adapter proteins, where the length of α-actinins and talins is about 60 nm [59]. For the molecular size of F-actins, the 70 nm critical spacing refers to a necessary interval of adjacent F-actins.

2.2.1  RGD nanospacing in 2D substrates with different stiffness The RGD nanospacing effects on chondrogenic differentiation of MSCs on the two-dimensional (2D) substrates were demonstrated by Ding and colleagues [60]. With the help of a nanolithography technology, RGD nanospacing with hexagonal patterns was fabricated on nonfouling poly(ethylene glycol) (PEG) hydrogels. They prepared two nanospacings (63 and 161 nm) where one was below and the other was above the critical nanospacing. For in vitro chondrogenic induction after 1-day and 9-day incubation, both chondrocyte-specific gene expressions (SOX9 and aggrecan) and collagen II proteins were found (Fig. 2.2).

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Large RGD nanospacing

Small RGD nanospacing

α-actinin

More cell spreading and lower chondrogenic extent

F-actin Complex Adhesion and induction

Less cell spreading and higher chondrogenic extent

Adhesion and induction

Figure 2.2  Schematic presentation of the effects of the RGD nanospacing on the chondrogenic differentiation of MSCs. When an RGD nanospacing on a material surface is larger than the critical nanospacing of about 70 nm, focal adhesion complexes and intracellular cytoskeletons could not be effectively formed, contributing to a less cell spreading and a higher chondrogenic extent. In contrast, focal adhesion complexes were well formed in MSCs on the pattern of the small nanospacing of less than 70 nm, resulting in a more cell spreading but a lower chondrogenic extent due to unknown mechanisms [60]. Copyright 2015. Reproduced from Z. Li, B. Cao, X. Wang, K. Ye, S. Li, J. Ding, Effects of RGD nanospacing on chondrogenic differentiation of mesenchymal stem cells, J Mater Chem B 3 (2015) 5197–5209.

Despite the strong cell adhesion for cell differentiation (specific chondrocyte genes) on the small nanospacing, MSCs showed more analogous to the natural chondrocytes on the RGD nanopatterns with the large nanospacing as compared with those on the small nanospacing. Therefore, the large RGD nanospacing affected a suitable chondrogenic differentiation of MSCs on the 2D substrate with poor cell adhesion. For the large nanospacing beyond the critical value, MSCs generated indistinct FA and cytoskeletons, resulting in less cell spreading and unexpected higher chondrogenic differentiation accompanied by unknown mechanisms [60]. A poor spreading area via large RGD nanospacing led to a greater chondrogenic differentiation. Further experiment was conducted to confirm the positive regulation of the p38 phosphorelay cascade by the addition of an inhibitor SB203580 on the chondrogenic induction [60]. In this system, the positive regulation of the p38 was detected. The chondrogenic differentiation was induced by transforming growth factor-β (TGF-β).

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Apart from this, matrix stiffness strongly affected a physical cue for differentiation of stem cell [61]. Soft substrates are adaptive to a neurogenic or adipogenic differentiation. In contrast, stiff substrates are advantageous for an osteogenic differentiation. Substrates of an intermediate stiffness are favorable for a myogenic lineage commitment for tissue cells to feel the stiffness and respond to their substrate [46,62]. More detailed studies and reviews with several commentaries were also described elsewhere [61,63–65]. Soon after, Ding group reported new findings in 2015 [66]. The tethered protein on the substrate surfaces (instead of matrix stiffness) was the imperative cue to regulate stem cells. In their experiment, the nonfouling PEG hydrogels as the matrix were used to prevent nonspecific protein adsorption, at the same time, covalently bound RGD peptides onto the hydrogel surfaces to regulate specific cell adhesion. The decoupling of smart surface chemistry and matrix stiffness is used in this approach. Four substrates (two compressive moduli of the PEG hydrogels with two different RGD nanospacings) were employed for MSCs incubation with the mixed osteogenic and adipogenic medium (Fig. 2.3). This elegant experiment showed that although the situation of the large RGD nanospacing is complicated, the effect of hydrogel stiffness still exists on stem cell differentiation, indicating that matrix stiffness overrides other physical effects in directing stem cell fate. They also demonstrated that RGD nanospacing is an effective modulator of stem cells, no matter on the soft or the stiff hydrogels [66]. The results demonstrated clearly that the matrix stiffness affords an influential regulator of differentiation. At the same time, RGD nanospacing affects the MSCs differentiation regardless of the stiffness of the substrate. In this way, they have concluded that both RGD nanospacing of ligands and matrix stiffness direct stem cell fate. Hence, the effect of RGD nanospacing on stem cell differentiation could not be fully elucidated by cell tension (traction force). They have conjectured that this behavior might be attributed to an outside-in signaling pathway. ALP positive

Cells (%)

Cells (%)

40

130 kPa 3170 kPa

60

80 60 40

49 nm

135 nm

40

0

Undifferentiated 130 kPa 3170 kPa

30

20

20 0

Oil-droplet positive

Cells (%)

100

80

130 kPa 3170 kPa

20 10

49 nm

135 nm

0

49 nm

135 nm

Figure 2.3  Statistical results of stem cell differentiation as functions of hydrogel stiffness and RGD nanospacing. The percentages of ALP positive cells, oil-droplet positive cells, and undifferentiated cells after coinduction are shown. Mean values and standard deviations from four independent experiments are presented. The stiff hydrogels and/or the large RGD nanospacing favored osteogenesis; the soft hydrogels and/or the small RGD nanospacing favored adipogenesis in such a coinduction experiment [66]. Copyright 2015. Reproduced from K. Ye, X. Wang, L. Cao, S. Li, Z. Li, L. Yu, J. Ding, Matrix stiffness and nanoscale spatial organization of cell-adhesive ligands direct stem cell fate, Nano Lett 15 (7) (2015) 4720–4729.

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2.2.2   3D substrates Some of the aforementioned studies may be helpful to design scaffolds with 2D. The results of 2D scaffolds in an in vitro culture system should be transferred to the design of scaffolds with 3D because most cells reside in their native tissue with 3Ds, to understand true cell behavior. To this end, the researchers focus on hydrogels having several properties, such as biocompatible polymeric networks and swelling in water [67]. Matrices with moduli of 11–30 kPa enhanced osteogenic differentiation of MSC in 3D. This was strongly affected with cell traction using hydrogels having 3D RGD introduced. Adipogenesis in alginate hydrogels with matrix moduli of 2.5–5 kPa was confirmed. These results are consistent with previous results on 2D substrate with corresponding stiffness [67]. However, matrix stiffness directed stem cell fate by adjusting binding with integrin via nanoscale reorganization of ligand presentation but not cell morphology and cell tension. Furthermore, the matrix degradation mediates cell traction forces in directing cell fate that was reported in 2013 [68]. The differentiation behavior is same as that expected in hydrogels with 2D, where the traction force generation similarly promoted osteogenesis differentiation, whereas inhibition of the generation of traction forces leads to adipogenic differentiation. The introduction of RGD ligand in 3D is exploited to direct specific cell responses. Until now, however, the effects of clustering in 3D and ligand nanospacing have not yet been well explored.

2.3  Mechanotransduction Another major challenge is a mechanotransduction via nanoscale surface topography on the substrates, where the cells respond to applied forces and exert forces in the substrate, i.e., ECM [69]. Mechanotransduction is well known as a key role in physiological processes in regeneration, homeostasis, aging, and disease [70]. Fig. 2.4 shows the direct physical signal transmission pathway and indirect mechanochemical signal transduction from the ECM [71–73]. Via actin organization, the direct mechanical stimuli from ECM are transmitted, at the same time indirect biochemical signals are generated, where the Ras-Raf-MEK-ERK signaling is well known as the regulation of cell fate and proliferation of MSCs [74]. Changing in the material features of ECM influences the force balance between cells and ECM. Such forces can change nuclear morphology and cytoskeletal structure owing to the traction force generation, which influences cell response and cell fate (chromatin remodeling and DNA unzipping [75]). When the traction forces are generated in the cells underlying substrate, the cells feel stiffness essentially [76]. The cells apply greater traction forces to develop more stable and distinct FA. Owing to the well-organized actin fibers, the cells spread more extensively on the surfaces with rigid property than on soft (compliant) surfaces (Fig. 2.5). When cells embedded within a soft (collagen) 3D scaffold without RGD sequences, the FA proteins could not generate aggregates. However, the FA dispersed through the cytoplasm widely. Via mechanotransductive processes, these proteins are still indirectly affected in speed of motility and modulating cell traction despite the lack of stable FA [77].

ERK Akt YAP/TAZ

Chromatin

Lamins Nesprin MLC

MLC P

YAP/TAZ

Myosin

ROCK MLCK ERK

Akt

Zyxin

PI3K

MEK Raf

Actin

Rho GDP Rho GTP

Vinculin

GEF

Ras

Cell adhesion complex

Talin

Src

FAK

Paxillin

Integrins ECM niche

Figure 2.4  Direct physical signal transmission pathway and indirect mechanochemical signal transduction from the ECM [71–73]. Via integrin transmembrane molecules into the cytoplasm mechanical stimuli are transmitted (right). Copyright 2015. Reproduced from H. Miyoshi, T. Adachi, Topography design concept of a tissue engineering scaffold for controlling cell function and fate through actin cytoskeletal modulation, Tissue Eng B 20 (2014) 609–627.

Round

Compliant

Branched

Flatten

Substrate

Stiff Elasticity

Stiffness Traction force

Neuron

Adipocyte Chondrocyte Myocyte

Motility

Osteoblast

MSC differentiation

Figure 2.5  Cells apply traction forces their underlying substrate, essentially “feeling” this stiffness [71]. In general, cells generate greater traction forces, establish more stable and distinct FA, and form more defined actin stress fibers (red) and spread more extensively on rigid substrates than on compliant substrates. Soft substrates are beneficial for a neurogenic or adipogenic differentiation, stiff substrates are beneficial for an osteogenic differentiation, and substrates of an intermediate stiffness favor a myogenic lineage commitment, for “tissue cells feel and respond to the stiffness of their substrate” [46,62].

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33 Focal adhesion: traction force Cell attachment, proliferation, migration, differentiation

MSC fate

Smart surface: RGD spacing, stiffness, 2-D, 3-D

Figure 2.6  Interrelation between FA formation (F-actin network), traction forces, cell fate, and extracellular cues via nanoscale surface topography on the substrates.

To date, extensive efforts have been made in this complex interrelation between FA formation (F-actin network), traction forces, and extracellular cues via nanoscale surface topography on the substrates to precisely control stem cell fate (Fig. 2.6).

2.4  Surface topography–mediated stem cell fate Topographical surface modification with biochemical stimuli of the scaffolds is an effective way to induce MSCs differentiation with biochemical stimuli. Cells respond to stimuli via mechanotransduction to change their functions [78,79]. In this process, physical stimuli are sensed via the plasma membrane and intracellular signaling is transduced. It is well demonstrated that both cell alignment and migration with a guide are induced by line topographies, whereas a random orientation of cells causes by pillar with high modulus of the materials [79]. Thus, the surface patterns should be taken into account as an indicator to control stem cell fate, including differentiation and gene expression [80]. The combination of both surface topographies and supplemented media for osteogenic differentiation was used to evaluate the effects. Despite many reports, however, tumorigenicity is caused by supplemented media or soluble factors used to achieve high efficacy in vitro. Thus, the experimental results are difficult to translate to the situation [81]. Topographical modifications have an advantage for much stable differentiation control of the cell via the specific site for extracellular cues. Without a chemical stimulus, the effects of different geometric patterns on cell adhesion, proliferation, and osteogenic differentiation of MSCs were detected [82]. Currently, the precise control of MSC differentiation is improved in tissue engineering. Carvalho and colleagues [83] prepared the line and pillar micropatterns of bioactive silica by combination of sol-gel processing and soft lithography technique. For human MSCs, both micropatterns exhibited higher level of osteogenic differentiation with higher level of expression of mature osteoblast genes, higher EMC calcification

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Handbook of Tissue Engineering Scaffolds: Volume One Lines

Pillars

Day 14

Day 7

Flat

Figure 2.7  SEM images of human MSC morphology of the cells adhered to the flat control and micropatterned thin films [83]. Copyright 2016. Reproduced from A. Carvalho, A. Pelaez-Vargas, D.J. Hansford, M.H. Fernandes, F.J. Monteiro, Effects of line and pillar array microengineered SiO2 thin films on the osteogenic differentiation of human bone marrow-derived mesenchymal stem cells, Langmuir 32 (2016) 1091–1100.

as compared with the flat surface as control (Fig. 2.7). Progress of topography-mediated stem cell fate in 3Ds exploits the design criteria of implant surfaces for more effective osteoanagenesis.

2.5  Control of cell migration and cancer invasion As explained in Section 2.4, the cell–ECM interactions regulate signaling and gene expression that underlie cellular processes in migration and cancer invasion. Understanding the interaction between microenvironment and cancer cells is also critical subject to progress in the cancer treatment [84,85]. As mentioned in Section 2.3, cellular microenvironment plays a pivotal role to form tissues and maintain homeostasis by interacting each other. Cancer destroys the normal balance in the microenvironment via the induction of aberrant ECM reconstruction. These disruptions induced gene expression, proliferation, and migration to promote cancer malignancy [86]. Especially, ECM seems to have a crucial role for cancer progression because stiffening and aligned ECM is observed in the vicinity of tumors. During cancer progression, cancer cells and fibroblasts mainly reconstruct the aberrant ECM. The cells do not simply recreate the ECM, and the remodeled ECM provides biochemical and biophysical cues to the neighboring cells (cancer cells and stromal cells) to promote cancer progression [87–90]. Thus, the cell–ECM interactions should be elucidated

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to understand mechanism of the cancer progression. To solve this profound subject, artificial ECM is required, which mimics in vivo environment. As explained in Section 2.4, surface topography of substrate also modulates cellular phenotype, especially, aligned nanofibers or microgrooves elongate cells. These morphological changes of cells are sufficient to affect the gene expression [91]. As some tissues, such as skeletal muscle tissue, ligaments, articular cartilage, and blood vessel walls possess highly oriented organization structure. The aligned substrates might be effective to develop their functions. In fact, the aligned substrates produced by electrospun nanofibers or microgrooves enhance the differentiation of stem cells to chondrocytes [92,93], neurons [94], myocytes [95], and osteoblasts [96]. Cancer cells also receive mechanical cues from extracellular matrix [97]. It has been reported that rigid substrates progress malignant phenotypes via integrin-dependent regulation [98,99]. Furthermore, several researchers suggested that the stiffness of substrate is highly associated with the cellular apoptosis, growth, and motility [76,100,101]. Generally the cancer cells prefer stiffer substrates and the proliferation and motility were enhanced when they were cultured on stiffer substrate. In addition, Ishihara et al. reported that the transcription factor, nuclear factor κB (NF-κB), is activated by stiff substrate via actomyosin contractions [102]. As NF-κB is associated with the cellular proliferation, adhesion, and apoptosis [103], its activation should be inhibited. Collagen fibers nearby tumor tend to be aligned, and cancer cells reorganize collagen fibers to be aligned. This collagen alignment contributes to the tumor progression [104,105]. As well as tissue engineering and regenerative medicine, designing scaffolds are important to understand the mechanism of cancer progression. As aforementioned, cancer cells are significantly depending on ECM condition. Thus, several researchers have been challenging to establish in vitro tumor models for evaluation of cell behavior [106,107]. Claudia et al. and Bray et al. reported that tumor vascularization was enhanced and less drug sensitivities were observed when the cancer cells were cultured on porous poly(lactide-co-glycolide) (PLG) scaffolds or glycosaminoglycan-based hydrogel [108,109]. Jeon et al. and Bersini et al. have established the microfluidic system to investigate cancer metastasis behavior [110,111]. The capability of metastatic cancers to shift motility modes is one of the main features of invasion. The tumor microenvironment is able to elicit epithelial–mesenchymal transition (EMT), where an epigenic program leads epithelial cells to lose their cell–cell and cell–ECM interactions to undergo cytoskeleton reorganization and to gain morphological and functional characteristics of mesenchymal cells. Artificial nanofiber scaffolds can induce EMT for some breast and lung cancer cells [112–114]. Considering these great efforts, designing scaffold properties have possibilities to control vascularization, EMT, and inflammation phenomena. These phenomena are important not only for cancer progression but also for tissue regeneration [115–119]. However, the cell–ECM interactions have not been elucidated yet. The cancer therapies with poor prognosis prompt us to conduct a novel study on an effective cancer therapy. That is, breakthroughs in effective cancer therapies are imperative. Okamoto group reported new findings in 2017 [120]. Their study was aimed to assess the combination of both surface topographies (fiber alignments) and different stiffness

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of the polymeric substrates (PLLA and PCL) to evaluate the effect on the cellular morphologies, proliferation, motility, and gene expression regarding EMT of two different types of human breast cancer cells (MDA-MB-231 and MCF-7). The cellular morphologies (roundness and nuclear elongation factor), E-cadherin and vimentin expression, and cellular motility in terms of cellular migration speed, persistent time, and diffusivity were discussed comprehensively. They have demonstrated that the microenvironment of cell culture substrates influences cancer progression and metastatic potential.

2.6  Scaffold for gene delivery A scaffold-mediated gene transfer system is another major challenge, where DNA/ material complexes without viral vectors are directed into cells and intracellular compartments for gene expression [121,122]. DNA has a high negative charge and a high molecule more than 1 million Da. A central obstacle to clinical gene therapy remains the effective way of using nonviral delivery. Thus, the low versatility, toxicity, and tumorigenicity of viral vectors are overcome by using nonviral vehicles. The transfection process involves the uptake of extracellular molecules, e.g., DNA, RNA, or oligonucleotides through the cell membrane into the cytoplasm and also into the nucleus of eukaryotic cells. The DNA can be incorporated into a cell’s genetic material and produce specific proteins when it has penetrated the nucleus, [123]. The DNA cannot combine into the host chromosome, i.e., transient transfection. This is distinguished from stable transfection. That is, the foreign DNA is integrated into the chromosome and passed over to the next generation. In this regard, scaffold-mediated gene delivery enables localized delivery of a therapeutic gene. This is one of the advantageous strategies for transfection. The scaffold-mediated DNA delivery is mainly taken up by the surrounding cells at the implant sites, resulting in limiting unwanted exposure in other areas [124]. The structure of the scaffold is regulated for sustained gene delivery acting as a “reservoir,” which gradually releases DNA/calcium phosphate (CP) complex over time. The scaffold degradation rate can be also designed for required release rate [125]. The protection of the DNA/CP complex is another advantage of the scaffold materials. Current trends seek the scaffold-mediated gene delivery complex, as well as stimulatory functions promoting cell proliferation and new tissue formation in 3Ds [126]. Surface area, surface charge, crystallinity, and CP chemistry also play an important role in gene loading efficiency. Mg2+-doped HA particle increases the surface positive charge of the CP nanoparticle and hence increased its DNA loading capacity because of the electrostatically attraction forces. The presence of β-tricalcium phosphate phase into HA enhanced gene delivery properties by increasing its solubility inside endosome [127]. By using a laminin/DNA/HA complex, the successful improvement of the gene transferring was reported by Oyane et al. [128]. The gene transferring efficiency of the laminin/DNA/HA complex was one to two orders of magnitude higher as compared with that of a conventional DNA/CP complex. That is, laminin provides regions of high DNA concentration between cell and the surface of the complex, resulting in enhancement of cell adhesion and spreading.

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The elegant investigation was conducted to demonstrate the effect of mineral properties on plasmid DNA (pDNA) release from CP mineral coatings [129]. The coatings were conducted in simulated body fluid (SBF) solutions of varying ion content and concentration with pDNA. The pDNA-containing mineral coatings by coprecipitation had different Ca/P ratios and mineral structures. Under simulated physiological conditions, these differences resulted in varying pDNA release rates. The DNA-CP chemical coatings had feature sizes of around 500 nm, and their size reduction led to an increase in gene delivery. This was believed to be due to the ability of cells to endocytose the smaller particles, which facilitated the release of DNA from CP matrix. In addition, the efficiency of the gene transfer from the mineralized surface suggested that the amount of pDNA affected the gene expression [129,130]. Choi et al. [131] examined the pDNA release from CP coatings formed on PLGA substrates with the intrinsic properties of the CP chemical coating and the surrounding solution conditions. Both polyplexes and lipoplexes are created from cationic lipids, liposomes, or polymers (poly(l-lysine), poly(d-lysine), and polyethyleneimine) where nucleic acids can electrostatically condense into nanoparticles. The charged complexes have several drawbacks during delivery in vivo including inflammatory toxicity, rapid aggregation, and high clearance from the bloodstream [132]. Several attempts have been made to reduce inflammatory cytotoxicity and improve the pharmacokinetics of these nanoparticles by introducing PEG as a stealth moiety to the surface of the nanoparticle [133]. Although this attempt has been found to increase circulation time of cationic liposomes, PEGylated neutral liposomes with doxorubicin maintained higher plasma concentration, led to higher localization within the targeting tissue, and delivered more effective than the cationic liposomes themselves [134]. Fig. 2.8 illustrates the scaffold-mediated

Plasmid

Polymer Polyplexes

Scaffold

Scaffold-polyplex combinational system

Figure 2.8  The scaffold–polyplex combinatorial system and its components. Copyright 2010. Reproduced from S. O’Rorke, M. Keeney, A. Pandit, Non-viral polyplexes: Scaffold mediated delivery for gene therapy, Prog Polym Sci 35 (2010) 441–458.

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polyplex gene delivery as a combinatorial system [126]. The development of the scaffold-mediated gene delivery has been conducted in different sizes and shapes, from the macroscale to the nanoscale depending on the targeted clinical application.

2.7  Scaffold for multimodal drug delivery Biological phenomena including inflammation, angiogenesis, and tissue remodeling are quite complex. The interlinked processes are also regulated by a spatiotemporal manner of biological processes [135–142]. The factors in the local microenvironment are elaborated and follow a defined path. For this reason, the single-factor delivery is oversimplify biology and has not proven to be clinically efficacious in pathological states. In this regard, the strategies of the delivery of several factors to modulate several stages have paved the way for the pathology. The multiple deliver system including a number of strategies offers opportunities in a temporal multimodal release. These strategies are of importance and demonstrated as the methodologies (Fig. 2.9) [3]. An alginate system was used to sequential delivery of three factors to induce neovascularization [138]. The alginate sulfate is capable of the loading of vascular endothelial growth factor (VEGF), or heparin-binding factors, platelet-derived growth factor-BB (PDGF-BB), and TGF-β. A released VEGF by burst is followed by delayed release of PDGF-BB and TGF-β, which is close to normal blood vessel formation. In case of the use of alginate only, a significant difference was not appeared in the

Figure 2.9  Two typical strategies that have been used to achieve multi-modal release. (a) The use of microspheres in a scaffold to promote differential release of two factors, as the contents of the spheres (drawn in red) are released slower than the contents of the hydrogel (drawn in blue). (b) The use of materials that have a differential affinity for the biomaterial. In this case, the factor drawn in red is released slower than the factor drawn in blue. This may be a natural phenomenon due to the interaction between the biomaterial and the loaded therapeutics, or may be engineered into the material using linker systems [3]. Copyright 2014. Reproduced from S. Browne, A. Pandit, Multi-modal delivery of therapeutics using biomaterial scaffolds, J Mater Chem B 2 (2014) 6692–6707.

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release profile of these factors, i.e., the release also quickly occurs for each factor. Three months later, the blood vessel density was increased by this differential release profile in vivo. The releasing timing for the multiple factors and ingenious changes in characteristics of the materials affect biological processes. A PEG-maleimide gel was prepared with RGD sequences, and several growth factors bind to RGD domains to control the growth factor release through protease-cleavable linkers. Dual delivery of hepatocyte growth factor (HGF) and VEGF was demonstrated to improve the function of ischemia and reperfusion in the rat myocardium [139].

2.8  Scaffolds for bone tumor destruction In bone tumor detection, the development of multifunctional scaffolds loaded therapeutic genes and anticancer drugs has been reported [143]. This is new area in the research of metastatic cancer using bone scaffolds to tackle metastatic spread of cancer cells and their many associated issues (Fig. 2.10) [144]. Doxorubicin-loaded liposome for patients’ protection is used to treat ovarian cancer and Kaposi’s sarcoma in the past. Recent advances have been done using HA nanoparticles, where the nanoparticles used in scaffolds seeded with metastatic breast cancer cells and the adsorption of adhesive serum proteins was enhanced in comparison with much larger nanoscale 1. 3D porous bone scaffold

2. Drug attachment Collagen

Nanohydroxyapatite

Anticancer drugs

Calcium, phosphorus and other inorganic constituents

3. Anticancer drug-loaded bioscaffold implantation Cancerous bone tissue

Implanted drug-loaded bioscaffold

Healthy bone

Figure 2.10  Bone scaffolds constructed to encourage bone cell proliferation, and to load anti-cancer drugs in order to kill cancerous cells [144]. Copyright 2013. Reproduced from G. Blackburn, T.G. Scott, I.S. Bayer, A. Ghosh, A.S. Biris, A. Biswas, Bionanomaterials for bone tumor engineering and tumor destruction, J Mater Chem B, 1 (2013) 1519–1534.

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HA particles. The larger particles led to enhancement of expression of the osteolytic factor rather than smaller nanoparticles [143]. The potential application of nanoscale properties will continue to progress in controlling breast cancer. Drug-loaded bone scaffold is a new area to be addressed. Hopefully the drug-loaded bone scaffolds will encourage osteoblast proliferation for healthy cells, while combating malignant tumor with metastatic spread. This is extremely challenging for the treatment of cancer.

2.9  Scaffolds for cell separations In the field of cell injection therapy, purified cell suspension and cell separation methods are of importance for effective treatment with high transportation efficiency [145]. Temperature-controlled cell adhesion/detachment surface is elegant method for cell separation because this does not require modification of cells surface, unlike fluorescence activated cell sorting [146]. Finally, native nonlabeled cells can be recovered. Poly(N-isopropylacrylamide) (PIPAAm)-modified microfiber mat prepared by electrospinning methods was reported [147]. PIPAAm exhibits lower critical solution temperature of 32°C (near body temperature), where PIPAAm becomes hydrophilic at low temperature (20°C) owing to hydration, and becomes hydrophobic at high temperature (37°C) owing to dehydration. Adhesion and detachment of various cell types, such as human umbilical vein endothelial cells (HUVECs), normal human dermal fibroblasts (NHDFs), human adipose-­ derived stem cells (ADSCs), adipocytes, and human microvascular endothelial cells (HMVECs), were examined. The microfibers exhibited thermally controlled cell separation by selective adhesion of NHDFs in a mixed cell suspension that also contained HUVECs. Furthermore, ADSCs exhibited thermally modulated cell adhesion and detachment, whereas adhesion of other ADSC-related cells was low. Finally, ADSCs can be separated from a mixture of adipose tissue-derived cells simply by changing the temperature. The PIPAAm-modified microfibers are potentially adaptive to temperature-modulated cell separation materials [147]. Detection of circulating tumor cells (CTCs) in peripheral blood is of great significance for the early diagnosis, the prediction of cancer development, and the evaluation of efficacy, prognosis, and individualized treatment of tumors. However, it is very difficult to isolate or capture CTCs from the blood via conventional means due to the extremely low concentration of CTCs in the blood (approximately one CTC among a billion blood cells). Circulating tumor cells (CTCs) are cancer cells shed from primary tumor tissue into the bloodstream, which are closely associated with cancer metastasis and recurrence [148]. Functional electrospun nanofibers and surface immobilization of targeting molecules allow for specific and effective capture of cancer cells. In general, the cell suspensions or blood samples containing cancer cells are first incubated with nanofibers for a certain time period under static conditions, and through the interaction of the topographic features of nanofibers and/or the immobilized targeting molecules, cancer cells can be attached onto the surface of nanofibers. Xiao et al. prepared hyaluronic acid (HyA)–modified poly(vinyl alcohol)/poly(ethyleneimine) (PVA/PEI) nanofibers and used for cancer cell (HeLa) capture [148].

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The capture efficiency of HeLa cells using HyA-PVA/PEI-Ac nanofibers is up to 85.0% at 240 min. Anti-CD20-modified nanofibers were incubated in the Granta-22 cell suspension for 45 min, and the captured cells were stained [149]. The anti-CD20 immobilization rendered the nanofibers with an ability to effectively capture Granta-22 cells. The cells were able to be wrapped or buried by fibers, which was another important factor for enhanced cell capture as revealed by microscopic analysis. Using inorganic nanofibers (epithelial cell adhesion molecule antibody [anti-EpCAM]-immobilized TiO2 nanofibers [TiNFs] coated on a silicon substrate), two EpCAM-positive cancer cell lines (colorectal cancer cell line HCT116 and gastric carcinoma cell line BGC823) and two EpCAM-negative cancer cell lines (cervical cancer cell line HeLa and chronic myelogenous leukemia cell line K562) were chosen for static cell capture assay to optimize TiNF packing density and incubation time [150]. In this way, even though much effort has been devoted to the development of nanofibers for cancer cell capture and release, some challenges still remain because of the extremely low concentration and heterogeneity of CTCs. For example, the nanostructures of nanofibers facilitate cancer cell adhesion but meanwhile also lead to nonspecific attachment and entrapment of blood cells, thus decreasing the purity of captured CTCs [148].

2.10  Future direction and conclusions It is well known that in tissue engineering the scaffold’s architecture from the macroto nanoscale is crucial for proper cellular interactions, nutrient transport, mechanical stability, and ultimately functional tissue formation. In the last decade, a variety of 3D cell culturing methodologies and materials have been reported. A large number of tissue-specific applications are presented throughout this chapter. Despite developments in material selection and scaffold design for 3D cell culturing, it can still be difficult to achieve tissue biomimicry in these scaffolds, which significantly affects the resulting tissue regeneration via ECM microenvironment. In addition, most attempts do not meet the requirements of practical and clinical applications. So far, tissue engineers may be limited by their scaffold fabrication techniques. Quite recently, for scaffold-free formation of 3D cell cultures, a powerful technology based on the magnetic levitation of cells was reported [151]. This technique is an alternative to both scaffold-based and scaffold-free 3D cell culture systems, which assemble into a 3D cell culture while secreting their own ECM. The magnetic levitation platform creates a suitable microenvironment for the cells, which retain their cellular activities while creating 3D organoid structures. Application of magnetic forces principle through magnetic levitation for biological systems has been investigated. The magnetic levitation platform is also utilized for the formation of a 3D cell culture while using agarose gel as a scaffold material [152]. This scaffold-free 3D cell culture system without using complex facilities or chemicals can apply regenerative medicine and 3D bioprinting [8]. Ultimately, some technologies will aid in bridging the gap between research and the clinic, making tissue engineering a viable strategy for the regeneration of whole organs in the body.

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R. Gater, W. Njoroge, H.A. Owida, Y. Yang Institute of Science & Technology in Medicine, University of Keele, Stoke-on-Trent, United Kingdom 

3.1   Introduction During embryonic development, primary cell layers known as “germ layers” work to give rise to all cell types required by the body. In humans, three germ layers develop known as the endoderm, mesoderm, and ectoderm. Once these cells aggregate, begin to communicate, and function together, tissue structures are formed [1]. The endoderm gives rise to the epithelial linings of digestive tract and lung tissues, as well as structures such as the stomach, liver, and pancreas. Alongside this the mesoderm aids in production of the cardiac, skeletal, and smooth muscle, as well as red blood cells and tissue surrounding the kidneys. Finally, the ectoderm is known to give rise to tissue with an epidermis such as hair, nails, and tooth enamel. Additionally, parts of the ectoderm develop further into the neural crest and neural tube, which aid to produce tissues such as the nervous system and brain. Because of this crucial role, some literature considers the neural crest as a fourth germ layer, despite being derived from the ectoderm. For these tissues to provide the specialized support mechanisms required for cellular function, native tissue structures contain an intricate microenvironment known as the extracellular matrix (ECM). Tissue ECM differs considerably between different cell and tissue types, playing key roles in the generation of diverse tissues and organs. Research into the generation of functional tissues through tissue engineering approaches necessitates a thorough understanding of the diversity of native ECM, in terms of its chemical and physical properties. This understanding gives rise to the design of artificial ECM in structures known as scaffolds, which can lead to the “development” journey of tissue formation in vitro, while still following in vivo paths.

3.2  Characterization of native tissues 3.2.1  Common chemical components in ECM ECM is a complex mixture of molecules secreted by cells to provide their surrounding structural and biochemical support. ECM characteristics can differ depending on the requirements of a particular tissue. Tissue-specific expression and synthesis of structural proteins and glycoprotein components result in unique functional and biological characteristics at distinct locations. Handbook of Tissue Engineering Scaffolds: Volume One. https://doi.org/10.1016/B978-0-08-102563-5.00003-4 Copyright © 2019 Elsevier Ltd. All rights reserved.

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The most abundant fibrous protein within ECM is collagen, which functions to provide the “scaffolding” of tissues for the attachment of proteoglycans and other ECM components. The name “collagen” is used as a generic term for proteins forming a characteristic triple helix of three polypeptide chains. All members of the collagen family form these supramolecular structures in the extracellular matrix, although their size, function, and tissue distribution vary considerably. There are 28 different types of collagen identified in vertebrates, with fibrillar collagen being the most abundant accounting for approximately 90% of all collagen in the body. Fibrillar collagen includes types I, II, III, V, XI, XXIV, and XXVII, with type I being the most common. Type I collagen is found in various tissues such as skin, connective tissue, vasculature, and the organic part of bone. Meanwhile, type II is found abundantly in tissues such as cartilage and the vitreous humor of the eye, accounting for up to 90% of collagen in these structures [2]. Collagen type III is widely distributed in collagen I containing tissues with the exception of bone and is an important component of reticular fibers in the interstitial tissue of the lungs, liver, dermis, spleen, and vessels. This homotrimeric molecule often contributes to mixed fibrils with type I collagen and is most prominent in highly compliant connective tissues [3–5]. Furthermore, type IV is the main collagen component of basement membrane, being a network-forming collagen that underlies epithelial and endothelial cells and functions as a barrier between tissue compartments [3,6]. Across all tissues, glycosaminoglycans (GAGs) are an important ECM component. GAG molecules are long unbranched polysaccharides with a repeated disaccharide unit. They attach to proteoglycans, adhesive glycoproteins, and fibrous proteins (e.g., collagen) to provide structural support [3,7] and play a major role in maintaining the equilibrium of healthy tissue by protecting and preserving ECM proteins and cytokines. The major GAG types with physiological significance in tissue can be categorized into four groups: chondroitin/dermatan sulfate, heparin/heparan sulfate, keratan sulfate, and hyaluronan.

3.2.2  Specific characteristics in ECM The characteristics of a tissue are ultimately determined by the type of cells, GAGs, proteoglycans, and proteins present. For example, connective tissue may typically have collagen fibrous protein present, surrounded by the proteoglycans decorin and biglycan, with chondroitin/dermatan sulfate GAG chains attached. Different tissues have differing properties and functional adaptations in ECM. There are diverse variations in ECM, for instance, soft or stiff ECM, stratifying (epithelial) ECM, layered/zonal topographic ECM, and vascularized ECM embedding multicellular tissues. These native scaffolds offer the principles in smart design and guidance for tissue engineers to produce artificial ECM (scaffolds) for tissue regeneration in vitro.

3.2.3  Mechanical properties - hard versus soft tissues The properties of native hard tissue (e.g., bone, teeth) are particularly unique in comparison with other tissues. Interestingly, water is a key component of bone tissue accounting for approximately 25% of its mass [8]. Approximately half of the dry weight of bone tissue is made up of mineral salts, mainly calcium and phosphate.

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The remaining 25%–30% of bone tissue mass is then comprised of fibrous proteins, primarily collagen. Research aiming to characterize the GAG and proteoglycan content of bone tissue has previously determined the presence of chondroitin sulfate and hyaluronic acid GAGs, with the associated proteoglycans (decorin, biglycan, and fibromodulin) and glypican [9]. This mixture of hard and soft material appears to be what gives bone its strength, with the hard material providing rigidity and the soft material providing some room for compression to avoid breakages. The mineral content of bone is believed to be the main influence of its mechanical properties, with increased mineralization making the bone stiffer, yet less capable of compression to withstand shock forces. Cancellous or “spongy” areas of bone tissue are believed to be better at withstanding strain forces, since it is porous and contains fluids such as blood vasculature and bone marrow. On the other hand, cortical or “compact” bone is stiffer and therefore better at withstanding stress forces, in comparison with strain, because of its high mineral content. These biological and mechanical properties are important factors to consider during the development of scaffolds aiming to mimic native hard tissue, such as bone. A common example of native soft tissue is the skin. Skin is a large, complex organ that provides the tissues, it encloses with protection from environmental stresses and serves as a sensory interface between the body and its surroundings. On a basic level, the skin provides a physical barrier that serves the dual function of excluding external stressors, including potential pathogens, and maintaining homeostasis. For the most part, this comprises heat regulation through sweating and vasoconstriction/vasodilation, control of fluid entry and loss by serving as a semiimpermeable barrier, synthesis of vitamin D through ultraviolet radiation in sunlight, sensory perception, and immune cell action [10–13]. Skin is a soft tissue exhibiting key mechanical behaviors. It must be flexible enough not to impair body motion, while being tough enough to resist tearing and piercing, as well as maintaining the ability to return to its original state. Skin can be described as anisotropic, viscoelastic, and nonlinear, resulting in an ability to endure large deformations. Because of the viscoelastic properties, skin undergoes a phenomenon known as preconditioning, where under cyclic loads the stress–strain relationship continuously alters until a steady state is reached [11,12,14,15]. The stratum corneum is the stiffest of the skin layers, therefore the least extendible under applied load. This layer exhibits less viscoelastic and preconditioning behavior compared with other layers but still maintains a nonlinear stress–strain relationship under applied tension. The underlying dermis contributes a large amount to the overall mechanical characteristics of skin; this layer consists of a dense network of collagen and elastin fibers, which allow for high levels of deformation. The underlying hypodermis is the softest of the three layers and evenly transfers loads from the upper skin layer to the underlying tissues. Hence it is important when applying loads perpendicularly to the surface of the skin [15–18].

3.2.4  Tissue with stratified epithelium (skin, lung, cornea, conjunctiva) Some native tissue types within the body possess a layer of cells lining the tissue surface, known as an epithelium. Functions of these cells include barrier protection,

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selective absorption/secretion, transcellular transport, and environmental sensing. Tissue epithelium does not contain blood vessels and therefore receives nourishment from the tissue it lines via a membrane known as the basement membrane. A common example of tissue with an epithelial layer is the lungs. In lung tissue, the alveoli are lined with thin, flattened cells known as the squamous epithelium. This is advantageous for lung gaseous exchange because it creates a short distance between air inside the alveoli and the blood capillaries. The lung trachea and respiratory tract are also lined with cells known as ciliated epithelium. These have tiny hairs known as cilia that function to sweep away any inhaled debris. All GAG types are believed to be present within lung tissue, with hyaluronic acid being the most abundant nonsulfated GAG and heparan sulfate being the most abundant sulfated GAG [19]. The skin is a highly organized, stratified structure consisting of three main layers: the epidermis, dermis, and hypodermis. The superficial epithelium layer, the epidermis, is approximately 75–150 μm in thickness [12,13,20] and consists largely of outward moving cells, keratinocytes, which are formed by division of cells in the basal layer of the epidermis. The second layer is the dermis, which is a dense fibroblastic connective tissue layer of thickness 1–4 mm. It mainly consists of collagen fibers, ground substance, and elastin fibers, and it forms the major mass of the skin. As a result of the collagen and elastin fibrils, the dermis has a mainly mechanical function, allowing for high levels of deformation, as the fibers stretch and reorientate [10,21]. The most numerous of the dermal cells are fibroblasts. They are responsible for the manufacture of all dermal connective tissue elements or their precursors. The third layer is the hypodermis, which contains areolar connective and adipose tissues, connecting the underlying muscles to the skin. The thickness of the layers varies considerably over the surface of the body. The areolar tissue consists of collagen and elastin fibers much like the dermis, and many migrating white blood cells that aim to destroy any pathogens that enter the body. The adipose tissue stores fats and nutrients as a potential energy source and provides cushioning for bony prominences [10,13,17]. Other examples of tissues, which possess an epithelial layer, are eye tissues, such as the cornea and the conjunctiva. Corneal epithelium functions to offer a barrier of protection for the ocular surface, including resistance from excess tear fluid and prevention of bacteria from entering the corneal stroma. Corneal epithelium contains several cell layers, including columnar/basal cells, polyhedral/wing cells, and squamous cells with flattened nuclei. It is reported that the corneal epithelial layers undergo constant mitosis because of the migration of columnar and polyhedral cells, as well as the constant loss of squamous cells as they are washed away by tear film. Underneath the corneal epithelium, the fibrous protein present within corneal tissue itself is primarily collagen. This collagen is highly aligned to enable tissue transparency for vision. Meanwhile, the GAG content of the corneal tissue is reported to be a combination of chondroitin sulfate/dermatan sulfate, heparan sulfate, and keratan sulfate [22]. Conjunctival tissue is a mucous membrane that covers the inside of the eyelids and fore part of the sclera [23]. The area of conjunctiva most commonly associated with disease is the bulbar conjunctiva, making it an interesting area for clinical research. The bulbar conjunctiva sits loosely on top of the sclera, separated by a thin, fibrous tissue layer known as Tenon’s capsule. Similar to other tissues, conjunctiva consists

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of three layers; fibrous, adenoid, and epithelium. Conjunctival epithelium contains several cell types including goblet cells, melanocytes, Langerhan’s cells, and the epithelial cells. Similar to the cornea, conjunctival epithelium functions to offer a barrier of protection for the ocular surface. Here, the goblet cells secrete mucus to trap and prevent bacteria from entering ocular structures such as the cornea. The fibrous protein present within conjunctival ECM itself is primarily collagen, with a sparse distribution of elastin [24]. Minimal previous literature reports the precise GAG content of conjunctival ECM; however, as a connective tissue with the presence of collagen, it can be assumed that chondroitin sulfate/dermatan sulfate GAG chains are present and attached to the proteoglycans decorin and biglycan. One study confirms a content of mainly chondroitin sulfate within conjunctival tissue, with no other GAG molecules reported [25]. The GAG content of scleral tissue (beneath the conjunctival layer) is reported to contain primarily chondroitin sulfate/dermatan sulfate, with a small quantity of hyaluronan [26].

3.2.5  Zonal, layer-specific tissues One example of layer-specific tissue is blood vessels. Blood vessels facilitate gaseous exchange, waste/nutrient transport, and cellular immune defense. Native blood vessel walls, excluding capillaries, are arranged into three concentric layers, which include intima, media, and adventitia. The innermost layer (intima) is comprised of a continuous single layer of endothelial cells that are in contact with the blood. This layer is the central barrier to the escape of plasma and is sealed with tight junctions. Underneath the intima is a dense elastic layer known as the internal elastic lamina, separating it from the subsequent layers. This is then followed by the middle layer (media), which is composed of smooth muscle (spindle-shaped) cells that provide structural support, mechanical strength, and contractility. Smooth muscle cells are arranged helically surrounded by elastin and collagen fibers forming a matrix (subendothelial matrix). The medial layer is separated from the adventitia (outer layer) by a dense elastic membrane, theexternal elastic lamina. The outermost (adventitial) layer consists of a connective tissue sheath without a distinct external border and has a function of anchoring the blood vessel to extracellular matrix (ECM). Thus, this layer has a role in tethering the blood vessel to surrounding tissues [27,28]. Articular cartilage is anisotropic in nature and is also organized into distinct zones. There are at least three architectural zones (superficial, middle, and deep) with striking variations between their structure, chondrocyte phenotype, ECM composition, and mechanical properties [29–31]. In the superficial zone, chondrocytes are elongated as a result of the tightly packed collagen fibers, which have a parallel orientation to the surface to dissipate high tensile strength. Also the concentration of proteoglycans is lower than that in middle and deep zones [30,32]. The middle zone contains rounded chondrocytes with a random collagen fiber orientation, while also having a large amount of collagen II and proteoglycans. In this zone, the collagen fibers are randomly oriented to provide resistance to the multidirectional compressive force [30,33]. The deep zone contains chondrocytes stacked in columns with radial collagen architecture and high proteoglycan concentration. This radial orientation provides cartilage with a high compressive stress resistance [30,32].

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3.2.6  Vascularized tissues In the human body, the vascular system forms an extensive, highly branched, and hierarchically organized network. This majority of tissues and organs are vascularized. The highly organized blood vessel network maintains tissues’ homeostasis through the mass transport of gas, liquids, nutrients, cells, signaling molecules, and waste products. Bone is a composite organ that fulfills several interconnected functions, including locomotion, involvement in phosphate and calcium metabolism, synthesis of endocrine molecules, and hematopoiesis, which may conflict with each other in pathological conditions. Vasculature is indispensable for appropriate bone development, regeneration and remodeling, and is a key interface between the various interconnected functions. It serves as a source of oxygen, nutrients, hormones, neurotransmitters, and growth factors [34,35]. The main blood supply of healthy long bones is derived from the principal nutrient arteries, which penetrate the cortex and perfuse the medullary sinusoids and then exit via multiple small veins [36]. Estimates of the proportion of the cardiac output received directly by the skeleton range from approximately 5.5%–11%. The rich perfusion of bone reflects not only the requirements of the bone cells (i.e., osteoblasts, osteocytes, and osteoclasts), but also the requirements of the marrow (i.e., hematopoietic lineage cells, stromal cells, and adipocytes), as well as endothelial cells. The vascular supply of bone enables the rapid growth and remodeling (including mechanical responsiveness) that would not otherwise be possible in essentially avascular cartilage [36,37]. Bone formation and development occurs through two distinct processes: intramembranous and endochondral ossification. The vascularization is the prerequisite of two different processes of ossification. In intramembranous bone formation, mesenchymal stem cells (MSCs) can be transported through capillaries and differentiate into mature osteoblasts, which then deposit bone matrix and lead to bone formation. During the endochondral ossification, the chondrocytes secrete angiogenic growth factors promoting the invasion of blood vessels, which then transport a number of highly specialized cells and replace the cartilage with bone and bone marrow [38]. In the absence of adequate blood supply, bone has been shown to display reduced growth and repair, experience loss of density, and eventually undergo necrosis. The blood vessels in skin tissue are comprised of a number of cellular and noncellular elements. Endothelial cells (ECs) form the interface between intravascular and extravascular compartments and serve as a selective barrier for the diffusion of cells and macromolecules between compartments. Along with a basement membrane that surrounds the ECs and additional cells such as smooth muscle cells and pericytes, dermal blood vessels are also surrounded by a variety of cells originating from the bone marrow such as mast cells, macrophages, dendritic cells, and mononuclear cells. These bone marrow–derived cells preferentially reside in a perivascular location in the skin. Blood vessels in the skin form a deep plexus in the subcutaneous fat and provide nutrition to the sweat glands and the hair papillae, and a superficial plexus in the papillary dermis [39,40]. The skin is supplied by arteries that originate from their underlying source vessels and are either destined for the skin (direct perforators)

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or from secondary branches of vessels that supply other underlying tissues (indirect perforators). The plexuses that form cutaneous vasculature are formed from either the direct or indirect perforators. The arteries and their associated microvasculature are referred to as angiosomes. The density of angiosomes varies from site to site, and this is primarily determined by whether the skin served is fixed to underlying tissues or is freely moving. Fixed sites tend to have more angiosomes that demonstrate limited branching, whereas large areas of movable skin may be served by a single feeder vessel that branches extensively in the superficial structures [40,41].

3.3  Scaffold designs to mimic the native structure of tissues A current strategy in the tissue engineering field is to induce formation of the specialized features of native tissue, through the design of biomimetic scaffolds. It is hoped that the ability to produce natural tissues through biomimetic scaffolds will aid in the development of cell and tissue replacement therapies, as well as provide a better understanding of health and disease development mechanisms. However, the development of scaffolds to mimic native ECM structurally and chemically is particularly challenging. Attempts have been made to replicate the ECM utilizing a variety of materials both natural and artificial, and these have been used with varying levels of success. Learning from nature, various culture and fabrication techniques have been developed to generate diverse tissues with differences in composition and architecture.

3.3.1  Scaffolds for soft tissue To mimic native soft tissue structures such as skin and conjunctiva, scaffolds using hydrogels and fibrous materials are common. Biocompatible polymers for these applications are considered to be either natural, semisynthetic, or synthetic. Polymers containing natural ECM components provide enhanced biocompatibility to hydrogels, providing mechanical integrity to tissues, as well as enhanced support and regulation for cellular processes. Although synthetic polymers can aim to mimic natural ECM components, a naturally occurring polymer such as collagen, elastin, or fibrinogen is thought to provide cells with a more physiologically relevant substrate on which to reside. A suitable native skin substitute should ideally have the ability to be kept sterile, act as a protective barrier, have a low inflammatory response, and allow for water vapor transmission across the material [42]. There are several scaffold designs attempting to mimic native skin tissue, a common type being porous scaffolds and hydrogels. Most recent examples include a fibrinogen-modified sodium alginate sponge scaffold developed by Soloveiva et al. [43], as well as a silk fibroin and functionalized citrus pectin derived scaffold developed by Turkkan et al. (2018) [44]. High porosity allows for better ECM secretion and nutrient supply for cells, as well as ensuring an even spread of cells across the scaffold without clustering together. However, optimization for the best pore size intended for particular cell types can be challenging.

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Hydrogel-based scaffolds are another common scaffold design in skin tissue engineering. Some of the most recent examples include a functional chitosan-based hydrogel developed for skin wound dressing and drug delivery by Liu et al. [45], plus a poly vinyl alcohol–poly acrylic acid–derived double layered hydrogel for wound dressings by Tavakoli et al. [46]. Hydrogel scaffolds can also allow for good porosity, as well as a high biocompatibility and controlled biodegradation rate. However, because hydrogels are soft structures often with a high water content, reduced mechanical strength can be a limitation. Fibrous scaffold designs can also offer a highly microporous structure, as well as improved mechanical strength for the material. Recent examples include a soy protein/cellulose-derived nanofiber scaffold to mimic skin ECM developed by Ahn et al. [47] and an electrospun poly(lactic-co-glycolic-acid)-derived nanofibrous scaffold by Zhu et al. [48]. However, the surface functionalization of nanofibers can be challenging to develop in this type of scaffold design. Future work in this area may eventually aim to incorporate drugs and biological molecules into fibrous scaffolds, for growth factor or drug release applications to aid in skin wound healing. Model constructs to mimic native conjunctiva are advantageous for the study of wound healing mechanisms in conjunctival tissue after glaucoma surgery [49], as well as for the study of drug treatments to treat diseases of the conjunctiva. Notable studies for polymeric scaffold designs to model conjunctiva include He et al. (2016) [50], who found poly-d-lysine-coated silk to be the most biocompatible polymer substrate for maintaining human conjunctival goblet cells in culture. Being a fibrous tissue, fibrous scaffold designs are also advantageous for mimicking conjunctival tissue to study ocular surface disease. Garcia-Posadas [51] also previously developed a fibrinbased 3D model for the successful culture of human conjunctival fibroblasts. Because of the fibrous protein content of native conjunctival tissue being primarily collagen, collagen hydrogel-based scaffolds to seed conjunctival cells have also been popular in past research [52]. Similar to scaffold design mimicking skin tissue, future work to incorporate biological molecules, such as growth factors, into fibrous and hydrogel scaffolds modeling conjunctiva would be advantageous. This would allow researchers to more closely mimic disease mechanisms of native conjunctival tissue and improve the testing of new drug treatments.

3.3.2  Tissue models with epithelium (coculture + multilayer scaffolds) As outlined previously, an epithelial layer can offer an additional barrier of protection, as well as improve the transcellular transport and environmental sensing of particular tissues. In an attempt to mimic this, multilayer scaffold designs and coculture models are common in the tissue engineering field. Development of lung tissue models can vary depending on the area of lung tissue that researchers would like to mimic. For example, models to mimic the alveoli may include a squamous epithelium, whereas models mimicking the trachea/airway may incorporate a ciliated epithelium instead. There are several multilayer and coculture scaffold designs attempting to mimic areas of native lung tissue. Some recent

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examples include an epithelial–mesenchymal coculture model of human bronchial tissue by Ishikawa et al. [53], as well as a triple layered coculture model of human respiratory tract by Blom et al. [54]. By including multiple cell layers, these models offer an improved opportunity for the study of full-thickness lung tissue. Similar to lung tissue, multilayer and coculture scaffold designs are also popular in research trying to mimic native skin tissue. As outlined previously skin tissue consists of three main cell layers, the epidermis, dermis, and hypodermis, making multilayer model designs particularly appropriate. A previous study of this includes [55] who assembled a multilayer model with hyaluronic acid and poly-l-lysine (for the epidermal layer) onto a porous hyaluronic acid scaffold (for the dermal layer), with human keratinocytes seeded on top. Models such as these are an example of how several scaffold design ideas are frequently combined in current research, such as using coculture techniques in addition to porous and hydrogel scaffold designs. Wilson et al. [56] have developed a multiple-layered corneal model with epithelial cells grown on a stromal layer (Fig. 3.1). The stromal layer was constructed by collagen type I scaffolds. The stromal cell orientation could be controlled by adding orthogonally aligned nanofiber [57]. Epithelial cells could be easily incorporated to the top of the stromal layer. The cross-talk of epithelial–stromal cells regulated stromal cells’ phenotype significantly.

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Combined scaffold design ideas are also frequently seen in the development of recent conjunctival membrane models. Examples include a model developed by Garcia-Posadas et al. [51] consisting of conjunctival fibroblasts seeded within a fibrin-based scaffold with epithelial cells seeded on top. Yao et al. [58] also previously developed a coculture model consisting of a collagen and poly(l-lactic acid co-ε-caprolactone)-derived porous nanofibrous scaffold seeded with conjunctival epithelial cells to stimulate stratification. Furthermore, a recent ocular surface and tear film model was developed by Lu et al. [59] using the coculture of rabbit conjunctival epithelium and lacrimal gland cell spheroids to mimic the aqueous and mucin tear film layers within an optimized air–liquid interface. Although multilayer and coculture methods offer a more realistic mimic of native tissue structures in comparison with monocultures, these are still limited and do not completely mimic a natural ECM structure. To overcome this challenge, continuous research into more accurate scaffold organization, as well as improved mechanical strength, porosity, and biocompatibility for the desired tissue type, is required [60]. Application of surface modifications is one of the approaches to modify scaffolds and cell sheets. For example, Yang et al. [61] recently found the use of collagen and alginate nanofilms improved the mechanical property of cell sheets, which could then be assembled as multilayers. 3D printing technology is another approach, which can be used to produce ECM scaffolds with improved accuracy to native tissue. For example, Hoshiba and Gong [62] recently worked on 3D printed poly(d, l-lactic acid)-derived scaffolds. These could be deposited with ECM constituents using the culture of fibrosarcoma cells on the scaffolds, before subsequently being decellularized for use in other applications. Improved techniques such as these are likely to revolutionize the way in which researchers design scaffolds to mimic native tissue structures in the future.

3.3.3  Scaffolds with zonal, layered structure Considering the prevalence and importance of zonal variations in normal articular cartilage, recent studies have aimed to engineer cartilage with zonal structure or function, or both. Replication of the zonal organization of tissue-engineered cartilage is one of the multiple strategies to generate functional tissue. Yucekul et al. [63] previously demonstrated the potential repair and regeneration of cartilage tissue by using a multilayered biomimetic scaffold. This study proposed a biodegradable, trilayered (poly(glycolic acid) mesh/poly(l-lactic acid)-colorant tidemark layer/collagen I, and ceramic microparticle-coated poly(l-lactic acid)-poly(ε-caprolactone) monolith) osteochondral plug indicated for the repair of cartilage defects. The porous plug allowed for continual transport of bone marrow constituents from the subchondral layer to the cartilage defect site for a more effective repair of the area. Owida et al. [64] developed zonal-specific three-dimensional hybrid scaffolds and a simple method for biomimetic cartilage regeneration, which can induce the generation of zonal-specific cellular morphology and ECM composition within articular cartilage zones. Scaffolds were formed using polylactic acidnanofiber meshes with different alignments: aligned nanofibers for the superficial zone, randomly aligned fibers for the middle zone, and microchannels in hyaluronic acidhydrogel (Fig. 3.2).

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Figure 3.2  Schematic illustration of the assembled three separately cultured zonal scaffolds with corresponding thickness (a). An OCT image of the assembled 3D zonal scaffold showing the aligned nanofiber in superficial zone, random nanofiber in middle zone, and the vertical channels in deep zone (b). Scale bar is 250 μm [64].

McCorry et al. [65] developed a simplified enthesis model, which can be used to mimic the native enthesis morphology and serves as an ideal test platform via test collagen integration with decellularized bone. Injecting collagen into tubing loaded with decellularized bone plugs resulted in a scaffold with three regions: bone, bone–collagen, and collagen. Furthermore, collagen formation was directed in the axial direction using mechanical fixation at the bony ends. Tellado et al. [66] provided new insights into the combinational effects of scaffold internal architecture and growth factors; biphasic silk fibroin scaffolds with integrated anisotropic (tendon/ligament-like) and isotropic (bone/cartilage like) pore alignment, which presented a promising strategy for tendon/ligament-to-bone regeneration. Human primary adipose-derived mesenchymal stem cells were cultured on biphasic silk fibroin scaffolds with integrated anisotropic (tendon/ligament-like) and isotropic (bone/cartilage like) pore alignment. Furthermore, the scaffolds were functionalized with heparin and the ability to deliver transforming growth factor ß2 and growth/differentiation factor 5 was explored.

3.3.4  Scaffolds to promote vascularization In the ever advancing field of tissue engineering, vascularization of tissues is an important aspect that allows the creation of specialized tissues that are able to maintain growth, function, and viability. Vasculature has a vital role to play in creating and maintaining healthy tissues. As such, creating artificial tissues that are able to effectively mimic and integrate with native tissues is a key challenge with tissue engineering. Common approaches to achieve this include the use of cells, growth factors, ECM proteins, and biophysical stimuli [67].

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Bone grafts are crucial for the treatment of a number of conditions that represent a great global burden, including segmental bone defects caused by trauma, tumor excision, or chronic osteomyelitis to name a few. The need for bone constructs stems from the limited availability of donor tissues, which can be categorized as autografts, allografts, and xenografts, with the current gold standard treatment being autografts. However, these are limited in availability, prolong the operation times, and are often associated with donor site morbidity [68,69]. As an alternative, allografts are widely available. However, they place the patients at risk for infections and rejection by the immune system. More importantly, allografts demonstrate decreased integration with the host tissue when compared with autografts, with high failure rates of approximately 25% and reaching up to 60% in patients requiring large grafts. Allografts, as well as synthetic grafts, are associated with complications such as osteonecrosis, as these options lack an endogenous vascular network to facilitate host integration. The current focus looks at developing vascularized bone grafts to counter these limitations [68]. The aim of bone tissue engineering is to provide a bone environment rich in functional vascular networks to achieve efficient osseointegration and accelerate restoration of function after implantation. To attain both structural and vascular integration of the grafts, a large number of biomaterials, cells, and biological cues have been evaluated [70]. An example of an approach that has shown some measure of success in improving graft vascularization is the use of a polymethyl methacrylate (PMMA) cement block, which is placed on the bone defect and is surrounded with the adjacent soft tissues. The host slowly creates a membrane rich in vasculature around the block, which is then removed and replaced with an autograft. Bone repair is then prompted by host remodeling in this vascularized environment [70,71]. Another established approach is the use of synthetic fillers, which provide an osteoconductive scaffold into which bone-depositing cells migrate and deposit new bone tissue [70,72]. Although promising results have been obtained, it is also increasingly evident that new research should be aimed at hierarchical integration of bone and vascular devices to yield fully functional bone tissue engineering constructs with enhanced biological properties that can promote concurrent osteogenic and angiogenic growth and seamlessly assimilate with the host bone matrix and vasculature [70]. The materials selected as the scaffolds for tissue engineering need to be inherently biocompatible, biodegradable, and promoters of cell adhesion. The materials also need to be porous and mechanically stable and exhibit a 3D structure that can be obtained through facile manufacturing processes [73]. Various strategies have been attempted to enhance the establishment of vascular networks within engineered constructs for bone regeneration. These include (1) directing cell behavior through growth factor delivery, (2) using coculturing systems, (3) applying mechanical stimulation, (4) using biomaterials with appropriate properties, and (5) incorporating microfabrication techniques. The development and use of 3D printed scaffolds represents a huge opportunity for the world of tissue engineering. This methodology represents an accurate, in terms of design, and reproducible method for the fabrication of porous scaffolds, which facilitate cell attachment, vascularization, and tissue ingrowth. Development of a vascular network has been shown to be successful with porous, degradable scaffold sleeve designs with a lumen [69,74].

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With numerous established and emerging methods of scaffold fabrication available, the incorporation of vascular cells within the scaffolds provides an ideal opportunity to develop sufficient vasculature within the engineered tissue. The use of endothelial cells in coculture with other cell types can be used to achieve in vitro prevascularization and generate vascularization within the scaffold construct, and it has been reported that there is a mutual cell–cell interaction between ECs and osteoblast-like cells during osteogenesis. Cross-talk between human umbilical vein endothelial cells (HUVECs) and MSCs not only enhanced the osteogenic differentiation of MSCs but also increased the proliferation of differentiated MSCs [75,76]. In addition, cocultures of HUVECs and primary human osteoblasts (pOB) in 3D spheroids seeded into collagen gels demonstrated the formation of microcapillary-like structures resembling capillaries. It was found that during cell–cell contact between spheroids of HUVECs and pOB, an upregulation of VEGFR-2 was observed on the HUVECs, whereas a downregulation of VEGF production and an upregulation of alkaline phosphatase activity were observed in the pOB. This is an example of one of the first studies showing a bidirectional regulation of gene expression of HUVECs in coculture with pOB that required a direct contact with pOB to form capillary-like structures [77]. The choice of endothelial cell type for prevascularization of grafts is crucial. Mature ECs have been traditionally thought to be responsible for postnatal angiogenesis by which capillaries sprout from preexisting structures, until recent identification of peripheral blood-derived endothelial progenitor cells (PB-EPCs) by Asahara et al. [78]. These cells were found to be 10 times more proliferative than HUVECs. The use of human progenitor-derived endothelial cells (PDECs) in a 3D coculture with MSCs showed new osteoid formation as demonstrated by Guerrero et al. [79]. The parallel formation of a vascular network, surrounding the tissue when implanted in vivo, represents a promising approach for achieving vascularization [76,79]. The proangiogenic effect exerted by EPCs appears dependent on their ability to stimulate the growth of endothelial cells and interact with mature endothelial cells to support vascular anastomosis. Nevertheless, the mechanism by which the EPCs exert beneficial effects on endothelial cell growth is likely multifactorial and might include the transdifferentiation of subpopulations of EPCs into mature endothelial cells [68]. Vascularization is a key process in skin tissue engineering, which determines the biological function of artificial skin implants. Consequently, efficient vascularization strategies are a vital prerequisite for the safe clinical application of these implants. Current approaches include (1) modification of structural and physicochemical properties of dermal scaffolds, (2) biological scaffold activation with growth factor–releasing systems or gene vectors, and (3) generation of prevascularized skin substitutes by seeding scaffolds with vessel-forming cells [80]. Attempts are being made to incorporate vasculature into engineered skin although applications in humans are thus far limited. An example of vascular inclusion in tissue-engineered skin is provided by Tremblay et al. [81] where they incorporated keratinocytes, fibroblasts, and endothelial cells (ECs) in a collagen sponge implanted in mice. The implanted skin substitute then spontaneously formed a network of capillary-like structures (CLS) in vitro. After transplantation to nude mice, they demonstrated that CLS containing mouse blood were observed underneath the epidermis in the endothelialized reconstructed

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skin (ERS) in less than 4 days versus 14 days required to have the same observation in non-endothelialized reconstructed skin. This result was associated with inosculation of the CLS network with the host’s capillaries, rather than neovascularization. Another approach is tissue engineering of a vascular network in human dermoepidermal skin substitutes (DESS). Klar et al. [82] utilized adipose stromal vascular fraction (SVF)–derived endothelial cell population to tissue-engineered DESS containing a highly efficient capillary plexus. To develop vascular networks in vitro, they employed optimized 3D fibrin or collagen type I hydrogel systems and upon transplantation onto immune-deficient rats; these preformed vascular networks anastomosed to the recipient’s vasculature within only 4 days. As a consequence, the neoepidermis efficiently established tissue homeostasis; the dermis underwent almost no contraction and showed sustained epidermal coverage in vivo. They conclude that adipose-derived SVF represents a convenient source of endothelial cells and pericytes and when submerged within an appropriate 3D environment, these cells allow the in vitro prevascularization of human autologous dermoepidermal skin grafts.

3.3.5  Scaffolds from decellularized tissues A convenient alternative to manufactured scaffolds is the use of decellularized tissues to obtain ECM. The most obvious advantage of decellularized tissue is the resultant ECM is perfectly designed to host cells in a 3D environment in vitro. This ECM has the natural conformation and chemical composition of specific tissues, generating the specific cues that cells need to grow in a more native environment. The removal of cellular content and antigens from the tissue-derived scaffolds reduces foreign body reaction, inflammation, and potential immune rejection. A disadvantage of this approach is the possible structural alteration of ECM or loss of some important components in ECM because of overexposure to chemicals during the decellularization process [83,84]. The clinical use of decellularized scaffolds has been applied for blood vessels, cardiac valves, and renal bladders. The current applications may be limited to tissue-level and anatomically simple organs; despite this, they ultimately provide the foundation for future complex and functioning organ regeneration [85]. There are a variety of approaches for decellularizing tissues. Effective decellularization methods include chemical, enzymatic, physical, or combinational approaches. The working principle of these methods is that the cell membrane is disrupted, and the cellular contents are released and rinsed away [84]. One method is the use of sodium dodecyl sulfate (SDS) as described by Schaner et al. [86]. They used intact human greater saphenous vein specimens that were decellularized using sodium dodecyl sulfate and assessed their viability for use as scaffolds for vascular tissue engineering. By evaluating burst and suture-holding strength and handling and durability of decellularized vein in vivo, they determined that the vein rendered acellular with SDS has well-preserved extracellular matrix, basement membrane structure, and strength sufficient for vascular grafting [86]. SDS has been used in a number of studies as the preferred chemical agent for decellularization. The method described by Guler et al. [83] involves the use of SDS to decellularize porcine aorta. Dimethyl sulfoxide (DMSO) was used as a penetration enhancer in the

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decellularization process to enhance the penetration of SDS, consequently reducing the exposure time of SDS to treated tissues. They determined that the addition of DMSO resulted in the removal of 64.4% more DNA compared with SDS on its own over a 3 hours period. The inclusion of DMSO also resulted in denser and more organized collagen fibers [83]. Another approach to decellularization is described by Tapias and Ott [87]. Their approach involved the use of perfusion decellularization. This method uses perfusion via the intrinsic vascular network as the most efficient way to deliver decellularizing agents, even to thick tissues, as it greatly decreases the diffusion distance of the decellularizing agent while preserving the three-dimensional macro- and microarchitecture [87]. The work done by Quint et al. [88] demonstrates practical use of decellularized scaffolds. Their approach involved the use of tissue-engineered vessels (TEVs) grown from banked porcine smooth muscle cells that were allogeneic to the intended recipient, using a biomimetic perfusion system. The engineered vessels were then decellularized, leaving behind the mechanically robust extracellular matrix of the graft wall. The acellular grafts were then seeded with either endothelial progenitor cells (EPC) or endothelial cells (EC), which were derived from the intended recipient, on the graft lumen. The TEVs were then implanted as end-to-side grafts in the porcine carotid artery, which is a rigorous testbed because of its tendency for graft occlusion. The EPC- and EC-seeded TEV all remained patent for 30 days in their study, whereas the contralateral control vein grafts were patent in only 3/8 implants [88]. Deviating from animal sources of decellularized matrix, Gershlak [89] utilized decellularized plant tissue as a prevascularized scaffold for tissue engineering applications. Their study exploited the similarities between the vascular structure of plant and animal tissues and using perfusion-based decellularization modified for different plant species, provided different geometries of scaffolding. After decellularization, plant scaffolds remained patent and able to transport microparticles. Plant scaffolds were recellularized with human endothelial cells that colonized the inner surfaces of plant vasculature. Human mesenchymal stem cells and human pluripotent stem cell–derived cardiomyocytes adhered to the outer surfaces of plant scaffolds. Cardiomyocytes demonstrated contractile function and calcium handling capabilities over the course of 21 days [89]. Decellularized scaffolds have vast potential for the regeneration of either small sections of tissue or whole organs and have demonstrated regenerative capabilities in vivo and in vitro, suggesting their value in emerging treatment approaches. There are still limitations with the technique such as the achievement of fully functional organs that bear all the necessary native properties and the limitations of tissue-specific decellularization methods and the resultant variation in efficacy. A combination of multiple approaches, as demonstrated by Guler et al. [83], led to improved outcomes. With an ever increasing understanding of underlying mechanisms and development of new materials and techniques, the generation of engineered tissues and organs that are as good as “the real thing” becomes less of a possibility and more of a certainty. Recellularization of the decellularized scaffolds by corresponded cell types is an other big challenge. The dense ECM network prevents seeded cells infiltrating to

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the related layers, especially the media layer. Recellularization of right phenotype cells in right layers for multiple cellular tissue is even tough challenge. A few nice projects have been undertaken for thin tissue with single cell type. Chani et al. for example,recellularized decellularized rat kidney [90]. The pattern of distribution of the injected cells was organized into vascular structures within the glomerulus, similar to freshly isolated kidney. Dynamically and sequentially seeding with perfusing system is one of the feasible techniques to achieve high seeding efficiency.

3.4  Summary The characteristics of native tissue structures are ultimately determined by the type of GAGs, proteoglycans, and proteins present in the ECM environment, in addition to the arrangement of cell types present. Factors such as mechanical properties, cell layer arrangement, vasculature, and the presence of an epithelial layer can each influence how a tissue adapts to its function. To mimic these native tissue structures in vitro, tissue engineers have developed various scaffold designs with increasing complexity over recent years. These include hydrogel, porous and fibrous scaffolds, coculture and multilayered scaffolds, zonal structure scaffolds, as well as scaffolds to promote vasculature. Continued work in this area is likely to revolutionize the way in which we study and treat disease mechanisms in the future.

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[69] L. Nguyen, N. Annabi, M. Nikkhah, H. Bae, L. Binan, S. Park, Y. Kang, Y. Yang, A. Khademhosseini, Vascularized bone tissue engineering: approaches for potential improvement, Tissue Eng B Rev 18 (5) (2012) 363–382. [70] Á. Mercado-Pagán, A. Stahl, Y. Shanjani, Y. Yang, Vascularization in bone tissue engineering constructs, Ann Biomed Eng 43 (3) (2015) 718–729. [71] A. Masquelet, F. Fitoussi, T. Begue, G. Muller, Reconstruction of the long bones by the induced membrane and spongy autograft, Ann Chir Plast Esthet 45 (2000) 346–353. [72] J. Kirkpatrick, C. Cornell, B. Hoang, W. Hsu, T. Watson, W. Watters, C. Turkelson, J. Wies, S. Anderson, Bone void fillers, Am Acad Orthop Surg 18 (9) (2010) 576–579. [73] N. Thadavirul, P. Pavasant, P. Supaphol, Development of polycaprolactone porous scaffolds by combining solvent casting, particulate leaching, and polymer leaching techniques for bone tissue engineering, J Biomed Mater Res A 102 (10) (2014) 3379–92. [74] M. Wang, C. Vorwald, M. Dreher, E. Mott, M. Cheng, A. Cinar, H. Mehdizadeh, S. Somo, D. Dean, E. Brey, J. Fisher, Evaluating 3d-printed biomaterials as scaffolds for vascularized bone tissue engineering, Adv Mater 27 (1) (2014) 138–144. [75] Y. Liu, J. Chan, S. Teoh, Review of vascularised bone tissue-engineering strategies with a focus on co-culture systems, J Tissue Eng Regenerat Med 9 (2) (2012) 85–105. [76] N. Paschos, W. Brown, R. Eswaramoorthy, J. Hu, K. Athanasiou, Advances in tissue engineering through stem cell-based co-culture, J Tissue Eng Regenerat Med 9 (5) (2014) 488–503. [77] R. Unger, E. Dohle, C. Kirkpatrick, Improving vascularization of engineered bone through the generation of pro-angiogenic effects in co-culture systems, Adv Drug Deliv Rev 94 (2015) 116–125. [78] T. Asahara, T. Murohara, A. Sullivan, M. Silver, R. van der Zee, T. Li, B. Witzenbichler, G. Schatteman, J. Isner, Isolation of putative progenitor endothelial cells for angiogenesis, Science 275 (5302) (1997) 964–966. [79] J. Guerrero, S. Catros, S. Derkaoui, C. Lalande, R. Siadous, R. Bareille, N. Thébaud, L. Bordenave, O. Chassande, C. Le Visage, D. Letourneur, J. Amédée, Cell interactions between human progenitor-derived endothelial cells and human mesenchymal stem cells in a three-dimensional macroporous polysaccharide-based scaffold promote osteogenesis, Acta Biomater 9 (9) (2013) 8200–8213. [80] F. Frueh, M. Menger, N. Lindenblatt, P. Giovanoli, M. Laschke, Current and emerging vascularization strategies in skin tissue engineering, Crit Rev Biotechnol 37 (5) (2016) 613–625. [81] P. Tremblay, V. Hudon, F. Berthod, L. Germain, F. Auger, Inosculation of tissue-engineered capillaries with the host’s vasculature in a reconstructed skin transplanted on mice, Am J Transplant 5 (5) (2005) 1002–1010. [82] A. Klar, S. Güven, T. Biedermann, J. Luginbühl, S. Böttcher-Haberzeth, C. MeuliSimmen, M. Meuli, I. Martin, A. Scherberich, E. Reichmann, Tissue-engineered dermoepidermal skin grafts prevascularized with adipose-derived cells, Biomaterials 35 (19) (2014) 5065–5078. [83] S. Guler, H. Aydin, L. Lü, Y. Yang, Improvement of decellularization efficiency of porcine Aorta using dimethyl sulfoxide as a penetration enhancer, Artif Organs 42 (2) (2017) 219–230. [84] S. Patnaik, B. Wang, B. Weed, J. Wertheim, J. Liao, Chapter 3: Decellularized scaffolds: concepts, methodologies, and applications in cardiac tissue engineering and whole-organ regeneration, Tissue Regen (2014) 77–124. [85] Y. Yu, A. Alkhawaji, Y. Ding, J. Mei, Decellularized scaffolds in regenerative medicine, Oncotarget 7 (36) (2016).

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[86] P. Schaner, N. Martin, T. Tulenko, I. Shapiro, N. Tarola, R. Leichter, R. Carabasi, P. DiMuzio, Decellularized vein as a potential scaffold for vascular tissue engineering, J Vasc Surg 40 (1) (2004) 146–153. [87] L. Tapias, H. Ott, Decellularized scaffolds as a platform for bioengineered organs, Curr Opin Organ Transplant 19 (2) (2014) 145–152. [88] C. Quint, Y. Kondo, R. Manson, J. Lawson, A. Dardik, L. Niklason, Decellularized tissueengineered blood vessel as an arterial conduit, Proc Natl Acad Sci USA 108 (22) (2011) 9214–9219. [89] J. Gershlak, S. Hernandez, G. Fontana, L. Perreault, K. Hansen, S. Larson, B. Binder, D. Dolivo, T. Yang, T. Dominko, M. Rolle, P. Weathers, F. Medina-Bolivar, C. Cramer, W. Murphy, G. Gaudette, Crossing kingdoms: using decellularized plants as perfusable tissue engineering scaffolds, Biomaterials 125 (2017) 13–22. [90] B. Chani, V. Puri, R.C. Sobti, V. Jha, S. Puri, Decellularized scaffold of cryopreserved rat kidney retains its recellularization potential, PLoS One 12 (3) (March 7, 2017) e0173040.

Computational design of tissue engineering scaffolds

4

Esther Reina-Romo1,a, Ioannis Papantoniou2,3,a, Veerle Bloemen2,4, Liesbet Geris2,5,6 1University of Seville, Department of Mechanical Engineering and Manufacturing, Seville, Spain; 2Prometheus, LRD Division of Skeletal Tissue Engineering, KU Leuven, Leuven, Belgium; 3Skeletal Biology & Engineering Research Center, KU Leuven, Leuven, Belgium; 4Materials Technology TC, Campus Group T, KU Leuven, Leuven, Belgium; 5Biomechanics Research Unit, GIGA - In silico Medicine, Université de Liège, Liège, Belgium; 6Biomechanics Section, KU Leuven, Leuven, Belgium

4.1   Introduction Next to in vitro and in vivo models, a third model system is increasingly used in biomedical sciences, namely in silico models. In silico refers to silicium, the basic component of computer chips, and means the use of computer modeling and simulation in the broadest sense of the word. These models can be purely data driven (empirical, black box) or they can be based on already identified mechanisms (hypothesis driven, white box). The use of computational tools presents multiple advantages for aiding scaffold design by identifying suitable design options before laborious and costly experimental effort. Given recent advances in bioprinting and biomanufacturing technologies, which possess the required accuracy for producing scaffolds with the necessary morphometric properties, the use of in silico models becomes indispensable as a compass for rational production of tissue-engineered implants. In this chapter, we will present examples where computational models have successfully supported the design and production of scaffolds and fabrication technologies. In this chapter, we will follow the typical technological phases of the scaffold fabrication process [1–3]. • Step 1—preprocessing: design of the scaffold. This design will take into account structural, mechanical, degradation, and mass transport properties. • Step 2—the fabrication process. • Step 3—postprocessing: bioreactor culture.

We will limit our discussion in all steps to the questions for which in silico models have been or are being developed. We continue the chapter with a discussion on (multiobjective) optimization strategies as the need for these approaches becomes imperative because there is a dramatic increase in both computational and experimental a These

authors contributed equally.

Handbook of Tissue Engineering Scaffolds: Volume One. https://doi.org/10.1016/B978-0-08-102563-5.00004-6 Copyright © 2019 Elsevier Ltd. All rights reserved.

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capacity and throughput. Finally, we end with a glance toward the regulatory aspects related to the inclusion of in silico generated evidence in dossiers submitted to the regulatory agencies. Fig. 4.1 gives a schematic summary of this outline.

4.2  Preprocessing: design of the scaffold In this section, we present an overview of how in silico models have allowed the design and optimization of key scaffold design properties such as topology, mechanical properties, degradation, and mass transport (cf. Chapter 1 in this volume [112]). Topology, including pore shape, is a factor affecting tissue growth volume and nutrient transport. Mechanical properties affect scaffold deformation under load and, therefore, tissue stimulation and integration upon implantation. The degradability influences the release of bioactive constituents affecting the biologic state of cells surrounding scaffold struts. Finally, mass transport considerations (within scaffold struts and/or pores) are crucial for cell survival and efficient differentiation. Many of the aforementioned scaffold properties might have synergistic and/or antagonistic effects on cell function. Computational tools allow to decouple these effects and to contribute in improved understanding and hence rational use in scaffold design. Optimal combinations of the aforementioned might depend on tissue type and target organ.

4.2.1  Scaffold structural properties In silico models can help link observed tissue-scale dynamics with unknown cellular activity [6]. Initial work by Hollister et al. [7–11] focused on the design of bone scaffolds as an optimization problem to obtain a microstructure as similar as possible (in terms of mechanical properties, porosity, pore size, etc.) to that of the implanted region. For this purpose, homogenization theory was extensively applied in their studies. In other work, mechanical properties and permeability were homogenized over a representative element of a bioceramic scaffold microstructure [12]. The obtained results were corroborated by an experimental setup showing the potential of numerical tools for the characterization of scaffold properties. A series of experimental studies in fact showed that pore geometry influences stem cell behavior [13] through various mechanisms [14]. However, the underlying mechanism regarding geometric regulation of collective cell crowding and neotissue formation was the impact of curvature [15] and specifically of its interplay with linear tension [16]. In addition, the cellular stress dictated by the scaffold geometry through local curvature translates in the secretion of prestressed extracellular matrix architectures following stress patterns [17]. Hence, instead of a random trial and error approach, various studies focused on a design approach for scaffold bioprinting whereby fundamental properties were investigated rather than trivial shapes and sizes. First single pore shapes models were used to probe the rate of infilling by cells in convex and concave surfaces [6] for simple geometries, while later on Guyot et al. [18] developed a curvature-driven computational model to capture neotissue growth kinetics across scaffold pores of increased complexity able to simulate up until complete infilling of all the pores (Fig. 4.2).

Postprocessing bioreactor

0

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Flow velocity, u (m/s)

0,192

0,153

Flow (1ml/min)

Fabrication

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Computational design of tissue engineering scaffolds

Preprocessing design

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Figure 4.1  Schematic overview of this chapter, illustrating the different steps discussed in the text. (1) Preprocessing: design of scaffold geometries [4]. (2) Fabrication: simulation of mechanical environment during 3D extrusion-based bioprinting (unpublished). (3) Postprocessing: bioreactor culture showing quantification of local oxygen concentration and fluid flow velocity [5].

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Figure 4.2  Curvature-driven neotissue (cell + the ECM they produce) growth dynamics in complex 3D scaffolds. (a) Simulation of neotissue growth in 3D-printed titanium scaffolds with three different unit cell geometries (triangle, square, hexagon). Calibration with experimental findings by comparing surface fraction of the unit cell in simulations and experiments. Calibration point used is hexagon (iteration 7000 = 7 days in culture). Right: visual comparison of obtained neotissue geometry (left) and experimentally observed neotissue formation (right). (b) Simulation of complex diamond-shaped unit cell, demonstrating the capacity of the in silico model to simulate neotissue growth up to complete filling. Adapted with permission from Springer from Y. Guyot, I. Papantoniou, Y.C. Chai, S. Van Bael, J. Schrooten, L. Geris, A computational model for cell/ECM growth on 3D surfaces using the level set method: a bone tissue engineering case study, Biomech Model Mechanobiol 13 (6) (November 2014) 1361–1371, https://doi.org/10.1007/s10237-014-0577-5. Epub 2014 Apr 3.

4.2.2  Mechanical properties A major role of scaffolds is to provide structural support to the cultured cells while tissues are engineered in vitro. In addition, scaffold stiffness regulates tissue regeneration upon implantation. Therefore, the mechanical properties of the produced scaffolds have long been a focus of computational investigation (cf. the aforementioned work by Hollister, Fernandes et al.). When assessing whole-implant scale scaffolds, computational models can define mechanical properties that will be comparable with those encountered at the implantation site aiding incorporation and host integration [19,20]. Additionally, the mechanical loading encountered after implantation can be coupled, for instance, to vascular development with bone tissue formation [21]. Fluid–structure interaction (FSI) approaches have been applied to understand the role of scaffold stiffness and architecture on the wall shear stress distribution. In fact, McCoy [22] determined that the applied flow rate dominated the mechanical stimulation when compared with the pore size in collagen–GAG scaffolds. More recently, Zhao et al. [23] applied this method to investigate the role of scaffold geometry

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(architecture, pore size, and porosity) on pore wall shear stress (WSS) under a range of different loading scenarios (being fluid perfusion, mechanical compression, and a combination of perfusion and compression), finding that scaffold geometry (spherical and cubical pores), and in particular the pore size, has a significant influence on the stimulation within the scaffolds. In addition, they concluded that the combination of loading conditions would allow amplifying these wall shear stresses. Fluid–structure interaction has been used to simulate how the fluid movement deforms the cell body, modeling the cell as a solid in many different applications of tissue engineering. Vaughan [24] developed a fluid–structure interaction model to characterize the deformation of integrin and primary cilia-based mechanosensors in bone cells under fluid flow stimulation. Recently, immersed boundary models were used to quantify compressive and shear stresses developed over deformable virtual cells of various shapes and scaffold locations when exposed in fluidic environment [5].

4.2.3  Modeling scaffold degradability The ability to quantitatively decipher physical, chemical, and biological phenomena involved in the controlled release of ions or molecules [25,26] requires the use of in silico models. Concerning the development of optimized degradable scaffolds, the importance of such models lies in their relevance during the designing stage as well as the experimental verification of degradation and release mechanism(s). However, it is unlikely that there will be one single in silico model that will be able to describe any type of release of ions or molecules from biomaterials. Scaffold degradation should give way gradually to new native tissue, leading to complete scaffold disappearance. Scaffold degradation usually takes place by chemical pathways (hydrolysis) in the case of polymeric scaffolds and has been a focus of numerical analysis studies by, e.g., Adachi et al. [7,27]. Thorough understanding of hydrolysis kinetics can control the rate of scaffold mass loss, enabling the design of polymer biomaterials for tailored applications. Pioneering studies on species (drug) release from scaffold–polymer systems has been extensively carried out [25,28]. However, given the particularities of bioprinting and tissue engineering, this domain is still not adequately studied. Over the past few years, a few lattice-based three-dimensional (3D) in silico models have been proposed to study the in vivo bone formation process in porous biodegradable CaP scaffolds [29,30]. Byrne et al. [29] developed an in silico model of in vivo tissue differentiation and bone regeneration in a degrading scaffold as a function of porosity, Young’s modulus, and dissolution rate. Sun et al. [30] proposed a multiscale model of a biodegradable porous calcium phosphate (CaP) scaffold to examine the effects of pore size and porosity on bone formation and angiogenesis. However, the aforementioned models did not capture the actual geometry of the degrading CaP scaffolds, which was demonstrated by Manhas et al. [31]. Lastly, a series of investigations reported on the development of in silico models for bioglass scaffolds and their degradation properties for bone tissue engineering [32], whereas agent-based models were used to capture degradation in function to invading vascularization [33]. Even more intricate models capturing geometric complexities such as the design of biodegradable interbody fusion cages have been recently carried out [34]. These models

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had phenomenological description of the degradation process and were interested in the degradation products themselves but also in the changing stiffness and porosity of the scaffold.

4.2.4  Mass transport Before the printing process, the performance of implants can already be verified by simulating the transport of oxygen, nutrients, and waste products in the printed materials. Shipley et al. [35] investigated the design of an extrusion-based printed construct (gel + cells) in terms of printed geometrical configuration (strand thickness and density) and speed of perfusion of medium throughout the construct in a perfusion bioreactor system. Besides transport of nutrients and waste products, also other transport can be simulated. For example, Carlier et al. [36] demonstrated using a previously established computational model of bone regeneration [37,38], spatially patterned constructs enhance bone regeneration compared with constructs with a uniform cell distribution. The model accounts for cells, their extracellular matrix as well as the presence (and diffusion) of growth factors. The models allow testing the printing pattern of complex implants before printing during the design process in bioprinting.

4.3  The fabrication process Scaffolds have been fabricated using various fabrication methodologies ranging from decellularization techniques to the use of additive manufacturing as discussed in Chapter 6 in this volume [113]. The choice of the production process determines the range of materials that can be used and the scaffold design characteristics that can be fabricated. Moreover, when materials and cells are combined in the same fabrication process, as is the case in the field of bioprinting, an extra level of complexity is added to the design of the production process to obtain a viable construct with the desired scaffold properties. To ensure that the fabricated scaffolds meet the requirements linked to architecture and biocompatibility, the production process should be well designed and robust. In conventional fabrication methods, the degree of control over the micro- and nanostructure is limited and therefore difficult to predict. Although additive manufacturing techniques allow for a precise spatiotemporal material deposition, the shape fidelity is often decreased by the fabrication process. Optimization is needed but challenging and tedious because of the multifactorial nature of the process, and a trial-and-error approach is currently mostly used in which one parameter is changed and the effect of the change is investigated experimentally [39–41]. To overcome these hurdles, computational modeling can offer an important tool that allows for a more efficient screening of the influence of process parameters on the robustness of the fabrication process and the quality of the outcome. In the following paragraphs, a few examples are given of how computational modeling can play a role in the design of the fabrication process.

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4.3.1  Shape fidelity in function of the fabrication process The theoretical design of a scaffold often differs from the actual architecture because of the influence of the fabrication process on parameters such as layer height and wall thickness, thereby reducing the shape fidelity of the actual object. Although great advances in scaffold design have been obtained by using computational modeling for optimizing the scaffold’s topology, surface, and size, models describing how the fabrication process influences these parameters are still scarce. Castilho et al. suggested that it might be considered to add a correction factor to the computational model describing the design of the scaffold to correct for the observed deviations introduced by the setup of the printing process [42]. In an attempt to predict the shape of a printed object taking into account the effect of its printing process, Suntornnond et al. developed an in silico model that describes the resolution of the printed width of a continuous hydrogel line as a function of nozzle size, pressure, and printing speed [43]. In addition, the behavior of materials during printing is dependent on their rheological profile and the design of the printing setup. Recently, a model has been proposed to describe the printability of inks in function of the design of the needles used in a pressure-driven, shear-thinning, extrusion-based printing process [44], and the influence of different printing parameters on viscoelastic stresses within the inks and print fidelity has been simulated in a computational fluid dynamics analysis [45]. To analyze the shape of the hydrogel extruded, Lee and Yeong [46] developed a model for a time–pressure extrusion-based biodispensing system (Fig. 4.3).

4.3.2  Biocompatibility of the fabrication process conditions In bioprinting strategies, it is key to understand, characterize, and select the optimal process parameters needed to produce a cell-based construct that guides tissue development. These parameters will depend on the fabrication technology and the formulation of the bioinks used. The ideal bioink process interplay should ensure printability and biocompatibility at all phases of the bioprinting process and should lead to mechanical integrity and structural stability of the construct [47,110]. Different additive manufacturing technologies exist to fabricate scaffolds or cellbased constructs, and each setup has its own properties that can influence the production process as discussed in other chapters. Several computational models have been developed to gain insight in how the printed object is affected by the specific system specifications and help in determining the window in which the bioinks are printable and ensure cell survival (Fig. 4.2). Evaluating the flow forces acting onto cell–material mixtures for different setups, for instance, is difficult to observe with biological analyses only and requires a description of the physics related to the fabrication process to quantify said forces. Numerical techniques have the potential to predict and mimic the mechanochemical microenvironments of cells during the bioprinting process under various protocols as well as to optimize process parameters. Both classical theoretical calculations and finite element analyses may be applied. Through these in silico models, more insights can be obtained into construct properties at the different steps of the bioprinting process [2].

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Figure 4.3  Predicting bioprintability of nanofibrillar inks using computer simulations of the 3D bioprinting process. (A) Comparison of experimental observations and model simulations for printed line height (a) and printed line width (b) for two different nanofibrillar bioinks. (B) Simulated distribution of viscoelastic stresses in the printed ink incurred during the printing process for two different nanofibrillar bioinks. Adapted from J. Gohl, et al., Simulations of 3D bioprinting: predicting bioprintability of nanofibrillar inks, Biofabrication 10 (3) (2018) 034105 under the CC BY license.

To predict the shape of the scaffold and cell survival in inkjet printing, it is important to understand the droplet ejection in thermal inkjet printheads [48]. Pepper et al. modeled, both analytically and by means of FE methods, the cell settling effects in inkjet printing [49,50]. Tirella et al. used a finite element model to investigate the role of the stiffness of the deposition substrate during droplet impact during inkjet printing process [51]. For laser-induced forward transfer (LIFT) printing, Mezel [52] presented a 2D axisymmetric model to analyze jet formation. This analysis contributes to the understanding of the ejection process and aims to reduce the biological damage during printing. In extrusion-based bioprinting, the compromise between printability and biocompatibility may be achieved by manipulating the temperature, the geometry of the dispensing setups, the dispensing pressure, shear stress, and the bioink concentration

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among others [53]. The flow rate has a direct effect on the pore size and porosity of the scaffolds formed and therefore on its mechanical and biological properties. To analyze the effect of the nozzle geometry on the flow rate, non-Newtonian flow is considered. The flow rate in a cylindrical needle can be expressed as:

Q=

πR3 n 1 + 3n

[

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]1/n



(4.1)

where R and L are the nozzle length and radius, K is the consistency index, the constant n is the power law index, and ΔP is the pressure drop in the needle. The flow rate in a tapered needle is given by [54]:

[ ]1/n πD3i D3o 3n Δ P (Di − Do ) Q= ( ) 3n 32 4KL D3n i − Do

(4.2)

where Di and Do are the entrance and exit diameters of the tapered nozzle, respectively. Therefore, according to these equations, the flow rates in the tapered nozzles are much higher under the same pressure conditions, owing to the smaller diameter size at the exit. During the extrusion-based bioprinting process, cells are subjected to shear stresses. It is well known that higher shear stresses lead to lower cell viability [53,55]. Blaeser et al. [56] found that shear stress should be controlled within 5 kPa to obtain more than 90% living cells for mouse fibroblasts in a valve-based jet printing process. In fact, the percentage of cell damage (I) may be directly related to the shear stress cell experience (τ) through the very simple and well-established power law [57,58].

I ( % ) = Cta τb

(4.3)

where t is the exposure time. C, a, and b are constants for a given type of cells. To reduce cell damage due to shear stress, instead of directly printing cells loaded within the biomaterial, they may be encapsulated in spheroids [59]. The level of shear stress is directly related to different bioprinting parameters, such as viscosity of the bioink, pressure, or nozzle geometry [55,56]. The shear stress is related to the viscosity (μ) through the shear rate ( γ˙ ):

τ = μ˙γ = K˙γ n

(4.4)

where K is the consistency index and the constant n is the power law index. A sudden decrease of shear rates during deposition may cause an increase in viscosity, resulting in a high printing fidelity and higher cell viability. The relationship between the maximum shear stress in the wall and pressure drop is given by Ref. [55]:

τmax =

(

n 3n + 1

)n

D 4

Δ P

(4.5)

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where D is the nozzle diameter. In addition, the shear stresses increase with the nozzle radius, from zero at the needle center to its maximum value in the wall. Thus, at equivalent flow rates, shear stress and consequently cell damage in a tapered needle is lower than in a cylindrical one owing to its lower value of exit radius. This has been already analyzed both in vivo and in silico [54]. To favor cell survival in extrusion-based bioprinting, the dispensing pressure must be maintained as low as possible. A higher dispensing pressure can allow ejecting highly viscous bioinks, but this could increase the shear stress, which reduces cell viability (Eq. 4.3). Billiet et al. [53] have compared cell viabilities under different dispensing pressures and needle geometries. They found that at low inlet pressures (high passage time), tapered needles are preferred over cylindrical ones. At high inlet pressure (low passage time), substantially higher shear stresses are induced and a higher viability level is observed for the cylindrical type. Nair et al. [55] characterized the viability of endothelial cells during extrusion-based printing, and the results indicated that dispensing pressure had a more significant effect on cell viability than the nozzle diameter. However, under the same pressure conditions, the level of force cells experienced in tapered nozzles is much higher than in cylindrical nozzles owing to its much larger surface at the entrance (pressure = force/area). This may cause cells to be harmed and even die.

4.3.3  Biological functionality after the fabrication process After cell printing, three possible outcomes may occur: (1) cell survival with desired phenotype, (2) cell survival but cells become quiescent and they may recover and differentiate into diverse specialized cell types or die; and (3) immediate necrosis due to high shear stress, clotting in the nozzle or no tissue printed. Computer models can help to assess whether tissues are able to function as intended after the fabrication process. For the particular case of extrusion bioprinting, complex in silico modeling via cellular particle dynamics (CPD) simulations can be used to predict postprinting structure formation even in the case of volume changing bioink units [60,61]. A method based on kinetic Monte Carlo simulations has also been used to describe the shape evolution of multicellular systems postbioprinting [62–69]. Yang et al. [70] studied the morphological development of the printed bioconstructs during fusion by means of an in silico model based on the phase field formulation.

4.4  Postprocessing: bioreactor culture To deal with the increasing mass transport requirements of growing engineered tissues in vitro, the majority of scaffolds will need to be coupled to bioreactor systems for stem cell growth and differentiation. This requires the use of in silico models of higher complexity to decipher the increased complexity posed by this interplay between the dynamic culture environment and scaffold properties [71].

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4.4.1  Incorporating the neotissue domain Initially, computational fluid dynamics (CFD) simulations were developed and used to characterize flow patterns in (mostly empty) scaffolds to quantify the local flow-induced shear stresses acting on the attached cells [72–76]. For scaffolds with ordered pores structures, Truscello et al. [77] showed that the permeability can be defined accurately through CFD approaches. These computational studies supported initial experimental tissue engineering studies where cells were attached on scaffold struts and wall shear stresses experienced by the cells were important in determining their differentiation [22,78–80]. However, during culture, cells and the extracellular matrix they produce (together indicated with the term “neotissue”) eventually fill the scaffold (as characterized, for instance, in Ref. [81]), which will significantly affect the flow profile. This means that modeling empty scaffolds is inadequate for the determination of the true mechanical environment that seeded cells will experience over culture time. Initial efforts to address incorporate neotissue growth were undertaken; however, the neotissue volume was modeled as an impermeable structure leading to overestimated surface shear stresses [82,90]. Because the neotissue is a permeable structure permitting flow even at complete filling of the scaffold [83], it is important to consider the fluid velocity field developed within the neotissue to correctly determine the gradients developed within it. More recent studies have managed to carry out CFD analysis, while neotissue growth is occurred by using the level set method coupled to the Brinkman equation [84]. In addition, the neotissue growth kinetics were also coupled to shear stresses experienced by the cells, which were influenced in a dose-dependent manner [85]. Along these lines, Williams et al. used a lattice Boltzman model for quantifying the fluid dynamics in their system and they were able to monitor time-dependent mineralization of neotissue during growth [86].

4.4.2  Multiphysics models for scaffolds in bioreactors Deciphering the microenvironment that defines stem cell survival and fate is a daunting task because a multitude of factors need to be quantified instantaneously. To date, very few experimental studies report on important environmental cues such as dissolved oxygen tension, glucose, lactate, and pH all impacting stem cell state and neotissue properties. For instance, very low glucose and oxygen concentrations in the system could lead to cell death, whereas an important amount of lactate will decrease the medium pH and inhibit cell proliferation capacity [87]. In addition, pH can play a crucial role in bone tissue engineering applications as human MSC osteogenic differentiation has been seen to be inhibited in specific pH ranges [88]. Despite the importance of these physicochemical factors on the outcome of the bioreactor process, their local quantification in growing neotissues (through, e.g., sensors) remains problematic, thereby severely limiting the product quality control and the translation to clinical practice. Limited computational studies have attempted to model the (multiparametric) physicochemical environment experienced by growing cells within a bioreactor setup (for instance, [89–91] and [4]). In those studies, the authors introduced oxygen, glucose, and lactate with the help of diffusion–convection–reaction equations

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to determine the local concentration of these metabolic species in the context of bioreactor processes geared toward in vitro cartilage or neotissue growth. In addition, a model for coupling mechanical properties of struts and mass transport was also recently published [8]. However, certain limitations were present in terms of confluence levels that the growing domain could reach as well as in terms of the global (length) scale that could be modeled. This is a broad field where computational studies are required for mapping dynamic and time evolving biological systems from the niche to the whole-implant scale.

4.5  Discussion 4.5.1  Multiparametric optimization The determination of the optimal combination of the aforementioned scaffold and process properties is a daunting task. It is difficult to define what constitutes an optimal solution to a problem as there may be multiple, conflicting objectives. Hence the use of multiobjective optimization methods (MOOs) can allow to identify optimal trade-off between “costs” (e.g., cost of materials, time, and negative side effects) and “rewards” (e.g., successful neotissue growth). Furthermore, design of experiments (DoE) is an essential tool for quality by design (QbD) that allows the systematic and parallel investigation of multiple process parameters on scaffold properties and printed cell characteristics. By using this method, a ranking of the process parameters is achieved in terms of the most influential parameters in light of the target output panel. This method takes into account the interaction of parameter interrelationship, using linear regression and analysis of variance (ANOVA) in silico models [92]. Recent work by Ruiter, Nazir, Desai, and their respective coworkers focused on finding optimal electrospinning process parameters (polymer composition, dispensing distance, voltage, and flow rate), which significantly influenced the fiber diameter, morphologies, and bead distribution observed in electrospinning of poly-D-L-lactic acid (PDLLA) fibers [93–95]. Similarly, few DoE studies exist for melt extrusion bioprinting where temperature, pressure, and nozzle diameter have been optimized for scaffold compression strength [96]. Ravi et al. designed a modular 3D printing setup combining different fabrication technologies and they used a DoE to characterize the setup and to assess the relation between fabrication process parameters and their effect on printed scaffold characteristics [97]. For scaffolds cultured in bioreactors, DoEs have been used to investigate optimal seeding density and fluid flow to produce scaffolds with optimal presence of neotissue [80]. As the multivariable complexity of new processes and materials increases, the task of identifying the best combination of process settings to achieve output material property targets becomes complex and time-consuming and DoE still requires considerable experimentation. High-throughput experimental setups can be developed to assess certain aspects of scaffold (printer) design such as the case for surface topography in the TopoChip setup [111]. These data sets need to be handled using appropriate algorithms from the big data and -omics domain. Alternatively, advanced optimization methods capable of making better use of less data are required. Topological

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optimization algorithms have been recently published [98] together with mechanobiology-driven algorithms for bone tissue engineering applications [99]. Relevant blackbox (i.e., data-driven or empirical) optimization methods include genetic algorithms, trust-region methods, and Bayesian optimization [100–104]. Black-box methods give flexibility with respect to the black-box contents. If there are no restrictions on the black box, the optimization methods are equally relevant to in vitro experiments as legacy computer code. MOO methods explicitly find the trade-offs between conflicting objectives [105–107].

4.5.2   Future prospects In silico models are increasingly present in the life cycle of medicinal products, not only in the design phase but also in the development phase, the clinical trial phase as well as for postmarketing surveillance purposes. To build model credibility, the so-called VVUQ (verification–validation–uncertainty quantification) needs to be established. Verification answers the question whether the computed results correspond to the mathematical equations. Validation answers the questions whether the computed results correspond to physical reality. This needs to be complemented with uncertainty quantification, in which the effect of the uncertainties in the model assumptions and parameters on the simulation outcome is determined. For medical devices, reporting guidelines have been established by the FDA for inclusion of in silico evidence in regulatory dossiers (https://www.fda.gov/downloads/ MedicalDevices/DeviceRegulationandGuidance/GuidanceDocuments/UCM381813. pdf). Furthermore, in collaboration with industry and academia, a clear procedure has been established to perform verification and validation of the developed in silico models that are (part of) a medical device or have delivered digital evidence that was important in the R&D process of a medical device presented in a regulatory dossier [108,109]. Currently, ASME and its partners are also working on a VVUQ standard specifically dedicated to advanced (including additive) manufacturing (https://cstools. asme.org/csconnect/CommitteePages.cfm?Committee=101978604). These focus on development of standards in a clear sign of maturity of the in silico technology. Adding the biological aspects into the models moves the application from a medical device toward an advanced therapeutic medicinal product (ATMP), which brings along additional challenges in terms of model establishment and validation. Nevertheless, given the recent advances in fabrication technologies and the increasing demands on the complexity of the applications, computational models will become indispensable for rational production of tissue-engineered implants.

Acknowledgments Research was funded by the Research Foundation Flanders (FWO Vlaanderen; I.P: 12O7916N), the Fund for National Research (FNRS; T.0256.16), and the European Research Council under the European Union’s Horizon 2020 framework program ERC/CoG 772418 (L.G.). This work is supported by Regenerative Medicine Crossing Borders (www.regmedxb.com), powered by EWI-Vlaanderen.

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The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. This work is part of Prometheus, the KU Leuven R&D division for skeletal tissue engineering (http://www.kuleuven.be/prometheus).

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[85] Y. Guyot, I. Papantoniou, F.P. Luyten, L. Geris, Coupling curvature-dependent and shear stress-stimulated neotissue growth in dynamic bioreactor cultures: a 3D computational model of a complete scaffold, Biomech Model Mechanobiol 15 (2016) 169. https://doi. org/10.1007/s10237-015-0753-2. [86] C. Williams, O.E. Kadri, R.S. Voronov, V.I. Sikavitsas, Time-dependent shear stress distributions during extended flow perfusion culture of bone tissue engineered constructs, Fluid 3 (2) (2018) 25, https://doi.org/10.3390/fluids3020025. [87] K. Wuertz, K. Godburn, J.C. Iatridis, MSC response to pH levels found in degenerating intervertebral discs, Biochem Biophys Res Commun 379 (2009) 824–829. [88] L.E. Monfoulet, P. Becquart, D. Marchat, K. Vandamme, M. Bourguignon, E. Pacard, et al., The pH in the microenvironment of human mesenchymal stem cells is a critical factor for optimal osteogenesis in tissue-engineered constructs, Tissue Eng 20 (2014) 1827–1840. [89] M.S. Hossain, D.J. Bergstrom, X.B. Chen, Prediction of cell growth rate over scaffold strands inside a perfusion bioreactor, Biomech Model Mechanobiol 14 (2) (2014) 333–344, https://doi.org/10.1007/s10237-014-0606-4. [90] M.S. Hossain, D.J. Bergstrom, X.B. Chen, Modelling and simulation of the chondrocyte cell growth, glucose consumption and lactate production within a porous tissue scaffold inside a perfusion bioreacto, Biotechnol Rep 5 (2015) 55–62. [91]  M.M. Nava, M.T. Raimondi, R. Pietrabissa, A multiphysics 3D model of tissue growth under interstitial perfusion in a tissue-engineering bioreactor, Biomech Model Mechanobiol 12 (6) (November 2013) 1169–1179, https://doi.org/10.1007/ s10237-013-0473-4. [92] R.A. Fisher, The arrangement of field experiments, J Ministry Agric Great Britain 33 (1926) 503–513. [93] K. Desai, C. Sung, DOE optimization and phase morphology of electrospun nanofibers of PANI/PMMA blends, NSTI Nanotechnol 3 (2004) 429–432. [94] A. Nazir, N. Khenoussi, L. Schacher, T. Hussain, D. Adolphe, A.H. Hekmati, Using the Taguchi method to investigate the effect of different parameters on mean diameter and variation in PA-6 nanofibres produced by needleless electrospinning, RSC Adv 5 (2015) 76892–76897. [95] F.A.A. Ruiter, C. Alexander, F.R.A.J. Rose, J.I. Segal, A design of experiments approach to identify the influencing parameters that determine poly-D,L-lactic acid (PDLLA) electrospun scaffold morphologies, Biomed Mater 12 (5) (September 25, 2017) 055009, https://doi.org/10.1088/1748-605X/aa7b54. [96] P. Sheshadri, R.A. Shirwaiker, Characterization of material–process–structure interactions in the 3D bioplotting of polycaprolactone, 3D Print Addit Manuf 2 (1) (2015). https://doi.org/10.1089/3dp.2014.0025. [97] P. Ravi, P.S. Shiakolas, J.C. Oberg, S. Faizee, A.K. Batra, On the development of a modular 3D bioprinter for research in biomedical device fabrication, in: ASME International Mechanical Engineering Congress and Exposition, vol. 2A, ASME, 2015, https://doi. org/10.1115/IMECE2015-51555. Advanced Manufacturing:V02AT02A059. [98] H.A. Almeida, P.J. Bártolo, Topological optimisation of scaffolds for tissue engineering, Procedia Eng. 59 (2013) 298–306. [99] A. Boccaccio, A.E. Uva, M. Fiorentino, L. Lamberti, G. Monno, A mechanobiology-based algorithm to optimize the microstructure geometry of bone tissue scaffolds, Int J Biol Sci 12 (1) (2016). 26722213. [100] F. Boukouvala, M.G. Ierapetritou, Surrogate-based optimization of expensive flowsheet modelling for continuous pharmaceutical manufacturing, J Pharm Innov 8 (2) (2013) 131–145.

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[101] H.M. Chi, H. Moskowitz, O.K. Ersoy, K. Altinkemer, P.F. Gavin, B.E. Huff, B.A. Olsen, Machine learning and genetic algorithms in pharmaceutical development and manufacturing processes, Decis Support Syst 48 (1) (2009) 69–80. ISSN 0167-9236. https://doi. org/10.1016/j.dss.2009.06.010. [102] H.J. Kushner, A new method of locating the maximum point of an arbitrary multipeak curve in the presence of noise, J Basic Eng 86 (1) (1964) 97–106. [103] J. Mockus, Bayesian Approach to Global Optimization: Theory and Applications, Kluwer Academic Publishers, 1989. [104] J. Nogueira, et al., Unscented Bayesian Optimization for Safe Robot Grasping, IROS, 2016. [105] J.R. Banga, Optimization in computational systems biology, BMC Syst Biol 2 (1) (2008) 47. [106] F. Boukouvala, et al., Global optimization advances in mixed-integer nonlinear programming, MINLP, and constrained derivative-free optimization, CDFO, Eur J Oper Res 252 (2016) 701–727. [107] C.-L. Hwang, A.S.M. Masud, Multiple Objective Decision Making – Methods and Applications: A State-of-the-Art Survey, Springer, 1979. [108] T.M. Morrison, M.L. Dreher, S. Nagaraja, L.M. Angelone, W. Kainz, The role of computational modeling and simulation in the total product life cycle of peripheral vascular devices, J Med Dev Trans ASME 11 (2) (2017) 024503. [109] T.M. Morrison, P. Pathmanathan, M. Adwan, E. Margerrison, Advancing regulatory science with computational modeling for medical devices at the FDA’s office of science and engineering laboratories, Front Med 5 (2018) 241, https://doi.org/10.3389/ fmed.2018.00241. [110] S. Kyle, Z.M. Jessop, S.P. Tarassoli, A. Al-Sabah, I.S. Whitaker, Assessing printability of bioinks. (Chapter 9), in: 3D Bioprinting for Reconstructive Surgery, 2017, pp. 173–189. [111]  H.V. Unadkat, M. Hulsman, K. Cornelissen, B.J. Papenburg, R.K. Truckenmüller, A.E. Carpenter, M. Wessling, G.F. Post, M. Uetz, M.J. Reinders, D. Stamatialis, C.A. van Blitterswijk, J. de Boer, An algorithm-based topographical biomaterials library to instruct cell fate, Proc Natl Acad Sci USA 108 (40) (2017) 16565–16570. [112]  F. Sefat, A. Atala, M. Mozafari, An introduction to tissue engineering scaffold. (Chapter 1), in: F. Sefat, A. Atala, M. Mozafari (Eds.), The Handbook for Tissue Engineering Scaffolds, Elsevier, 2019. [113] J.P. St-Pierre, Fabrication techniques of tissue engineering scaffolds. (Chapter 6), in: F. Sefat, A. Atala, M. Mozafari (Eds.), The Handbook for Tissue Engineering Scaffolds, Elsevier, 2019.

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Poornima Ramburrun, Sunaina Indermun, Mershen Govender, Pradeep Kumar, Lisa C. du Toit, Yahya E. Choonara, Viness Pillay Wits Advanced Drug Delivery Platform Research Unit, Department of Pharmacy and Pharmacology, School of Therapeutic Sciences, Faculty of Health Sciences, University of the Witwatersrand, Johannesburg, South Africa 

5.1   Introduction Biomaterials, as defined by the American National Institute of Health and Nature, refers to any material that is of natural or synthetic origin or a combination thereof, alive or lifeless and consisting of multiple components with the ability to interact with biological systems to augment or replace partially or totally any natural function, tissue, or organ of the body for any length of time [1]. The earliest record of materials implemented by mankind for the repair and replacement of damaged or missing tissues from the human body included animal parts, nacre, and wood. This later evolved to include the use of metals and ceramics for basic repair of bone injuries (Fig. 5.1). The early twentieth century presented the development of new materials that replaced its ancient counterparts and consisted of an array of naturally derived and synthetic polymers, metal alloys, and ceramics engineered for enhanced reproducibility, performance, and functionality [3]. Thus, this chapter provides a general outline of the biomaterial classifications and an overview of material criterion for hard and soft tissue engineering applications with mention of common and emerging materials and composites for tissue engineering applications. The research evolution of scaffold materials in terms of different generations is discussed with indication of specific tissue engineering goals associated with the research efforts of each biomaterial generation.

5.1.1  Types of biomaterials The repair of damaged tissues using scaffold biomaterials that integrate cells from the body to initiate and guide the growth of new tissues is the objective of tissue engineering. Scaffolds serve as structural support and templates for tissue regeneration applications and are composed of a variety of synthetic or natural polymer materials or combinations thereof consisting of two or more material types from either class (Fig. 5.2). Extensive research is focused on the development of composite materials to compensate for disadvantages associated with individual materials to enhance biocompatibility, biodegradability, and biomimetic functions [4]. Handbook of Tissue Engineering Scaffolds: Volume One. https://doi.org/10.1016/B978-0-08-102563-5.00005-8 Copyright © 2019 Elsevier Ltd. All rights reserved.

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Development of decellularized extracellular matrix biological materials

2000

2010-present

Electrospun nanofibrous materials and hydrogels for cell and bioactive encapsulation for physical, chemical and therapeutic cues

Figure 5.1  Timeline representation of biomaterial use through the ages. Adapted from K. Sadtler, A. Singh, M.T. Wolf, X. Wang, D.M. Pardoll, J.H. Elisseeff, Design, clinical translation and immunological response of biomaterials in regenerative medicine, Nat Rev Mater 1 (7) (2016) 16040. https://doi.org/10.1038/natrevmats.2016.40 with permission from Springer Nature © 2016.

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5.1.2  Synthetic biomaterials Synthetic materials encompass a range of synthetic polymers, metals, ceramics, and glass whose widespread use is owed to the ability of such materials to be custom-synthesized according to specific requirements in terms of shape, architecture, chemistry, degradation, and mechanical properties via manipulation of polymerization parameters and composition of individual monomer proportions [5]. Although synthetics offer high reproducibility and improved functionality and mechanical attributes, it presents with poor bioactivity in terms of hydrophilicity and cellular interaction sites compared with most naturally derived materials [6]. Polycaprolactone (PCL), used in tissue engineering applications since the 1930s, has still maintained its popularity over the years along with its polyester counterparts, including the common poly(l-lactic acid) (PLLA) and poly(lactic-co-glycolic) acid (PLGA) [6] where composites with metals, glass, ceramics, and naturals materials are continuous in development for application to various tissue types.

5.1.3  Natural biomaterials Natural materials comprise biological materials derived from human, animal, marine, plant, algae, or microbial sources. Such materials can be broadly classified into protein-based materials, polysaccharide-based materials, and the emerging decellularized tissue-derived materials obtained through a process involving the complete removal of cellular components, thereby preserving only the native extracellular matrix (ECM) architecture [5,7]. Natural biomaterials are considered beneficial because of their inherent structural resemblance to native ECM, thereby offering enhanced biocompatibility and bioactivity compared with synthetics. The degradation and mechanical properties of natural materials are adjustable; however, control and predictability of degradation kinetics may be challenging. Despite their improved capacity for cellular interactions and adhesions, natural materials are often weak in mechanical strength and pose inconsistencies in composition and physical characteristics because of variations in batch production [5].

5.2  Biomaterials for tissue engineering applications 5.2.1  Biomaterials for hard tissue engineering Hard tissue engineering includes scaffolds for the repair and replacement of bone, cartilage, and dental tissues. Materials for the fabrication of scaffolds for such applications require high structural integrity, rigidity, and durability, in addition to a porous matrix network. Early investigated materials for hard tissue engineering scaffolds included metals (stainless steel, titanium, and cobalt–chromium alloys), ceramics (zirconia and alumina), and synthetic polymers (PMMA [polymethylmethacrylate] and PLGA, and Teflon-based materials) as single-phase constructs [8,9]. Newer materials of interest include ceramics (hydroxyapatite and tricalcium phosphate) and bioactive glass (silicate, phosphate, and borate-based), which are commonly investigated for use

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in bone and teeth repair because of their high mechanical stiffness, low elasticity, and hard brittle surfaces [4,10]. Widely investigated synthetic bioresorbable polymers for bone and cartilage tissue engineering include PCL, PLLA, poly(glycolic acid) (PGA), and polyvinyl alcohol (PVA) for the requirement of mechanical stability formulated as solid, hydrogel, or aligned fibrous constructs [10]. Polysaccharides (chitosan, alginate, and agarose), protein-based materials (collagen, gelatin, and fibrin), and glycosaminoglycans (hyaluronic acid and chondroitin sulfate) are synthesized as semisynthetic composites or surface modification materials in the common form of coatings and confer bioactivity to the scaffold [11]. As bone repair scaffolds necessitated the need for maintained mechanical strength during its degradation, surface-eroding polymers such as poly(ortho esters), polyphosphazenes, and polyanhydrides became of interest as opposed to the conventional bulk-eroding PLGA and PLLA polyesters [8]. For improved simulation of native bone mechanical stresses, the aforementioned materials have been fabricated as a variety of polymer–polymer and polymer–ceramic composites to confer rigidity to deformation-prone polyesters and brittle ceramics. Similarly, metal alloys have been explored for application in load-bearing scaffolds with growing research progress toward biodegradable porous magnesium alloys as alternatives to the conventional titanium derivatives [12,13].

5.2.2  Biomaterials for soft tissue engineering Soft tissues include skin, nerve, muscle, tendons, ligaments, and blood vessels. In contrast to hard tissue requirements, soft tissues display anisotropic behavior and endure large deformations, depending on the composition of elastin and collagen in the respective tissue type, thus requiring the need for elastic and pliable materials for scaffold fabrication [14]. The requirements for surface functionality and biomimicry of soft tissues potentiate the use of predominantly natural materials for scaffold fabrication, whereas synthetic polymers are utilized to confer mechanical support of the scaffold system [15]. Silicone-based materials were initially utilized for the repair of skin, neural, and vascular defects, thereby resulting in the commercialization of several silicone-based grafts either formulated alone (for tubular neural and vascular tissue repairs) or with layers or coatings of collagen (for skin healing grafts) [16–18]. Collagen-based materials (formulated as gels, sponges, hydrogels, and fibrous sheets) are considered to be the most popular natural material used in tissue engineering scaffolds for skin, nerve, and cardiovascular repair because of its high structural resemblance and abundance in native ECM and tissues [19]. Other commonly investigated natural materials are gelatin, fibronectin, chitosan, elastin, and silk, which are gaining rapid interest because of unfavorable material contraction and fast degradation rates observed with collagen during soft tissue healing [20,21]. Likewise, PLA, PGA, and PCL synthetic polyesters and PLGA and PLA-PCL synthetic copolymers have long been explored for soft tissue scaffolds because of their high elastic performance and strength. Likewise, polyhydroxyalkanoates and poly(polyol sebacate) (PGS)-based elastomeric polymers are gaining attention as promising materials for ligament, blood vessel, and cardiac tissue engineering [22,23].

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5.3  Research development of tissue engineering biomaterials 5.3.1  First-generation biomaterials The first generation of materials developed for use inside the human body was during the 1960s and 1970s and focused on bioinertness with the primary goal to “achieve a suitable combination of physical properties to match those of the replaced tissue with minimal toxic response in the host,” as described by Hench [24]. During those times, the intent of biomaterial use was to simply replace lost or damaged tissue with research emphasis placed on biomaterial development to minimize adverse foreign body immune responses [25]. Most implants were fabricated of single-phase materials derived from commercially available resources of higher purity grade as an attempt to reduce cytotoxicity and inflammatory responses [26]. Common first-generation biomaterials used to replace diseased soft and hard tissue included silicone rubber (soft tissue prosthetics), Teflon (tubular tissue constructs), 316L stainless steel, titanium and cobalt–chromium alloys (bone and joint replacement), Dacron mesh (blood vessel repair) and PMMA (bone repair cement), PVA (cartilage replacement), and PCL (skin replacement) synthetic polymers [24].

5.3.2  Second-generation biomaterials The second generation of biomaterials saw the shift in material research to include bioactivity as a biomaterial property as opposed to solely bioinertness [25]. Research was directed at synthesizing biomaterials that were capable of eliciting controlled actions and responses in the physiological environment of the tissues under repair to enhance interactions at the biomaterial–tissue interface for promotion of cell adherence to the implant surface for subsequent tissue growth [26,27]. A variety of bioactive materials such as bioactive glass, ceramics, glass–ceramic composites, and hydroxyapatite were clinically used in the 1980s for orthopedic and dental repairs [25]. A sequence describing the bioactive mechanisms involved in the bonding of 45S5 Bioglass to bone, the first synthetic material to bond to living tissue, was further elucidated by Hench from 1971 to 1998 [26,28]. This generation further included the development of biodegradable polymer materials, PLA and PGA, utilized clinically as biodegradable sutures. The feature of chemical disintegration, resorption, and eventual replacement of foreign polymer materials by regenerating tissues in the body instigated interest in the concept of tissue regeneration instead of tissue replacement. Second-generation biomaterial research and development was based on the principles of bioinertness, bioactivity, and bioresorbability [25]. The development of bioresorbable polymers heralded the innovation of controlled-release drug-delivery systems using pharmaceutical actives dispersed within polymeric matrices. The concepts of polymeric drug-delivery applications soon became an integral aspect to engineering the next generation of biomaterials for tissue regeneration scaffolds.

5.3.3  Third-generation biomaterials The need for biomaterials with the potential to respond to physiological and biological stimuli as living tissue was realized, and the third generation of biomaterials for scaffold

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fabrication commenced development. The goal of third-generation biomaterials involved the ability of such materials to stimulate cellular responses at a molecular level by either promoting or inhibiting specific cell activities [27]. The desirable features of bioinertness and bioresorbability of second-generation biomaterials converged to form the term biocompatibility. The research of third-generation biomaterials (from 2000s onward) was dedicated to the development of materials able to mimic native tissue architecture at macro- and microlevels with progression toward the nanoscale. Research efforts were concentrated at the development of composite materials encompassing combinations of synthetic and natural materials via cross-linking strategies across different material classes and molecular modification of such materials and composites utilizing specific proteins, peptides, and other biomolecules to attain increased similarity to the structure of native ECM constituents. An outline of the basic strategies and material types utilized in third-generation biomaterials is represented in Fig. 5.3. Despite the favorable property of textural resemblance to native tissues, natural-based materials presented with weakened mechanical strength upon hydration, leading to the utilization of mostly aldehyde chemical cross-linking strategies to enhance mechanical integrity. Later established cytotoxicity of aldehyde cross-linking promoted the exploration of ionic cross-linking methods (multivalent cations and anions), chemical cross-linking using natural substances (e.g., the extensively investigated genipin for collagen and chitosan cross-linking), photocross-linking, and thermal cross-linking for synthesis of synthetic, natural, or semisynthetic composite materials utilizing common elastomeric polyesters and natural-based polymers. Recent interests include the investigation of hydrogel materials (cross-linked hydrophilic polymeric networks capable of absorbing large amounts of water while remaining water insoluble in a solid swollen gel-like state) composed from natural proteins and polysaccharides with controlled drug-delivery strategies. This offers replication of microarchitectural features of native soft tissues and provision of mechanical, biochemical, and therapeutic cues to enhance functional tissue regeneration [17]. A variety of hydrogel materials composed of polysaccharides, proteinaceous materials, and synthetic polymers have been designed with stimuli-responsive mechanisms (i.e., temperature, pH, and ionic strength) that initiate processes of swelling and dissolution for the release of encapsulated cells and bioactives while offering the capacity for textural modification for customizable tissue resemblance [20,29]. The significance of the extent of ECM resemblance of biomaterials was recognized in terms of cell adhesion and migration on the material surface following gradual cell infiltration, proliferation, and differentiation throughout the scaffold structure. Consequently, the emergence of electrospinning techniques led to the widespread fabrication and investigation of micro- and nanofibrous materials as scaffolds for both hard and soft tissues. Electrospun materials presented improved resemblance to ECM microand nanoarchitecture, where cells could interact intricately with the surface topography of the materials’ structures for improved adhesion, migration, and proliferation. The use of nanomaterials offered an essential feature of physical and topographical cues in addition to macrosized mechanical cues of scaffolding materials. Recent research demonstrated improved efficacy of an R-peptide-modified electrospun PCL nanofibrous scaffold for biomimicry of native ECM of mesenchymal stem cells through provision of micro- and nanoarchitectural elements and biochemical molecules [30].

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In addition, the inclination toward bioregeneration entailed the synthesis and modi­ fication of scaffold materials via implementation of various growth factors, lowmolecular-weight drugs, and other suitable bioactives contained within drug-delivery controlled-release systems to promote tissue regeneration by means of serving as bioactive chemical and therapeutic cues for cell proliferation and direction [31]. These approaches were further combined with cell-seeding strategies to enhance vascularization, biocompatibility, and stability for long-term scaffold constructs. Similarly, decellularized tissue constructs in conjunction with cell-seeding strategies have been investigated as potential scaffold materials because of the preservation of original tissue microarchitecture, thereby producing structures of higher resemblance to native tissues and organs [7,32]. These multifunctional facets incorporated into the design and fabrication of third-generation scaffold materials aimed to enhance cellular interactions at the biomaterial–tissue interface and improve its similarity to native tissue ultrastructure as the gradual steps toward achieving maximum tissue biomimicry.

5.3.4   Fourth-generation biomaterials Initiation of endogenic bioelectrical signals from the activity of ion channels and pumps in the cell membrane of most tissue types has been extensively studied with the potential of offering insight into single cell behavior and control of proliferation, migration, and differentiation [33]. It is believed that all tissue types have the ability to generate and receive bioelectrical signals. Therefore, an additional prerequisite toward achieving maximum tissue biomimicry requires the replication of such electrophysiological features, thus motivating the development of the next-generation biomaterials: electroactive and conductive polymers and carbon-based and piezoelectric materials [33,34]. Such electroactive, electroconductive, and piezoelectric materials are considered as fourth-generation biomaterials with the goal of utilizing and manipulating cellular bioelectricity and electric signals for enhanced tissue regeneration and monitoring of cellular responses and activities for communication with host tissues via modulation of material–surface electric charge and biosensing capabilities (Fig. 5.4 depicts an example of the provision of electrical stimuli and signaling in cardiac tissue engineering) [33,36]. In promotion of the further development of such fourth-generation scaffold materials for regenerative medicine, a commentary by Ning and coworkers [33] postulated that “bioelectric signals can provide detailed information about a host material’s response to external environmental stimuli as well as generating desired feedback to the host and external environment to guide the behaviour of cells using electrical signals.” Advances in material research involve the investigation of conducting and electroactive materials for neural tissue engineering: polypyrrole (PPY), polyaniline (PANI), poly-3,4-ethylenedioxythiphene (PEDOT), and carbon nanotubes [16,17]. Graphene-based nanomaterials and piezoelectric materials have captured much attention for regenerative applications to the largely electroresponsive bone, neural, skeletal muscle, and cardiac tissues [33,34,37]. Although preliminary studies have investigated the regenerative potential of graphene-based materials, piezoceramics (e.g., zinc oxide, barium titanate, and lithium sodium potassium niobate), and piezopolymers (e.g., polyvinyl fluoride [PVDF], copolymers, and PLLA) on cell proliferation, direction, and differentiation, the concerns of cytotoxicity, biocompatibility,

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Figure 5.4  Electrospun albumin mats with 600-nm-thick Au electrodes assembled using an ECM-based hydrogel. Following, a layer of dexamethasone-loaded polypyrrole deposited onto selected electrodes. Finally, neonatal rat ventricular cardiomyocytes are seeded onto the device and are allowed to organize into a functional cardiac tissue. By connecting the device to an external amplifier, extracellular signals may be recorded from the tissue and stimuli may be delivered to the device for pacing and drug release. Reproduced with permission (Elsevier B.V. Ltd. © 2018) from R. Feiner, S. Fleischer, A. Shapira, O. Kalish, T. Dvir, Multifunctional degradable electronic scaffolds for cardiac tissue engineering, J Contr Release 281 (2018) 189–195.

and biodegradability of such materials is yet to be fully elucidated and resolved using appropriate in vitro and in vivo models. This compels the need for composite material development with a range of nontoxic biodegradable synthetic and natural-derived materials with the additional development of organic electroactive and conductive materials as opposed to the conventional inorganic derivatives [36–38].

5.4  Recent techniques in tissue engineering fabrication 5.4.1  Bioprinting: bioink materials for tissue engineering scaffolds Three-dimensional (3D) bioprinting has emerged as a versatile technique for the fabrication of biomimetic scaffolds [39]. This technology involves a computer-assisted process of bioink (a combination of biomaterials [natural and/or synthetic polymers], cells, biologicals, and bioactive molecules) deposition in a spatially controlled layer-by-layer pattern for

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the synthesis of bioidentical tissues, living tissue, and organ scaffolds [40,41]. In addition to efficient and multipurpose bioprinters, the success of bioprinting modalities is equally dependent on the development of materials for utilization as bioinks, which demand the requirement for a range of specially designed materials. Currently, hydrogel materials are considered the most favorable for 3D-bioprinting applications because of its instantaneous gelation and capacity for encapsulation of a variety of bioactive molecules and cells [41]. Although a range of ample materials and bioinks are presently available (as shown in Fig. 5.5), not all are compatible with the existing bioprinting modalities [41]. Hence, the need for increased research activity directed at further refinement and engineering of properties of existing third-generation and upcoming fourth-generation materials and development of new materials specifically for the domain of bioprinting [39–41]. Advancements in 3D bioprinting technologies have led to the inclusion of a fourth dimension resulting in the emergence of 4D bioprinting, which utilizes an assortment of stimuli-responsive materials (electrical-, thermal-, humidity-, magnetic-, pressure-, and photoresponsive or sensitive materials) for the biofabrication of smart scaffold materials with the capacity to undergo transformation in structure and function postprinting in response to internal and/or external stimuli and the environmental and structural dynamics of regenerating native tissues [39,42]. An example of a 4D approach would be the development of restricted and asymmetrical swelling bioprinted scaffolds for controlled and directional swelling conforming to the arrangement and shape of native tissues upon hydration [42]. The desire for bioink materials to confer concurrent properties for optimal tissue regeneration (mechanical strength and cellular viability and interaction) and bioprinting

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Collagen fibrils formation

Figure 5.6  Schematic illustration of the bioprinting process. Cells (indicated in orange) are seeded onto porous PLGA microspheres. After stirred culturing, cells infiltrate and proliferate into microspheres, producing cell-laden microspheres (CLMs). CLMs are encapsulated with a thin layer of thermoresponsive agarose–collagen composite hydrogel for bioprinting on a chilled platform, where agarose gelation occurs. Collagen fibrils are formed after culturing the construct at 37°C. Reproduced under CC license (Creative Commons CC BY 4.0) from Y.J. Tan, X. Tan, W.Y. Yeong, S.B. Tor, Hybrid microscaffold-based 3D bioprinting of multi-cellular constructs with high compressive strength: a new biofabrication strategy, Sci Rep 6 (2016) 39140. https://doi. org/10.1038/srep39140.

processes (rheological properties and printing compliance) is contradictory, thereby posing a significant challenge to the selection and development of suitable materials that are able to maintain a fine balance between structural, physiomechanical, and biological properties without causing major compromise to cellular viability and functionality of the materials [39,43]. Currently available conventional biomimetic materials often lack favorable mechanical properties required for bioprinting. The projected approaches for the improvement of bioink material properties include chemical and physical multicross-linking reactions, introduction of new functional groups, formation of composites from natural and synthetic polymers, microfiber reinforcement, and photocross-linking and thermoresponsive mechanisms for rapid gelation and solidification of synthesized constructs (Fig. 5.6 depicts scaffold-based bioprinting using a thermoresponsive hydrogel) [40,43].

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5.5  State-of-the-art and future perspectives Future developments hold great anticipation for tissue engineering innovations emanating from the integration and application of the much admired nanomaterials, hydrogel materials, and bioprinting modalities to the forthcoming generation of electroactive and piezoelectric materials for attaining the next level of biomimicry. A vital step toward effectively applying this new generation of materials to tissue regeneration strategies requires the thorough elucidation of cellular and molecular mechanisms governing living cells and the phenomenon of bioelectricity. Presently, detailed investigations into the interactions of in vivo topophysical properties, chemical stability, cytotoxicity, and cytocompatibility of these materials are highly necessitated [36]. A profound understanding of the aforementioned mechanisms and properties would allow the development and modification of the finest class of smart biomaterials with the potential of offering programmable and controllable biosensing and bioresponsive features, in situ, to actuate a series of responses such as shape change, release of pharmaceutical actives, growth factors, and inflammatory modulators, while simultaneously performing enhanced and specific biological and physiological functions (i.e., electronic innervations and conduction systems of tissue engineering scaffolds and modulation of inflammatory mediators) similar or equivalent to that of the native tissue requiring restoration of functionality.

5.6  Conclusions The research development of materials for the fabrication of tissue engineering scaffolds discussed in this chapter is recognized through the primary objectives of bioinertness, bioactivity, and bioregeneration where the next steps of material evolution build upon the outcomes of the previous phases. The third generation of biomaterials presented the major outputs of material modifications via utilization of hydrogels, nanomaterials, and controlled-release strategies for bioactives and cells for the goals of achieving biomimicry through physical, topographical, biochemical, and therapeutic cues for maximum cellular interaction with foreign scaffold materials. These preceding research outputs together with the investigations of bioelectrical cues utilizing upcoming electroactive and piezomaterials for the design of bioelectronics with advances in 3D- and 4D-bioprinting technologies’ open prospective for the creation of fully replicated living tissue engineering constructs encompassing biomimicry on macro-, micro-, and nanolevels where the intricate unification of abiotic and biotic components could be expected. Such hybrid systems termed as “bionic systems” may unlock potential for the synthesis of bionic cells through modification of active cells with stimuli-responsive nanomaterials to confer capabilities of sensing and relaying cellular signals, metabolic processes, and proliferation conditions of regenerating tissues as further advancement of fourth-generation materials [31].

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List of abbreviations 3D Three-dimensional 4D  Four-dimensional CLMs  Cell-laden microspheres ECM  Extracellular matrix PANI  Polyaniline PCL  Polycaprolactone PEDOT  Poly-3,4-ethylenedioxythiophene PGA  Poly(glycolic acid) PGS  Poly(polyol sebacate) PLGA  Poly(lactic-co-glycolic) acid PLLA  Poly(l-lactic acid) PMMA  Polymethylmethacrylate PPY  Polypyrrole PVA  Polyvinyl alcohol PVDF  Polyvinyl fluoride

Acknowledgments This work was supported by the Wits’ University Research Committee (URC) and the National Research Foundation (NRF) of South Africa.

References [1]  C.P. Bergmann, A. Stumpf, Biomaterials, in: Dental Ceramics. Topics in Mining, Metallurgy and Materials Engineering, Springer, Berlin, Heidelberg, 2013, https://doi. org/10.1007/978-3-642-38224-6_2. [2] K. Sadtler, A. Singh, M.T. Wolf, X. Wang, D.M. Pardoll, J.H. Elisseeff, Design, clinical translation and immunological response of biomaterials in regenerative medicine, Nat Rev Mater 1 (7) (2016) 16040, https://doi.org/10.1038/natrevmats.2016.40. [3] N. Huebsch, D.J. Mooney, Inspiration and application in the evolution of biomaterials, Nature 462 (7272) (2009) 426–432. [4] F.J. O’Brien, Biomaterials & scaffolds for tissue engineering, Mater Today 14 (3) (2011) 88–95. [5] F.M. Chen, X. Liu, Advancing biomaterials of human origin for tissue engineering, Prog Polym Sci 53 (2016) 86–168. [6] S. Stratton, N.B. Shelke, K. Hoshino, S. Rudraiah, S.G. Kumbar, Bioactive polymeric scaffolds for tissue engineering, Bioactive Mater 1 (2) (2016) 93–108. [7] D. Rana, H. Zreiqat, N. Benkirane‐Jessel, S. Ramakrishna, M. Ramalingam, Development of decellularized scaffolds for stem cell‐driven tissue engineering, J Tissue Eng Regenerat Med 11 (4) (2017) 942–965. [8] B.L. Seal, T.C. Otero, A. Panitch, Polymeric biomaterials for tissue and organ regeneration, Mater Sci Eng R Rep 34 (4–5) (2001) 147–230. [9] X. Yu, X. Tang, S.V. Gohil, C.T. Laurencin, Biomaterials for bone regenerative engineering, Adv Healthc Mater 4 (9) (2015) 1268–1285.

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[10] H.E. Jazayeri, M. Tahriri, M. Razavi, K. Khoshroo, F. Fahimipour, E. Dashtimoghadam, L. Almeida, L. Tayebi, A current overview of materials and strategies for potential use in maxillofacial tissue regeneration, Mater Sci Eng C 70 (2017) 913–929. [11] A.R. Armiento, M.J. Stoddart, M. Alini, D. Eglin, Biomaterials for articular cartilage tissue engineering: learning from biology, Acta Biomater 65 (2018) 1–20. [12] J. Yang, G.L. Koons, G. Cheng, L. Zhao, A.G. Mikos, F. Cui, A review on the exploitation of biodegradable magnesium-based composites for medical applications, Biomed Mater 13 (2) (2018) 022001. [13] M. Yazdimamaghani, M. Razavi, D. Vashaee, K. Moharamzadeh, A.R. Boccaccini, L. Tayebi, Porous magnesium-based scaffolds for tissue engineering, Mater Sci Eng C 71 (2017) 1253–1266. [14] G.A. Holzapfel, Biomechanics of soft tissue, Handb Mater Behav Models 3 (2001) 1049–1063. [15] Y. Zou, L. Zhang, L. Yang, F. Zhu, M. Ding, F. Lin, Z. Wang, Y. Li, “Click” chemistry in polymeric scaffolds: bioactive materials for tissue engineering, J Contr Release 273 (2018) 160–179. [16] S. Mobini, B.S. Spearman, C.S. Lacko, C.E. Schmidt, Recent advances in strategies for peripheral nerve tissue engineering, Curr Opin Biomed Eng 4 (2017) 134–142. [17] P. Sensharma, G. Madhumathi, R.D. Jayant, A.K. Jaiswal, Biomaterials and cells for neural tissue engineering: current choices, Mater Sci Eng C 77 (2017) 1302–1315. [18] S.P. Tarassoli, Z.M. Jessop, A. Al-Sabah, N. Gao, S. Whitaker, S. Doak, I.S. Whitaker, Skin tissue engineering using 3D bioprinting–an evolving research field, J Plast Reconstr Aesthetic Surg (2017), https://doi.org/10.1016/j.bjps.2017.12.006. [19] E. Sachlos, J.T. Czernuszka, Making tissue engineering scaffolds work. Review: the application of solid freeform fabrication technology to the production of tissue engineering scaffolds, Eur Cells Mater 5 (29) (2003) 39–40. [20] A.A. Chaudhari, K. Vig, D.R. Baganizi, R. Sahu, S. Dixit, V. Dennis, S.R. Singh, S.R. Pillai, Future prospects for scaffolding methods and biomaterials in skin tissue engineering: a review, Int J Mol Sci 17 (12) (2016) 1974, https://doi.org/10.3390/ijms17121974. [21] R.A. Kamel, J.F. Ong, E. Eriksson, J.P. Junker, E.J. Caterson, Tissue engineering of skin, J Am Coll Surg 217 (3) (2013) 533–555. [22] Y. Xue, V. Sant, J. Phillippi, S. Sant, Biodegradable and biomimetic elastomeric scaffolds for tissue-engineered heart valves, Acta Biomater 48 (2017) 2–19. [23] H. Ye, K. Zhang, D. Kai, Z. Li, X.J. Loh, Polyester elastomers for soft tissue engineering, Chem Soc Rev (2018), https://doi.org/10.1039/c8cs00161h. [24] L.L. Hench, Biomaterials, Science 208 (1980) 826–831. [25] L.L. Hench, J.M. Polak, Third-generation biomedical materials, Science 295 (5557) (2002) 1014–1017. [26] L.L. Hench, I. Thompson, Twenty-first century challenges for biomaterials, J R Soc Interface 7 (Suppl. 4) (2010) S379–S391. [27] R.J. Narayan, The next generation of biomaterial development, Philos Trans R Soc 368 (2010) 1831–1837. [28] L.L. Hench, An introduction to materials in medicine, Bioceram J Am Ceram Soc 81 (1998) 1705–1727. [29] E.A. Kamoun, E.R.S. Kenawy, X. Chen, A review on polymeric hydrogel membranes for wound dressing applications: PVA-based hydrogel dressings, J Adv Res 8 (3) (2017) 217–233. [30] R. Mobasseri, L. Tian, M. Soleimani, S. Ramakrishna, H. Naderi-Manesh, Peptide modified nanofibrous scaffold promotes human mesenchymal stem cell proliferation and longterm passaging, Mater Sci Eng C 84 (2018) 80–89.

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[31] S.K. Srivastava, V.G. Yadav, Bionic manufacturing: towards cyborg cells and sentient microbots, Trends Biotechnol 36 (5) (2017) 483–487. [32] R.H. Fu, Y.C. Wang, S.P. Liu, T.R. Shih, H.L. Lin, Y.M. Chen, J.H. Sung, C.H. Lu, J.R. Wei, Z.W. Wang, S.J. Huang, Decellularization and recellularization technologies in tissue engineering, Cell Transplant 23 (4–5) (2014) 621–630. [33] C.Y. Ning, L. Zhou, G.X. Tan, Fourth-generation biomedical materials, Mater Today 19 (1) (2016) 2–3. [34] C. Ribeiro, V. Sencadas, D.M. Correia, S. Lanceros-Méndez, Piezoelectric polymers as biomaterials for tissue engineering applications, Colloids Surf B Biointerfaces 136 (2015) 46–55. [35] R. Feiner, S. Fleischer, A. Shapira, O. Kalish, T. Dvir, Multifunctional degradable electronic scaffolds for cardiac tissue engineering, J Contr Release 281 (2018) 189–195. [36] M. Acosta, R. Detsch, A. Grünewald, V. Rojas, J. Schultheiß, A. Wajda, R.W. Stark, S. Narayan, M. Sitarz, J. Koruza, A.R. Boccaccini, Cytotoxicity, chemical stability, and surface properties of ferroelectric ceramics for biomaterials, J Am Ceram Soc 101 (1) (2018) 440–449. [37] S.R. Shin, Y.C. Li, H.L. Jang, P. Khoshakhlagh, M. Akbari, A. Nasajpour, Y.S. Zhang, A. Tamayol, A. Khademhosseini, Graphene-based materials for tissue engineering, Adv Drug Deliv Rev 105 (2016) 255–274. [38] M. Gajendiran, J. Choi, S.J. Kim, K. Kim, H. Shin, H.J. Koo, K. Kim, Conductive biomaterials for tissue engineering applications, J Ind Eng Chem 51 (2017) 12–26. [39] Y.C. Li, Y.S. Zhang, A. Akpek, S.R. Shin, A. Khademhosseini, 4D bioprinting: the next-generation technology for biofabrication enabled by stimuli-responsive materials, Biofabrication 9 (1) (2016), https://doi.org/10.1088/1758-5090/9/1/012001. 012001. [40] E.S. Bishop, S. Mostafa, M. Pakvasa, H.H. Luu, M.J. Lee, J.M. Wolf, G.A. Ameer, T.C. He, R.R. Reid, 3-D bioprinting technologies in tissue engineering and regenerative medicine: current and future trends, Genes Dis 4 (2017) 185–195. [41] M. Hospodiuk, M. Dey, D. Sosnoski, I.T. Ozbolat, The bioink: a comprehensive review on bioprintable materials, Biotechnol Adv 35 (2017) (2017) 217–239. [42] W.G. Whitford, J.B. Hoying, A bioink by any other name: terms, concepts and constructions related to 3D bioprinting, Future Sci 2 (3) (2016) FSO133, https://doi.org/10.4155/ fsoa-2016-0044. [43] W. Aljohani, M.W. Ullah, X. Zhang, G. Yang, Bioprinting and its applications in tissue engineering and regenerative medicine, Int J Biol Macromol 107 (2018) 261–275. [44] Y.J. Tan, X. Tan, W.Y. Yeong, S.B. Tor, Hybrid microscaffold-based 3D bioprinting of multi-cellular constructs with high compressive strength: a new biofabrication strategy, Sci Rep 6 (2016) 39140, https://doi.org/10.1038/srep39140.

Further reading [1] A.H. Rajabi, M. Jaffe, T.L. Arinzeh, Piezoelectric materials for tissue regeneration: a review, Acta Biomater 24 (2015) 12–23. [2] Y. Sang, M. Li, J. Liu, Y. Yao, Z. Ding, L. Wang, L. Xiao, Q. Lu, X. Fu, D.L. Kaplan, Biomimetic silk scaffolds with an amorphous structure for soft tissue engineering, ACS Appl Mater Interfaces 10 (11) (2018) 9290–9300.

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Denis Fabricio Viera Rey, Jean-Philippe St-Pierre Department of Chemical and Biological Engineering, University of Ottawa, Ottawa, ON, Canada 

6.1   Introduction Materials of both biological and nonbiological origin have been used as implants and prostheses to replace the structural functions of lost, diseased, or damaged body parts or aid in healing for quite some time. Evidence was unearthed to show that the Etruscans devised dental appliances as far back as the seventh century BC [1], while a wood and leather prosthetic toe dated between the eleventh and eighth centuries BC was found in Egypt [2]. Sutures may have been in use for a much longer time, maybe as far back as the Neolithic period, if we are to judge from skull evidence to the survival of a number of individuals following trepanation surgeries [3]. History is indeed filled with examples of our early ventures into the field of biomaterials [4]. However, issues of biocompatibility with the host, limited control over implant properties, and elevated risks of infection considerably hampered the outcome of these interventions. The sum of these experiences instructed the development of implants with the avowed goal of achieving bioinertness, the absence of a host response, from the 1950s onward. Implants developed during this period are now considered the first generation of biomaterials and remain in broad use in clinical settings today [5]. A paradigm shift toward achieving bioactivity led to a second generation of biomaterials, whereby new constructs were developed to interact with biological tissues and elicit a controlled response. This approach produced clinical breakthroughs in the 1980s. A third generation of biomaterials, involving the design of temporary scaffolds (or templates [6]) that control cellular responses and modulate specific signaling pathways, has since emerged and developed hand-in-hand with the field of tissue engineering in the 1990s [7]. This approach has fostered the development of increasingly complex instructive environments to integrate the accelerating pace of discovery in biological sciences toward tissue repair/regeneration. Of course, any tissue engineering scaffold must first and foremost be biocompatible, that is to say that it must elicit an appropriate host response for its intended application. As the ultimate goal of tissue engineering is to regenerate diseased, damaged, or lost tissues, scaffolds must also be biodegradable so as to make space for and be replaced by newly synthesized tissue. In many applications, the scaffold must perform the biomechanical functions of the native tissue, while the engineered tissue matures. As such, the mechanical properties must be carefully tailored to match, or at least approach, those of the native tissue. Furthermore, Handbook of Tissue Engineering Scaffolds: Volume One. https://doi.org/10.1016/B978-0-08-102563-5.00006-X Copyright © 2019 Elsevier Ltd. All rights reserved.

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the biomechanical environment imparted by the scaffold to resident cells, both through direct interactions and as a response to the application of forces, can impact cell responses and should be considered. Similarly, scaffold architecture is critical to guide tissue deposition and has increasingly been used to provide structural cues required to recreate macroscopic and microscopic tissue anisotropies. This must be achieved while taking into account the need to facilitate cell migration and to ensure adequate nutrient and metabolite transport. Scaffold design must also consider the biological interface presented to resident cells. Tailoring of the biological microenvironment is a challenging proposition, which is receiving increasing attention as our understanding of the complexity of the native cell niche expands. This is especially evident when one considers the spatiotemporal biological, mechanical, and structural changes taking place during tissue development and maturation, the processes that we are often trying to recapitulate at an accelerated pace with tissue engineering. In this chapter, we will review some of the main scaffold fabrication techniques used in tissue engineering and regenerative medicine in the context of the aforementioned design criteria. We will discuss the concepts behind the most frequently used techniques to produce porous scaffolds and provide a brief examination of the impact of additive manufacturing (AM) on the way we approach scaffold fabrication. We will also discuss the different approaches to hydrogel formation and tissue decellularization, two tissue engineering scaffold classes that have garnered increasing interest in recent years. Finally, a brief overview of organ and tissue bioprinting that combines state-of-the-art AM technologies and hydrogel fabrication know-how will also be highlighted.

6.2  Scaffold fabrication techniques 6.2.1  Porous scaffolds A key principle behind the emergence of the field of tissue engineering from efforts by Joseph Vacanti and Robert Langer in the 1980s was the idea that a porous biocompatible and resorbable material could be used as a scaffold to guide tissue regeneration [7]. In the 30 some years since their seminal work, a number of porous scaffold fabrication techniques have been developed and adapted to produce structures deemed appropriate for a number of applications pertaining to tissue engineering. In this section, we will review some of the most commonly used porous scaffold fabrication technologies and highlight important features and limitations of each technique. Technologies will be evaluated in regard to the reliance on compounds that can elicit cytotoxic effects and thus impact biocompatibility, as well as for the ability to control pore size, shape and orientation, overall porosity, and interconnectivity of the porous network, with an eye on the impact of these parameters on resulting mechanical properties. Considerations for the selection of approaches to functionalize the resulting structures with biological signals that afford a degree of control over the responses of cells interfaced with the scaffolds will also be touched on briefly.

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6.2.1.1  Solvent casting and porogen leaching Solvent casting and porogen leaching (SCPL) is a relatively simple, two-step technique first proposed by Mikos et al. [8]. It typically involves mixing a polymer solution dissolved in an organic solvent with particulates (the porogen), which are insoluble in the solvent, to create a slurry. This mixture is then cast into a membrane, and the solvent is evaporated to produce a solid composite of the polymer and porogen. The particulates are subsequently leached out of the structure by immersion into an aqueous solution to yield interconnected pores within the membrane. An illustration of this process is provided in Fig. 6.1(a). Salt particles have been used extensively as the porogen; however, other compounds including sugar, gelatin, and even paraffin (for which an aliphatic solvent must be used as leaching medium) particles have also been used. The choice of porogen dictates the size, shape, and uniformity of pores within the structure, whereas the concentration of porogen added to the slurry dictates the porosity and to a degree, the extent of interconnectivity between pores. Nevertheless, this technique can suffer from a lack of uniformity in the scaffold porosity owing to porogen settling during the solvent evaporation phase. The formation of a dense polymer layer on the external surfaces of the material, termed skin layer, can limit access to the internal porous structure for cells and further impacts the applicability of scaffolds produced with this approach. The thickness of scaffolds produced by this technique is also limited to 2–3 mm, as the leaching process is impeded in thicker constructs [9]. This drawback has somewhat limited the use of SCPL scaffolds for applications requiring relatively thin membranes; however, it also inspired the design and implementation of laminated constructs prepared by stacking multiple membranes [10]. SCPL scaffolds are typically characterized by weak mechanical properties and have been associated with cytotoxicity concerns associated with the residual solvent and porogen, contributing to a limited applicability [11]. The use of an organic solvent must also be considered when selecting a functionalization approach for SCPL. Indeed, proteins can be denatured by contact with these solvents, such that postprocessing functionalization schemes are typically favored.

6.2.1.2   Phase separation Phase separation is an oft-used scaffold fabrication technique that exploits the fact that a homogenous polymer solution will separate into polymer-rich and polymer-poor phases when it becomes thermodynamically unstable [12]. This can be achieved by cooling the solution to below the freezing point of the solvent to initiate crystal nucleation, which drives this phase separation in a process referred to as thermally induced phase separation. Once the solution is completely frozen, the resulting solid material is sublimed to remove the solvent and the polymer-rich regions are left to form scaffold walls, whereas the polymer-poor regions yield pores within the structure. Efforts to control crystal nucleation and growth via optimization of fabrication parameters such as polymer concentration, temperature, and the use of surfactants can yield a degree of control over pore size and distribution [13]. These structures are typically characterized by a high degree of interconnectivity to facilitate tissue ingrowth.

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Figure 6.1  (a) Schematic illustration of SCPL fabrication process. (b) Schematic illustration of electrospinning process. Examples of scaffold microstructures in insets.

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A number of groups have adapted this technique to produce parallel channel-like porous structures with the application of directional temperature gradients to control solvent crystal nucleation and growth starting from one face of the receptacle containing the polymer solution [14]. It should be noted that this method typically produces pores with dimensions on the lower end of the useful range for tissue engineering applications (often smaller than 200 μm), especially as it pertains to directional freezing, representing a drawback of the method. The use of many synthetic polymers as base materials may require organic solvents such as dioxane, impacting the functionalization strategy in much the same way as for SCPL scaffolds; however, water-­soluble polymers including collagen and polysaccharides have also been used successfully with this technique.

6.2.1.3   Gas foaming Gas foaming has also been used extensively to produce scaffolds for tissue engineering applications. This class of techniques relies on incorporation of a blowing agent to generate gas bubbles within a solid polymeric sample and consequently form pores. In the original application of this technique for tissue engineering, Mooney et al. [15] exposed poly(lactic-co-glycolic) acid discs produced by compression molding to high-pressure CO2 for extended periods, before rapidly returning the samples to atmospheric pressure to allow for expansion of the CO2 entrapped within the polymer matrix. Since this first report, others have proposed a number of alternative foaming agents, including chemical agents that produce gas upon thermal degradation [16]. A major advantage of this approach over other traditional scaffold fabrication techniques is the fact that it does not rely on the use of solvents or porogens, doing away with associated risks of cytotoxic effects. However, control over pore size and interconnectivity has proved challenging. Nonhomogenous porosity and pore interconnectivity have caused issues with reproducibility, as well as complications with producing uniform tissue deposition [17]. Furthermore, the heating sometimes required during compression molding step or to activate a chemical foaming agent can preclude the incorporation of biological molecules during the process. Attempts to address the lack of control on pore characteristics and uniformity have also been met with developments in microfluidics to generate highly uniform gas bubbles within a liquid polymer medium [18], an approach that somewhat sacrifices processing simplicity and requires access to equipment and expertise that is not as widespread.

6.2.1.4   Sintering Sintering has been used extensively to bond together polymer and ceramic particles (and fibers) into a cohesive porous scaffold. Typically, a bed of randomly packed particles is heated to a temperature above the glass transition temperature of the base material, but below its melting point. This condition encourages the diffusion of atoms or molecules on the surface of these beads toward the points of contact between particles to fuse them together and yield a porous macrostructure. The driving force for this phenomenon is the reduced free energy that accompanies a decrease in constructs’ surface area.

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Alternative modes of sintering that have been used include mild solvent treatment and pressure. The use of heat or solvent to achieve bonding between the particles can be deleterious to biological molecules, such that this technique would also often be incompatible with functionalization during processing. Sintered scaffolds are typically characterized by a small pore size and a lower porosity than those produced by other techniques, which is often associated with improved mechanical properties [19]. As such, these structures are often used for bone and dental tissue engineering applications [20]. A certain degree of control over porosity is afforded by the selection of particle sizes and treatment conditions; however, important increases in scaffold porosity have come from combining this approach with other fabrication techniques (see Section 6.2.2 for examples). This discussion presents the opportunity to mention that one of the most successful biomaterials in bone tissue engineering, 45S5 bioactive glass, is a very difficult material to process into three-dimensional scaffolds; yet, it has been implanted directly as a powder to fill bone defects with impressive results toward bone regeneration [21].

6.2.1.5  Electrospinning The textile industry has provided a number of fabrication techniques to the field of tissue engineering. Some textile-based scaffold fabrication techniques offer a useful alternative to the strategies discussed thus far in that they produce fiber-based materials, thereby affording the opportunity to recreate aspect of the fibrous nature of ECM in tissues. Electrospinning is undoubtedly the most broadly applied textile technique in tissue engineering. This method relies on the generation of an electric field between a polymer solution delivered at a controlled flow rate (usually through a needle) and a collector to draw the solution into a fiber [22]. This occurs because the application of a sufficiently high voltage onto the polymer solution causes it to become charged, such that electrostatic repulsion overcomes the surface tension of the solution and leads to elongation of a thread. This fiber solidifies on its way to the collector owing to the rapid evaporation of the solvent influenced by its large surface area-to-volume ratio. The outcome is a membrane of nonwoven fibers deposited onto the collector. This process is illustrated in Fig. 6.1(b). The porosity can be described as interconnected; however, the resulting pore size is generally much smaller than those obtained from the techniques to produce foamlike scaffolds described previously in this section. This can be considered an advantage for many applications, including tissue engineering of blood vessels or membranes to retain transplanted cells within a tissue defect [23,24]. Conversely, the small pore size has been shown to limit cell migration through the structure, which can impact its applicability as a scaffold to guide the regeneration of thicker tissues, such as bone or even full thickness articular cartilage. A number of process parameters including polymer concentration, solvent selection, flow rate, humidity, voltage differential, needle dimensions, as well as the distance between the needle and the collector can be manipulated to control fiber diameter. Careful optimization of these parameters allows the production of fiber diameters ranging from a few micrometers down to the low nanometer range. These parameters also directly impact the amount of residual solvent present in fibers as they reach the collector, thereby controlling the strength of bonds created at contact points between fibers. Furthermore, the extent of fiber alignment can be tailored by the use of a rotating drum as a collector and

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adjusting the rotation speed, while the use of a fanner enables the production of a membrane with a fairly uniform thickness. Both fiber diameter and alignment have been shown to impact the phenotype of interfacing cells [25,26]. Adaptations to the collector have also resulted in the formation of interesting scaffold shapes, notably for applications in vascular tissue engineering [27]. The need for organic solvents to dissolve many polymers used to produce electrospun scaffolds means that postprocessing functionalization approaches are often used. However, methods have been devised to achieve surface functionalization during the electrospinning process, thereby allowing the generation of biomolecular gradients via temporal changes to the solution fed to the needle. One such example involves the incorporation of peptide–polymer conjugates, whereby the short peptides are not affected by contact with the organic solvent and the difference in polarizability between the polymer and peptide segments helps drive the latter to the surface [28].

6.2.1.6   Self-assembly A number of research groups have designed peptide amphiphiles that spontaneously organize into ordered structures including nanofibers [29]. The careful design of the building blocks offers a tremendous degree of control over the biological activity, degradability, and structure of these self-assembled materials, thereby making these attractive alternatives to electrospun membranes for a broad range of applications in tissue engineering [30].

6.2.1.7   Hybrid scaffolds Driven by increased efforts to direct tissue deposition through structural cues and to achieve engineered constructs with macro- and microstructural organization that replicates the anatomical structures of native tissues, a number of reports have demonstrated the benefits of combining or modifying the fabrication techniques described above to achieve anisotropic structures. A number of examples originate from the field of cartilage tissue engineering, whereby hybrid scaffolds have been developed to direct ECM deposition into a depth-dependent organization reminiscent of that found in the native tissue. A recent study combined SCPL with electrospinning to produce scaffolds with anisotropic mechanical properties [31], thereby outperforming scaffolds produced by porogen leaching alone, while providing more porosity for tissue deposition than anisotropic scaffolds produced by electrospinning of each layer according to distinct fabrication parameters [32]. Others have combined fabrication techniques to generate multilayered constructs with anisotropic pore structures [33,34].

6.2.2   Additive manufacturing The porous scaffold fabrication techniques discussed in the previous section represent some of the main tools used by tissue engineers in their efforts to produce functional replacements for damaged and diseased tissues over the past 30 odd years. However, these techniques impose limitations on the design of scaffolds, providing only a relatively narrow range of control over pore shape, dimensions, and interconnectivity. Ultimately, techniques that facilitate our ability to achieve intricate control of the porosity and structural, biological, and mechanical properties of templates and to

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recreate the complex macro- and microstructures found in the tissues that we set out to engineer are needed. The rapid evolution of AM technologies in recent years has provided the means to attain improved control over the spatial distribution, shape, dimension, and interconnectivity of pores within scaffolds and allowed the application of an increasingly robust scientific approach to scaffold fabrication. AM techniques allow the fabrication of scaffolds, or more broadly structures, in a layer-by-layer manner according to data provided by a computer-aided design (CAD) drawing. These methods also facilitate the generation of scaffolds with carefully tailored shapes to match the dimensions of tissue defects that require repair. Thus, combining AM with CAD drawings generated based on tissue imaging data contributes to the application of personalized medicine in tissue engineering. The following section will discuss some of the AM techniques that have emerged as key approaches in tissue engineering.

6.2.2.1  Powder-bed three-dimensional printing Powder-bed three-dimensional (3D) printing is an AM approach that relies on the selective spatial delivery of a binder onto a powder bed by an inkjet printer [35]. Sequential application of powder layers interspersed by the addition of the binder and a drying step leads to the fabrication of 3D structures. The CAD drawing of the scaffold design is used to generate the cross-sectional data required to control the spatial application of the binder. The loose powder bed that remains uncoated by the binder serves to support the structure as it is being built (Fig. 6.2). This loose powder is then removed and a postprocessing sintering step is typically performed to fuse the powder particles together and provide mechanical integrity to the structure. The resolution of this technique is highly dependent on the properties of the powder (e.g., size, ease of application as a thin layer onto the powder bed) and binder selected to construct the green material, as well as the actual printer technology [36]. Consequently, the number of base materials available in powder form and with appropriate properties to provide sufficient resolution for scaffold fabrication remains somewhat limited. In addition to its relatively low cost, powder-bed 3D printing generally does away with toxic components such as organic solvents. This technique also supports the generation of scaffold composition gradients across its depth (z-axis) via changes in the makeup of the powder bed [37]. It does, however, suffer from important construct shrinkage, deformations, and even cracks during the sintering step, such that volume loss must be predicted and accounted for in the initial CAD design, which is not a straightforward assignment [38]. Another limitation of this technique pertains to the depowdering stage, whereby complete removal of the loose powder is made more challenging by the fabrication of structures with small pore (8 months) • No immune-related reaction • Indentation testing performed comparable with articular cartilage • Fibrin glue with chondrocytes regenerated homogenous cartilage at 6–12 weeks with lower to comparable GAG content • Acellular fibrin glue alone does not regenerate cartilage • Regenerated hyaline-like cartilage when implanted in the knee • Comparable matrix content and mechanical properties can be achieved when TGF-β3 is added • Improved mobility and reduced pain over 3 years • Increase in articular cartilage chondrocyte densities • Proliferation of fibrous tissue even after 6 weeks • Macroscopic appearance and histology results of cartilage repaired with alginate seem comparable with native cartilage • Biochemical parameters, such as sulfated GAG content of the alginate repaired cartilage, are inferior at end of 40-day period • Repairs with only alginate had decreased articular cartilage and more disturbed collagen fiber arrangement compared wth group with cells over 3 months • Alginate–agarose hydrogel regenerated predominantly hyaline cartilage in patients after 2 years • Regenerated cartilage integrated well with surrounding tissue in an osteochondral defect after 8 weeks • Growth factors increase collagen deposition in matrix • CS addition regenerated more neocartilage with superior structure and antiinflammatory activity after 12 weeks • Cartilage tissue forms after 4 weeks with positive staining for collagen type II, Safranin-O, and toluidine blue without added cells • With cells, cartilage defects are fully repaired with mostly hyaline cartilage after 6 months but only partially repaired with less hyaline cartilage content without cells • No

tendon as the preferred source for tissue engineering applications [41]. Marine organisms, such as salmon, marine sponges, and jellyfish, have been studied for collagen isolation as well and presumed to be safer compared with mammals [41–43]. As collagen type I is more readily available than collagen type II, those scaffolds are less expensive and easier to make. However, fabricating a scaffold with collagen type I alone is disadvantageous. Section 11.1.1.1 of this chapter describes the differences between the three types of cartilage; from there, it was learned that articular cartilage is composed mostly of collagen type II, not type I. This could be an explanation for why chondrocytes can dedifferentiate in just collagen type I scaffolds, whereas chondrocytes seeded onto collagen type II scaffolds show phenotype retention [44,45].

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Despite this, a study has also shown that in comparison with a collagen type II scaffold, a type I scaffold degenerated slower and was more uniform in wall thickness and pore structure [45]. Collagen scaffolds of many types and forms have been successful in facilitating the growth of cartilage. An ovine pepsin-digested, acid-soluble collagen type I sponge showed deposition of cartilage by chondrocytes over 4 weeks when cultured in vitro with bovine articular chondrocytes [46]. A collagen type II-based hydrogel that was embedded with chondrocytes exhibited significantly more hyaline cartilage than the control group after 8 weeks of injection into damaged rabbit cartilage [47]. Overall, hydrogels made out of natural materials such as collagen and gelatin are a good choice for cartilage scaffold because of their ability to present ECM like structure and function [48].

11.2.1.2   Fibrin Fibrin is a protein that makes up the insoluble network that forms a blood clot. It has been studied in a variety of applications, including as a scaffold itself and as a method for chondrocyte [49] and growth factor [50–52] delivery. Fibrin is mostly used in the form of fibrin glue, which is created by adding thrombin to the soluble fibrinogen. Fibrin glue is fully biodegradable and can be injected before it solidifies [53]. Fibrin with perichondral mesenchymal stem cells (MSCs) that were transfected to carry bone morphogenic protein-2 (BMP-2) or insulin-like growth factor I (IGFI) cDNA demonstrated partial-thickness rat articular cartilage lesion repair [54]. However, fibrin does not have great mechanical strength, so its ability to be used alone as a scaffold is limited [53]. So, although pure fibrin has been used with chondrocytes to regenerate cartilage [55], it has also been used with other materials such as alginate [56,57] or collagen [58] to improve mechanical properties. Results from these studies did not show major biochemical differences from other scaffold materials [53]. Another study investigating genipin cross-linked fibrin scaffolds did show collagen type II and aggrecan accumulation and increased compressive and shear moduli [59]. In addition to weak intrinsic mechanical strength, fibrin could trigger a humoral immune response because the thrombin used in fibrin gel formation is commonly obtained from animal sources [60]. To avoid this, fibrin gel was recently formed via an intraoperative process that eliminates the use of animal-derived thrombin [61].

11.2.1.3   Hyaluronan Hyaluronan, also called hyaluronic acid, is a nonsulfated GAG that is composed of repeating glucuronic acid and N-acetylglucosamine disaccharide units. Hyaluronan is an attractive material as it plays many roles in the cartilage of the joint, from lubrication to facilitation of MSC migration. It also interacts with the chondrocytes to organize the ECM and increase production of and retention of proteoglycans [62]. To improve on the mechanical stability that basic hyaluronan has, researchers have cross-linked hyaluronan via methods such as esterification, photocross-linking, and thiol-modified PEG linker to successfully create biodegradable scaffolds [63–65].

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Although not confirmed in clinical studies, some ex vivo studies have shown that the degradation rate of the hyaluronan scaffolds can even be altered to equal the rate of cartilage ECM synthesis [66,67]. This improves scaffold integration and prevents premature breakdown of the scaffold. Hyaluronan scaffolds have been commonly used in the form of injectable hydrogels that can fill irregular defects in a minimally invasive way [53,66,67]. Hyaluronan has also been used in different forms. Hyalograft C is a tissue-engineered graft made of an esterified derivative of hyaluronic acid, HYAFF-11. It has been seeded with autologous chondrocytes and shown to grow hyaline-like cartilage when implanted in the knee with improved mobility and reduced pain over 3 years [68]. Another study investigated an HYAFF-11 (benzylated hyaluronan) sponge and an ACP (cross-linked hyaluronan) sponge, made of cross-linked hyaluronan, in the femoral condyles of 4-month-old rabbits. After 12 weeks, the surfaces of the osteochondral defects were mainly hyaline cartilage. The ACP scaffolds facilitated good integration of the cartilage to the surrounding cartilage and healed the defects significantly better than the untreated and the HYAFF-11 scaffolds [69].

11.2.1.4   Chitosan Chitosan is a biocompatible polysaccharide derived from the hard exoskeletons of shellfish. It is a good choice of material for a cartilage engineering scaffold because its molecular structure is similar to many GAGs present in the cartilage ECM; because of this, chitosan can interact with any adhesion proteins or growth factors that are present [44]. Chitosan is the deacetylated version of the polymer chitin that is actually found in exoskeletons, and this extent of deacetylation can be controlled to alter its degradability in vivo. When the chitin is either 0% deacetylated or 100% deacetylated, the material is at its peak crystallinity. It is at its most amorphous when halfway deacetylated. Because of the crystallinity, highly deacetylated forms are degraded the slowest and can last up to several months in the body [70]. Chitosan has also been used in multiple forms. It is easily made into porous scaffolds that can have different porosities and pore orientations because of processing methods to achieve different mechanical characteristics [71]. It has also been shown to significantly increase articular cartilage chondrocyte densities in rat knees when injected as a solution [72]. For cartilage TE applications, chitosan is cross-linked with chondroitin sulfate (CS), a sulfated GAG that is present in hyaline cartilage ECM. Because of the similarities with the native environment in articular cartilage, these GAG-augmented chitosan hydrogels have shown to be very biocompatible and chondrogenic [73]. Chondrocytes cultured in chondroitin 4-sulfate (CSA)-augmented chitosan hydrogels retained their differentiation and characteristic chondrocyte morphology, but although they did synthesize proteoglycans such as native articular chondrocytes, the amount synthesized was not enhanced by the hydrogel compared with chondrocytes cultured on standard polystyrene surfaces [70]. Chitosan scaffold composites with other materials have also been explored. Collagen type II is commonly used to enhance the adhesion of chondrocyte clusters in vitro [74,75]. Composites of chitosan with silk fibrils, polyester, and polycaprolactone have also been studied [75–77].

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11.2.1.5  Agarose and alginate Agarose and alginate are other biocompatible polysaccharides that are commonly used in an implantable or injectable hydrogel form as a scaffold for cartilage regeneration. Both are extracted from algae; agarose from a type of red algae and alginate from brown algae. Both polysaccharides can induce cartilage growth when used as a scaffold in the short term, but studies have also shown these materials increase the chances of an immune response [78–80]. The polysaccharides can be used in a hydrogel form to encapsulate cells in a more realistic, 3-D environment and keep them in a rounded shape. Reculturing dedifferentiated chondrocytes, previously passaged in a monolayer, in an alginate hydrogel helped to redifferentiate the cells to primary chondrocyte level [81]. Agarose is especially attractive in that its temperature-sensitive water solubility makes it convenient for cell encapsulation [44]. There are issues with using agarose and alginate as scaffolds for cartilage TE. The degradation properties of agarose cannot be easily altered to fit a desired rate. Alginate also does not have ideal degradation properties as it does not degrade quickly enough in vivo to allow room for new tissue growth. So, long-term alginate scaffolds tend to lose their integrity within a year [44]. One study showed that a cell-seeded alginate sponge gave good histological results but inferior biochemical parameters, including sulfated GAG quantity, when compared with native cartilage [78]. Many fundamental studies have used agarose scaffolds in vitro to study the effects of active mechanical stimuli on chondrocytes and its influence on ECM production [82–84].

11.2.1.6   Silk Silk is a naturally produced fiber made by silkworms. Silk scaffolds are composed of this natural silk fibroin and a filament core protein that is coated with a gluelike protein called sericin [40]. Silk is biocompatible, has very strong mechanical strength, slow degradation, and can be arranged into many different scaffold forms [85]. Highly porous silk sponges seeded with human MSCs facilitated greater collagen type II quantity, GAG production, and chondrocytic gene expression levels than collagen-based sponges in vitro [86]. Silk hydrogels have been compared with porous silk scaffolds to investigate the effect of the hydrogel, maintaining the spherical morphology of chondrocytes on cartilage regeneration ability. Both scaffold types supported chondrocyte cell proliferation and phenotype, but the properties of the porous scaffold leveled out after day 42; in contrast, DNA and GAG content, along with compressive modulus of the cultured silk hydrogels, continued to increase past this 42 day mark [87].

11.2.1.7  Native cartilage matrix One idea in tissue engineering is to use the ECM produced directly by native tissue as a scaffold for new tissue growth. Decellularized ECM would have all the native proteins present in the natural form and structure—eliminating the challenge of finding biocompatible materials and forming them into a biomimetic structure. It is also thought that other bioactive molecules, such as growth factors, left over in the scaffold, will promote better matrix and tissue production [53]. In addition, ECM scaffolds encourage more specific

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tissue formation instead of inferior scar tissue [88]. Different decellularization techniques can be used, such as thermal shock, freeze–thaw cycles, enzymatic treatments, or detergent treatments [89], but harsh decellularization techniques can result in decreased mechanical properties and loss of proteins and bioactive cues. Cell-derived ECM scaffold sheets can also be produced from cultured cells derived from patients that can eliminate disease transmission and are more easily decellularized and recellularized [90–93]. Cheng et al. harvested cartilage from porcine knee joints and created spongelike scaffolds by homogenizing, washing, freezing, and then lyophilizing the tissue [94]. When human adipose-derived stem cells (ASCs) were seeded onto the scaffold, chondrogenic differentiation was induced and resulted in abundant cartilage ECM protein production, rounded cells, and mechanical properties close to those of native cartilage [94]. Gong et al. used acellular cartilage sheets that were harvested from adult pig ear and layered them in a sandwich model that alternated the cartilage sheet and a layer of seeded chondrocytes for a total of 20 sheets; the engineered cartilage maintained the phenotype after 12 weeks and reached 87% of native ear cartilage mechanical properties [95]. This model is useful because the cartilage sheets can be cut and stacked to fit different sizes and shapes. Yang et al. investigated human cartilage-derived scaffolds seeded with canine MSCs implanted into mice [96]. The implanted cells were viable and regenerated new cartilage that contained GAGs and collagen type II [96]. These studies demonstrated the potential of the decellularized cartilage use for cartilage tissue engineering. However, further studies are required to establish the safety of the animal-derived and decellularized cartilage for human use.

11.2.2  Synthetic materials Many synthetic materials are popular for the use of cartilage tissue engineering scaffolds as well (Table 11.2). Compared with natural materials, synthetic polymers are advantageous because they can be produced as needed. They can also be customized with different chemical modifications to create a scaffold with tunable degradation rate and mechanical strength. These polymers have been made into many different forms, such as felts, meshes, and porous scaffolds. Common polymers such as polyglycolic acid (PGA), polylactic acid (PLA), and polylactic-co-glycolic acid (PLGA) are Food and Drug Administration (FDA) approved for certain applications in humans, which is why research into using these specific synthetic materials is so desirable.

11.2.2.1   Polyglycolic acid The most commonly used polymer for cartilage tissue engineering is PGA. PGA is a poly(α-hydroxy ester) that can degrade totally within 4–12 months by the mechanism of hydrolytic scission; this is faster compared with the degradation of other polyesters [97]. PGA is a polymer that is higher in crystallinity and hydrophilic and loses mechanical strength rapidly before total degradation. The main degradation product of PGA is glycolic acid, which is biocompatible and naturally resorbed into the body. PGA has been made into porous structures with various levels of porosity and interconnectivity by using the salt-leaching method [53]. PGA has also been developed into a nonwoven mesh or felt for cartilage engineering. However, although these forms are high in

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Table 11.2  In vivo evaluation of commonly used synthetic biomaterial scaffolds. Material PGA

In vivo performance [98]

• Repaired

cartilage with and without chondrocytes chondrocytes, the regenerated cartilage is smooth, has uniform GAG distribution, integrates well with underlying bone, and shows formation of the subchondral plate and columnar alignment of chondrocytes after 6 months • Regenerated cartilage is smooth, firm, comparable in overall color and appearance to native cartilage after 6 weeks, and filled most defects • Long-term subchondral bone reformation is inconsistent at 1 year • No completely normal results compared with native cartilage at 1 year • Neocartilage was collagen type II dominant and low in GAG content • At 12 weeks, neocartilage appeared smooth, shiny, and white • Regenerated cartilage appeared hyaline-like when examined histologically • After 6 months, scaffolds seeded with MSCs resulted in smooth, uniform cartilage surface with a color similar to that of the host tissue • Histological results showed hyaline-like cartilage using scaffolds seeded with MSCs • Scaffolds seeded with MSCs regenerated cartilage similar in color and texture to host tissue at 24 weeks • Regenerated cartilage showed an increase in GAG content and collagen type II, with the collagen levels similar to that in healthy cartilage when evaluated histologically • At 12 weeks, regenerated cartilage is smooth and well integrated into surrounding cartilage • Histological results showed smooth, hyaline-like cartilage as well as abundant GAG and collagen type II content • With

PLA

[105] [106]

PLGA

[132]

PCL

[178]

PLGA/ articular cartilage ECM

[179]

PLGA/nanoHydroxyapatite

[180]

porosity and interconnectivity, the initial mechanical strength of meshes is weak [53]. Because PGA degrades rapidly to allow new cartilage matrix to fill the empty space, the strength of the new matrix should soon be able to make up for the low mechanical integrity of the scaffold itself to be able to function in a load-bearing region. The predictable and quick degradation rates of PGA have allowed for good cartilage ECM production [98–100], with even greater proteoglycan production than collagen or PLGA copolymer scaffolds [101]. However, quick loss of mechanical integrity and fast degradation are the main reasons why PGA needs to be modified with slow-degrading polymers to develop cartilage scaffolds. A study in 2017 used scaffolds made of nonwoven PGA fibers that were prepared into 10-mm by 2-mm cylinders, with the addition of a 0.5% PLA solution to solidify its shape. Autologous bone marrow MSCs were isolated from the anterior superior

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(a)

(b)

(c)

(d)

(e)

(f)

(g)

Gross view grading table Grading

2-week group

4-week group

8-week group

Blank

Complete repair

1/9

10/18

11/19

0/6

Incomplete repair

6/9

5/18

5/19

0/6

No repair

2/9

3/18

3/19

6/6

Figure 11.3  Gross view and repair of osteochondral defects (indicated with black arrows) repaired with a nonwoven PGA fiber scaffold. (a) Comparison of the blank and 2-week groups. (b and c) The blank and 2-week groups showed formation of a fibrouslike tissue on top or deep inside the defect, with irregular surfaces. (d) Comparison of 4- and 8-week groups. (e and f) The 4- and 8-week groups showed regeneration of cartilage- and bonelike tissue with smoother surfaces and good integration into surrounding tissue. (g) Overall repair results. Image from A. He, L. Liu, X. Luo, Y. Liu, Y. Liu, F. Liu, et al., Repair of osteochondral defects with in vitro engineered cartilage based on autologous bone marrow stromal cells in a swine model, Sci Rep 7 (2017) 40489. https://doi.org/10.1038/srep40489. This work is licensed under a Creative Commons Attribution 4.0 International License and has been reproduced following the guidelines. To view a copy of this license, visit https://creativecommons.org/licenses/by/4.0/legalcode.

iliac spine of swine, passaged twice, seeded onto the scaffolds, and implanted into femoral condyle osteochondral defects of the same swine. The defects with the implant contained two constructs stacked together and stitched to the surrounding cartilage. Fig. 11.3 shows the gross view of the repaired regions; after 2 weeks, most defects show regeneration of fibrouslike tissue, whereas after 4 and 8 weeks, more than half of the defects showed complete regeneration of cartilage- and bonelike tissues. As shown in Fig. 11.4, histological results show cartilage-specific Safranin-O and collagen type II staining in the 4- and 8-week groups [102].

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Scaffolds for cartilage tissue engineering

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Figure 11.4  Cartilage-specific staining of osteochondral defects (indicated with black arrows) repaired with a nonwoven PGA fiber scaffold over a period of 8 weeks. Blank and 2-week defects show negative Safranin-O and collagen II staining. The 4- and 8-week groups show strongly positive Safranin-O and collagen II staining in cartilage defect areas with negative staining in the bone defect areas. Image from A. He, L. Liu, X. Luo, Y. Liu, Y. Liu, F. Liu, et al., Repair of osteochondral defects with in vitro engineered cartilage based on autologous bone marrow stromal cells in a swine model, Sci Rep 7 (2017) 40489. https://doi.org/10.1038/srep40489. This work is licensed under a Creative Commons Attribution 4.0 International License and has been reproduced following the guidelines. To view a copy of this license, visit https://creativecommons.org/licenses/by/4.0/legalcode.

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11.2.2.2   Polylactic acid PLA is a more amorphous, hydrophobic, poly(α-hydroxy ester) polymer than PGA and has a degradation time of greater than 2 years. Like PGA, a PLA scaffold will lose mechanical integrity before it is completely degraded. It is established that chondrocytes show less affinity for PLA surfaces than PGA surfaces [103]. However, over time, the total amount of cells attached is similar between PGA and poly-l-lactic acid (PLLA) scaffolds [103,104]. Because of the slower degradation rate, PLA is more suitable as a scaffold material for articular cartilage defects that experience long periods of load right after implantation. The slower degradation allows time for more ECM production before the scaffold loses integrity. PLA scaffolds are usually made as thin, nonwoven, fiber meshes because of the thinness of articular cartilage [105]. Hyaline-like cartilage has been found after 6-week implantation of undifferentiated perichondral cells on PLA meshes in a rabbit femoral condyle model [105]. PLA grafts seeded with autogenic and allogenic perichondrium cells in the same rabbit model have resulted in repairs that were grossly successful, but the newly formed cartilage tissue is biochemically inferior to native articular cartilage [106,107]. Like PGA scaffolds, PLA scaffolds also supported increased proteoglycan synthesis over the period of 3 weeks [101].

11.2.2.3   Polylactic-co-glycolic acid PLGA is a copolymer of PLA and PGA monomers that degrades in the same way, by hydrolysis. PLGA is advantageous because the PLA:PGA ratio can be altered to reach desired properties, such as degradation rate, hydrophilicity, and mechanical strength. Including more PLA will result in a more amorphous material with a longer degradation time; a 75:50 PLA:PGA copolymer degraded within 4–5 months, whereas a 50:50 PLA:PGA copolymer degraded within 1–2 months [108]. Like the individual polymers, PLGA degrades into nontoxic, resorbable lactic and glycolic acids that can be sequestered by the body. Like PGA and PLA, PLGA is primarily used as a nonwoven or woven fiber mesh for cartilage engineering. It is also commonly used with other natural materials to develop better cartilage scaffolds. For example, gelatin/hyaluronic acid–treated PLGA (PLGA-GH) sponge scaffold showed superior cell attachment, proliferation, and ECM secretion compared with pure PLGA scaffolds [109]. In a separate study, alginate was used to deliver chondrocytes uniformly throughout PLGA pads before implanting into rabbits and showed good early histological and biochemical results [110]. Together, PGA, PLA, and PLGA present a number of material choices to be developed into three-dimensional and porous scaffolds for articular cartilage regeneration.

11.2.2.4   Others Besides the most commonly used polymers, there are other synthetic materials used for cartilage engineering scaffolds as well. Polycaprolactone (PCL) has a long degradation time (1–2 years) and is stronger than PGA, PLA, and PLGA [97]. It is more resistant to rapid hydrolysis, which allows it to offer structural integrity to the defect

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over longer periods of time and right away. This material also has high drug permeability, but poor hydrophilicity. PCL can be formed into threads to create felt or mesh scaffolds or into a porous scaffold by salt leaching. An electrospun PCL scaffold seeded with adult bone marrow MSCs and transforming growth factor beta 1 (TGFβ1) differentiated the cells into a chondrocyte phenotype and facilitated the growth of a matrix containing collagen type II, cartilage proteoglycan link protein, and aggrecan [111]. Creating copolymers with PCL can result in a material with both the strength and elasticity of PCL, while changing the degradation time slightly to fit the scaffold needs [103]. Honda et al. used PCL and PLA copolymer sponge with cartilage-like structures. When implanted in mice, the scaffolds exhibited minimal degradation after 4 weeks [112]. Polyethylene glycol (PEG) is an extremely hydrophilic polymer, which prevents protein and cell adsorption and is used in copolymers often. Adding PEG increases hydrophilicity, which can increase biocompatibility by decreasing the adsorption of antibodies and other proteins. It can be added to alter overall cell attachment to the scaffold or even added in specific areas of an implant to allow cell attachment only on certain places [53]. PEG itself has similar compression properties to cartilage but is not ideal to be used alone as a cartilage engineering scaffold because it does not naturally degrade to allow new ECM ingrowth. It has been made into many copolymers such as a poly(propylene fumarate-co-ethylene glycol) hydrogel to carry cells and fill cartilage defects [113–120]. Some other synthetic polymers that have been studied are polyethylene oxide [121], polyurethane [122], polyethyleneterephthalate, and polytetrafluoroethylene [123].

11.2.3  Composite scaffolds 11.2.3.1   Chondroinductive approaches 11.2.3.1.1   Growth factors Growth factors function to promote cell proliferation, healing, and development in the body. Although growth factors can be added to cell media, many studies have investigated delivering growth factors from the implanted scaffold in a controlled manner to promote tissue growth over time. This is advantageous because simply injecting growth factors into a defect alone will not give a lasting effect, as they will diffuse before they can have an effect on the cells in the defect. A common family of growth factors used specifically in cartilage tissue engineering is the TGF-β superfamily, which is involved in the inflammatory repair response that follows injury [140]. TGFβ1 is a specific isoform used that has been shown to stimulate chondrogenesis and cell proliferation [141–143]. Bone morphogenic proteins (BMPs) are also part of this family and have effects on chondrogenesis and osteogenesis; for cartilage engineering, BMP-2 plays a role in regulating chondrocyte differentiation and altering extracellular matrix turnover [140]. Other commonly used growth factors are IGF-I and basic fibroblast growth factor 2 (FGF-2). One strategy for adding growth factors to scaffolds is to encapsulate them. This involves loading the factors into polymer scaffolds with controllable degradation times so they

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can be released in a predictable way as the scaffold degrades. A biphasic, porous PLGA scaffold loaded with TGF-β was implanted into a goat femoral condyle osteochondral defect and showed slightly better quality cartilage repair by biomechanical and histological results with the added factor than without [144]. The factors can also be encased in a bulk scaffold material, such as a hydrogel, which can restrict the diffusion of the factors out of the scaffold for a period of time unlike meshes or felts [53]. Changing the amount of cross-linking in the hydrogel can alter the level of free space available for diffusion of the molecules [145]. A fibrin hydrogel is also able to be modulated by changing the fibrinogen or thrombin concentrations [146]; fibrin gels have been used to release IGF-I in vivo [50,147]; and TGF-β1 in vitro to effectively promote cartilage regeneration. Microparticles encapsulating growth factors can also be incorporated into porous scaffolds. The second method to add growth factors to cartilage engineering scaffolds is to covalently bond the molecules to the scaffold material. Although this depends on the binding reaction and the specific molecule itself, this method can sometimes reduce the effectiveness of the growth factors by blocking their active regions [53]. If that can be avoided, such as tethering the factor with a longer chain, this method has an advantage because the movement of the factor away from the defect is restricted. TGF-β1 that was tethered to a polymer scaffold stimulated significantly more cartilage matrix production than an equivalent amount of soluble TGF-β1 [148]. Similarly, a PEG hydrogel system with encapsulated chondrocytes and tethered TGF-β1 increased DNA, GAG, and collagen content while maintaining articular cartilage phenotype better relative to soluble TGF-β1 [149]. Growth factors are an effective way to stimulate cartilage tissue growth, but also expensive. Optimal concentrations, ideal growth factor cocktails, and better methods of incorporation need to be studied to best take advantage of this resource for future regeneration methods.

11.2.3.1.2   Chondroitin sulfate Chondroitin sulfate (CS) is a sulfated GAG in cartilage that helps to draw water into cartilage and therefore helps it to resist compression. It has also been shown to inhibit cartilage destruction [150] and increase chondrocyte metabolism, therefore inducing the synthesis of more collagen, proteoglycans, and cartilage markers [151]. Many people actually take it as a dietary supplement to help treat osteoarthritis. Because it is a natural component of native cartilage and the benefits it provides, CS has been added into scaffolds to help increase amount and quality of regenerated cartilage. CS is readily water soluble and can be easily added into hydrogels by submerging them in media supplemented with CS. A fibrin–alginate hydrogel supplemented with CS increased chondrocyte numbers and GAG production more than the hydrogel not supplemented with CS [152]. CS can also be cross-linked to other polymers. For example, it has promoted chondrogenic phenotype when cross-linked to a chitosan scaffold [73]. CS also has been covalently attached to porous collagen type I scaffolds, which resulted in significantly higher cell proliferation and proteoglycan content than the collagen scaffolds without CS [153]. A biodegradable copolymer CS-grafted PLLA scaffold has also successfully promoted chondrocyte phenotype gene expression, morphology, and matrix production [154].

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11.2.3.2   Hybrid scaffolds Recently, there has been a lot of interest in hybrid scaffolds, or composites, which contain more than one type of cartilage TE material. Some of these hybrid scaffolds have already been mentioned. The goal of using multiple materials in one scaffold is to combine different benefits and make up for some drawbacks of another material. The scaffold can combine a natural material with a synthetic material or use multiple materials of the same general material type. The composites can also include two materials in the same form or in different forms. An example of using multiple synthetic polymers in a scaffold is one that combines PGA and PLGA. A porous 75:25 PLGA scaffold was reinforced with PGA fibers in a specific orientation to increase compressive modulus of the scaffold by 20% to make it more suitable for load-bearing applications [155]. A scaffold using two natural materials both in the same form is a fibrin/hyaluronan composite gel. When implanted into mice and compared with fibrin alone, the fibrin/hyaluronan composites formed cartilage-like tissue earlier, had more stable maintenance of chondrocyte phenotype, and produced more GAG, collagen, and overall ECM throughout the whole study [156]. Native cartilage is composed of many protein fibers with a gel-like ground substance dispersed throughout that protein network. Because of this, many composite scaffolds have taken the approach of combining a mesh or porous scaffold and a hydrogel. The idea is for the solid component to provide mechanical strength and structure, whereas the hydrogel allows uniform cell dispersion and maintenance of a rounded, chondrocyte-like morphology. Choosing different combinations of materials for these phases can also achieve desired characteristics such as degradation time, strength, and specific functions that natural materials play in cartilage. Filling a PLGA mesh with chondrocytes encapsulated in fibrin glue produced 2.6 times more GAGs in 4 weeks than the PLGA mesh did alone [157]. Fibrin hydrogel was also used in conjunction with a stiff, macroporous polyurethane scaffold to allow for more cell viability and even distribution, more GAG content, and higher collagen type II and aggrecan gene expression levels [122].

11.3  Osteochondral approach Sometimes an articular cartilage defect is so large and deep that it penetrates into the underlying subchondral bone. When that occurs, it is called an osteochondral defect. These defects are more difficult to treat than smaller cartilage defects because of the need to not only repair cartilage, but also regenerate bone and a cartilage–bone interface. Fig. 11.2 depicts the different zones of articular cartilage and how it transitions into the subchondral bone. Scaffolds for osteochondral defects should ideally mimic these zones to repair the defect in the most realistic way.

11.3.1  Allografts Osteochondral allografts are a common way to repair these defects. These allografts can be fresh, used within 7 days of donor death, or harvested and kept frozen until

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needed. Fresh allografts have shown good success over time. Follow-up studies have shown 100% graft survival rate after 3 years [158] and 96% graft survival rate after 2.9 years [159]. A long-term study of 66 repaired knees over 15 years showed 91% osteochondral allograft survival after 5 years, 76% after 10 years, and still 76% after 15 years [160]. Fresh allografts have to be used soon after harvest, whereas frozen allografts are more convenient because they can be stored and made available at any moment. However, frozen grafts do not contain viable chondrocytes like fresh allografts do, which reduces their regenerative ability. The frozen cartilage may also crack and delaminate because of damage caused to the ECM [161]. One study compared allografts after fresh or frozen storage and after 6 months of implantation in goat knees. Fresh allografts contained a high density of chondrocytes at the articular surface that secreted the lubricant proteoglycan-4 (PRG4) at a high rate, whereas frozen allografts did not contain any surface chondrocytes, which resulted in a low rate of PRG4 secretion [162].

11.3.2  OC scaffold configurations 11.3.2.1   Single phase Single-phase scaffolds are the most basic type of scaffold to treat osteochondral defects. These scaffolds are fabricated with the same materials in the same composition throughout. Considering the zonal structure and the corresponding compositional variation in OC tissue, it does not make the most sense that a single-phase scaffold would be the best choice. However, early TE approaches utilized single-phase scaffolds with and without cells and/or growth factors with some success. For example, PLLA scaffold seeded with perichondrocytes showed type I collagen formation and supported cell attachment and survival [105]. In a study by Coburn et al., electrospun poly(vinyl alcohol)-methacrylate with CS-methacrylate regenerated hyaline-like cartilage with increased collagen type II in vitro. However, although the scaffold increased proteoglycan content in a rat osteochondral defect, it did not reach the proteoglycan deposition of native articular cartilage [163].

11.3.2.2   Multiphase Because of the zonal differences in articular cartilage and subchondral bone, biphasic and triphasic scaffolds have been investigated for OC tissue engineering. These scaffolds have multiple layers, representing cartilage and bone in the case of a biphasic configuration, and cartilage, bone, and cartilage–bone interface in a triphasic scaffold. These multiphasic scaffolds can be formed using different materials, such as a softer one for the cartilage phase, a stiffer one for the bone phase, and an intermediate stiffness phase to support bone–cartilage interface formation. Because of simplicity, biphasic scaffold configuration has been extensively studied for OC tissue engineering [4]. One example of a biphasic scaffold is one using a collagen composite sponge as the cartilage phase and 75:25 PLGA as the bone phase [164]. After being seeded with bone marrow MSCs and implanted into a femoral condyle defect for 4 months, there was good integration, cartilage-like tissue with a smooth surface, and

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new underlying bone tissue. Liu et al. were able to repair osteochondral defects in rabbits with a biphasic scaffold that featured a freeze-dried Col-1/hylauronate sponge cartilage phase and a beta-tricalcium phosphate (TCP) bone phase [165]. There are many other cartilage/bone phases that have been used in biphasic scaffolds. Some combinations that were studied are collagen/GAG phosphate [166], agarose hydrogel/45S5 bioactive glass [4:1] microspheres [167], and HYAF-11 sponge/calcium phosphate [168]. Lately, studies focused on accurately regenerating the interfacial zone by adding another layer to biphasic scaffolds, which resulted in the development of triphasic scaffolds. In this configuration, there is a dedicated layer for bone, bone–cartilage interface, and cartilage. For example, a trilayered scaffold was developed with a collagen type I and hydroxyapatite (HA) bone layer, a collagen type I/II and HA transition layer, and a collagen type I/II and hyaluronic acid cartilage layer. The idea is that the different biomechanical properties in each layer would promote the desired cell differentiation in that section. When cultured in vitro, the scaffold’s high porosity and interconnectivity supported homogenous cellular infiltration and distribution and a seamless transition between the three layers [169]. Most recently, Kang et al. created a triphasic scaffold that varied in pore structure and material, going from a biomineralized bone layer, to an anisotropic macroporous cryogel transition layer, and then a hydrogel cartilage layer. When the top two layers were seeded with cells and subcutaneously implanted in mice, the scaffold successfully recruited cells into the bottom bone layer and regenerated cartilage tissue with similar zonal characteristics to native articular cartilage [134].

11.3.2.3   Gradient Multiphasic scaffolds have performed well in terms of forming the intended tissues, but their integration of the individual tissues has become an issue because the scaffolds are formed with biomaterials of varied strength and structure. To overcome this, an integrated scaffold approach was developed. In this, a single biomaterial is developed into a scaffold with varied porosity, stiffness, micro/nanostructure, and growth factor amount along the scaffold length to support bone, cartilage, and bone–cartilage interface in one integrated scaffold. For example, a graded scaffold with variable TGFβ1 and BMP-2 along the scaffold length showed high-quality cartilage regeneration of appropriate thickness and high GAG content. Additionally, there was proper bone ingrowth and integration of the neocartilage to both the underlying bone and surrounding cartilage [170]. More recently, Nukavarapu et al. developed a porosity gradient scaffold system, in which the scaffold with gradient pore volume is formed via particle sintering and porogen leaching method [4,139,171,172]. The template gradient scaffold was further combined with a hydrogel phase to form inverse gradient matrix, in which the polymeric structure and the infused gel supported osteogenesis and chondrogenesis, respectively, in vitro [63,173]. Current efforts are toward the development of an integrated scaffold with microstructural and stiffness variation along the length. This is to achieve stiffness or local microstructure–mediated stem cell osteogenic or chondrogenic differentiation. These efforts are mainly intended to develop growth factor–free approaches for osteochondral tissue engineering.

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11.4  Future perspectives In the past decades, cartilage regeneration efforts have come a long way—from simply implanting chondrocytes on their own to seeding them on advanced scaffolds engineered from different materials and with intricate architecture. With the rise of 3D printing and other technologies to create scaffolds, research studies are attempting to develop scaffolds with a highly organized structure, similar to the unique zonal organization of articular cartilage. Additionally, scientists are creating new types of copolymers and experimenting with new ways to modify materials to create the ideal scaffold material. Table 11.3 summarizes some of these more recently investigated cartilage scaffolds. Much research is being done outside of the scaffold itself, including exploring superior cell sources and additive components. The choice of the relevant cell population for cartilage tissue engineering still remains a large challenge. The difficulty lies with the dedifferentiation of primary chondrocytes during in vitro culture, which is necessary to produce the requisite number of cells needed for implantation. Also, the most readily accessible progenitor cell population, bone marrow MSCs, may differentiate into chondrocytes with the right treatment. However, in extended culture conditions and implantation, the cells often progress toward hypertrophy, which can lead to mineralization on the neocartilage and worsen the original condition. Although materials such as hydrogels have helped with maintaining chondrocyte morphology and therefore phenotype, creating cartilage-like extracellular environment to retain cells in the same phenotype over a long period of time is needed. One solution that is being explored is culturing the cells in vitro in a 3D spheroid or pellet environment before being placed in the 3D scaffold environment in the body. Other efforts include recreating the physiological conditions, including load, which chondrocytes experience in the body in vitro with advanced bioreactors. These specially engineered bioreactors offer a controlled environment with specific pH, temperature, CO2 level, and amount of mechanical stress, such as hydrostatic pressure or shear stress. Obtaining donor chondrocytes is limited and can lead to donor site morbidity, so more abundant cell types such as bone marrow–derived MSCs (BMSCs) and adipose-derived stem cells (ASCs) have been increasingly studied for their chondrogenesis and repair ability. Although BMSCs differentiate into chondrocytes more easily, ASCs are an attractive source because they are plentiful, can be harvested in large amounts with low donor site morbidity, and are easily isolated [174]. Furthermore, these cells actually showed enhanced chondrogenic potential after expansion in vitro, whereas it is decreased in BMSCs [175,176]. Although growth factors have been discussed, there are further questions on the best amount and type of growth factor to be used. An interesting issue that might arise is that different growth factors may be optimal for different cell lines, as it has been shown that ASCs respond better to BMP-6, whereas BMSCs respond better to TGF-β [177]. In the future, researchers will explore alternative ways to supplement cell culture both in vitro and in vivo. This includes coculturing cells so that “teacher” cells can communicate with the target cells by direct cell–cell contact or soluble factors to guide differentiation, using platelet-derived cytokines, or even DNA segments that include regulatory genes [174]. Within the field of cartilage tissue engineering, there are a myriad of areas to be explored to improve cartilage repair. In the past, research teams may have considered just one component of the scaffold, whether it was the cells, the architecture of the

Table 11.3  Recently developed cartilage tissue engineering scaffolds. Study

Scaffold

Evaluation

Results

[133]

Sodium hyaluronate and sodium alginate interpenetrating polymer network scaffold combined with berberine

In vitro evaluation and in vivo knee osteochondral defect model in rats

• Berberine

[134]

Trilayer scaffold: 1. Poly(ethylene glycol)-diacrylate (PEGDA) and N-acrylyl 6-aminocaaproid acid (A6ACA) bottom layer biomineralized with CaP 2. Cryogel PEGDA middle layer with anisotropic pore architecture 3. Hydrogel PEGDA top layer Shape memory native and denatured collagen scaffolds

In vitro evaluation and in vivo subcutaneous implantation in immunodeficient mice

[135]

[136]

PLGA-based porous scaffold with tunable inner surfaces that were functionalized with aromatic aldehyde groups via PEG and with added fibronectin

In vitro evaluation and in vivo subcutaneous Sprague Dawley rat model and full-thickness knee articular cartilage defect model in New Zealand white rabbits In vitro evaluation and in vivo implantation into a full-thickness articular cartilage defect model in New Zealand white rabbits

helps to promote bone regeneration by upregulating the Wnt pathway and promoting osteogenic differentiation of BMSCs • After 8 weeks, the scaffold with berberine resulted in the highest histological score and regenerated thick, hyaline-like cartilage with a smooth surface • After 9 weeks of culture, the chondrocytes and hMSCs that were loaded into the top two layers did not migrate to the mineralized bottom layer and only deposited GAGs and collagen in those layers • When implanted, the scaffold recruited cells to the bottom layer • After 8 weeks, the implanted scaffold showed good differentiation between the bone and cartilage layers with cellular organization that mimics that of the different articular cartilage zones • Native

collagen promoted proliferation, adhesion, and redifferentiation of chondrocytes and matrix interaction better than denatured collagen • After 12 weeks, native collagen scaffolds repaired cartilage defects better than denatured collagen, resulting in cartilage comparable to normal cartilage • The

scaffold allowed for formation of spheroids in situ and surface-mediated gene transfection of TGF-β1 during cell–scaffold adhesion • In vitro, the formed spheroids promoted TGF-β1 expression for 10 days and underwent enhanced chondrogenic differentiation • After 8 weeks, defects were filled with new cartilage tissue with similar organizational features and cell volume and density to native cartilage and no fibrous tissue Continued

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Table 11.3 Continued Study

Scaffold

Evaluation

Results

[137]

1:3 sodium alginate and gelatin 3D printed porous scaffold with various printing conditions and with or without icariin

In vitro evaluation

• The

[138]

Trilayer HA hydrogel scaffold 1. Hydrogel with multiple vertical channels in bottom 2. Hydrogel with randomly oriented PLA nanofiber mesh in middle 3. Hydrogel with aligned PLA nanofiber mesh on top Gradient PLGA scaffold with HA hydrogel 1. Gradient PLGA matrix 2. HA hydrogel infusion 3. Establishing a reverse gradient structure

In vitro evaluation

[139] [63]

In vitro evaluation

porous scaffold allowed for good distribution of cells throughout and had no adverse effect on cell viability • The composite ink used to print the scaffolds showed good thermal stability and swelling effect • Icariin addition increased cell proliferation and differentiation • After 14 days, the different layers facilitated growth of cartilage with zonal organization of cells and composition of ECM • The superficial zone properly facilitated the growth of elongated cells, whereas the middle zone properly facilitated the growth of rounded cells

• The

bottom PLGA layer facilitated MSC differentiation into osteoblasts • The top layer with PLGA scaffold and gel facilitated MSC chondrogenesis • The intermediate zone facilitated both osteogenesis and chondrogenesis, resulting interfacial layer formation

scaffold, or the material. In the future, research teams need to combine these efforts to come up with the best regenerative solution.

11.5  Conclusions Many different scaffolding materials for the purpose of cartilage tissue engineering have been explored. These materials range from natural materials obtained from other organisms, found in cartilage ECM, or cartilage ECM itself. Although many of these natural

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materials are abundant, they present issues with material composition and batch-tobatch variation based on the source. Synthetic biomaterials or gels are more versatile as they can be manufactured limitlessly and can be tailored to have specific characteristics to fit the needs of the scaffold. However, these polymers need to be modified to have the required characteristics such as biocompatibility and mechanical stiffness/strength. Not all cartilage defects are created the same; they may vary in size, depth, location, and shape. Different materials offer different advantages, which should be considered when deciding which material to use for specific defects. An injectable hyaluronan hydrogel may be suitable for an irregular defect, whereas a stronger PLGA scaffold may be a better choice for treating an osteochondral defect. Recent trend is to design composite scaffold systems that consist of synthetic and natural material combinations with the ability to encapsulate cells and present with the required biological or mechanical signaling to form cartilage or bone or cartilage–bone interface. This chapter summarizes the use of many commonly used cartilage scaffold materials and introduces the different types of composite scaffolds/structures and scaffold additives that are gaining popularity. Although much progress has been made in terms of developing biodegradable scaffolds with a combination of materials and bioactive cues, the efforts continue to develop scaffold systems that are simple yet effective for cartilage and osteochondral tissue engineering.

Acknowledgments The authors thank support from NSF EFMA (#1640008) and NSF EFRI (#1332329). Dr. Nukavarapu acknowledges funding from SPARK and Research Excellence Program (REP) through the University of Connecticut. Dr. Nukavarapu also acknowledges funding support from the National Institute of Biomedical Imaging and Bioengineering of the National Institutes of Health (#R01EB020640).

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[152] C.J. Little, W.M. Kulyk, X. Chen, The effect of chondroitin sulphate and hyaluronic acid on chondrocytes cultured within a fibrin-alginate hydrogel, J Funct Biomater 5 (3) (2014) 197–210. https://doi.org/10.3390/jfb5030197. [153] J.L.C. van Susante, J. Pieper, P. Buma, T.H. van Kuppevelt, H. van Beuningen, P.M. van Der Kraan, et al., Linkage of chondroitin-sulfate to type I collagen scaffolds stimulates the bioactivity of seeded chondrocytes in vitro, Biomaterials 22 (17) (2001) 2359–2369. [154] C.-T. Lee, C.-P. Huang, Y.-D. Lee, Biomimetic porous scaffolds made from Poly(llactide)-g-chondroitin sulfate blend with poly(l-lactide) for cartilage tissue engineering, Biomacromolecules 7 (7) (2006) 2200–2209. https://doi.org/10.1021/bm060451x. [155] M.A. Slivka, N.C. Leatherbury, K. Kieswetter, G.G. Niederauer, Porous, resorbable, fiber-reinforced scaffolds tailored for articular cartilage repair, Tissue Eng 7 (6) (2001) 767–780. https://doi.org/10.1089/107632701753337717. [156] S.-H. Park, S.R. Park, S.I. Chung, K.S. Pai, B.-H. Min, Tissue-engineered cartilage using fibrin/hyaluronan composite gel and its in vivo implantation, Artif Organs 29 (10) (2005) 838–845. https://doi.org/10.1111/j.1525-1594.2005.00137.x. [157]  G.A. Ameer, T.A. Mahmood, R. Langer, A biodegradable composite scaffold for cell transplantation, J Orthop Res 20 (1) (2002) 16–19. https://doi.org/10.1016/ S0736-0266(01)00074-2. [158] R.F. LaPrade, J. Botker, M. Herzog, J. Agel, Refrigerated osteoarticular allografts to treat articular cartilage defects of the femoral condyles. A prospective outcomes study, J Bone Joint Surg Am Vol 91 (4) (2009) 805–811. https://doi.org/10.2106/JBJS.H.00703. [159] P.C. McCulloch, R.W. Kang, M.H. Sobhy, J.K. Hayden, B.J. Cole, Prospective evaluation of prolonged fresh osteochondral allograft transplantation of the femoral condyle: minimum 2-year follow-up, Am J Sports Med 35 (3) (2007) 411–420. https://doi. org/10.1177/0363546506295178. [160] B.C. Emmerson, S. Görtz, A.A. Jamali, C. Chung, D. Amiel, W.D. Bugbee, Fresh osteochondral allografting in the treatment of osteochondritis dissecans of the femoral condyle, Am J Sports Med 35 (6) (2007) 907–914. https://doi.org/10.1177/0363546507299932. [161] M. Demange, A.H. Gomoll, The use of osteochondral allografts in the management of cartilage defects, Curr Rev Musculoskel Med 5 (3) (2012) 229–235. https://doi. org/10.1007/s12178-012-9132-0. [162] A.L. Pallante-Kichura, A.C. Chen, M.M. Temple-Wong, W.D. Bugbee, R.L. Sah, In vivo efficacy of fresh vs. frozen osteochondral allografts in the goat at 6 months is associated with PRG4 secretion, J Orthop Res 31 (6) (2013) 880–886. https://doi.org/10.1002/ jor.22319. [163] J.M. Coburn, M. Gibson, S. Monagle, Z. Patterson, J.H. Elisseeff, Bioinspired nanofibers support chondrogenesis for articular cartilage repair, Proc Natl Acad Sci USA 109 (25) (2012) 10012–10017. https://doi.org/10.1073/pnas.1121605109. [164] G. Chen, T. Sato, J. Tanaka, T. Tateishi, Preparation of a biphasic scaffold for osteochondral tissue engineering, Mater Sci Eng C 26 (2006) 118–123. https://doi.org/10.1016/j. msec.2005.07.024. [165] L. Shen, J. Wu, X. Liu, D. Chen, L. Bowlin Gary, L. Cao, et al., Osteochondral regeneration using an oriented nanofiber yarn-collagen type I/hyaluronate hybrid/TCP biphasic scaffold, J Biomed Mater Res 103 (2) (2014) 581–592. https://doi.org/10.1002/ jbm.a.35206. [166] A.M.J. Getgood, S.J. Kew, R. Brooks, H. Aberman, T. Simon, A.K. Lynn, N. Rushton, Evaluation of early-stage osteochondral defect repair using a biphasic scaffold based on a collagen-glycosaminoglycan biopolymer in a caprine model, Knee 19 (4) (2012) 422–430. https://doi.org/10.1016/j.knee.2011.03.011.

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[167] J. Jiang, A. Tang, G.A. Ateshian, X.E. Guo, C.T. Hung, H.H. Lu, Bioactive stratified polymer ceramic-hydrogel scaffold for integrative osteochondral repair, Ann Biomed Eng 38 (6) (2010) 2183–2196. https://doi.org/10.1007/s10439-010-0038-y. [168]  J. Gao, J.E. Dennis, L.A. Solchaga, A.S. Awadallah, V.M. Goldberg, A.I. Caplan, Tissue-engineered fabrication of an osteochondral composite graft using rat bone marrow-derived mesenchymal stem cells, Tissue Eng 7 (4) (2001) 363–371. https://doi. org/10.1089/10763270152436427. [169] T.J. Levingstone, A. Matsiko, G.R. Dickson, F.J. O’Brien, J.P. Gleeson, A biomimetic multi-layered collagen-based scaffold for osteochondral repair, Acta Biomater 10 (5) (2014) 1996–2004. https://doi.org/10.1016/j.actbio.2014.01.005. [170] N. Mohan, N.H. Dormer, K.L. Caldwell, V.H. Key, C.J. Berkland, M.S. Detamore, Continuous gradients of material composition and growth factors for effective regeneration of the osteochondral interface, Tissue Eng 17 (21–22) (2011) 2845–2855. https://doi. org/10.1089/ten.tea.2011.0135. [171] A.R. Amini, D.J. Adams, C.T. Laurencin, S.P. Nukavarapu, Optimally porous and biomechanically compatible scaffolds for large-area bone regeneration, Tissue Eng 18 (13–14) (2012) 1376–1388. https://doi.org/10.1089/ten.TEA.2011.0076. [172] D.L. Dorcemus, S.P. Nukavarapu, Novel and unique matrix design for osteochondral tissue engineering, MRS Proc (1621) (2014) 17–23. https://doi.org/10.1557/opl.2014.285. [173] S. Majumdar, P. Pothirajan, D. Dorcemus, S. Nukavarapu, M. Kotecha, High field sodium MRI assessment of stem cell chondrogenesis in a tissue-engineered matrix, Ann Biomed Eng 44 (4) (2016) 1120–1127. https://doi.org/10.1007/s10439-015-1382-8. [174] F. Hildner, C. Albrecht, C. Gabriel, H. Redl, M. van Griensven, State of the art and future perspectives of articular cartilage regeneration: a focus on adipose-derived stem cells and platelet-derived products, J Tissue Eng Regenerat Med 5 (4) (2011) e36–51. https://doi. org/10.1002/term.386. [175] B.T. Estes, A.W. Wu, R.W. Storms, F. Guilak, Extended passaging, but not aldehyde dehydrogenase activity, increases the chondrogenic potential of human adipose-derived adult stem cells, J Cell Physiol 209 (3) (2006) 987–995. https://doi.org/10.1002/ jcp.20808. [176] V. Vacanti, E. Kong, G. Suzuki, K. Sato, J.M. Canty, T. Lee, Phenotypic changes of adult porcine mesenchymal stem cells induced by prolonged passaging in culture, J Cell Physiol 205 (2) (2005) 194–201. https://doi.org/10.1002/jcp.20376. [177] B.O. Diekman, C.R. Rowland, D.P. Lennon, A.I. Caplan, F. Guilak, Chondrogenesis of adult stem cells from adipose tissue and bone marrow: induction by growth factors and cartilage-derived matrix, Tissue Eng 16 (2) (2010) 523–533. https://doi.org/10.1089/ten. tea.2009.0398. [178] W.J. Li, H. Chiang, T.F. Kuo, H.S. Lee, C.C. Jiang, R.S. Tuan, Evaluation of articular cartilage repair using biodegradable nanofibrous scaffolds in a swine model: a pilot study, J Tissue Eng Regenerat Med 3 (1) (2008) 1–10. https://doi.org/10.1002/term.127. [179] W. Guo, X. Zheng, W. Zhang, M. Chen, Z. Wang, C. Hao, Q. Guo, Mesenchymal Stem Cells in Oriented PLGA/ACECM Composite Scaffolds Enhance Structure-Specific Regeneration of Hyaline Cartilage in a Rabbit Model, Stem Cells Int 2018 (2018). 6542198. https://doi.org/10.1155/2018/6542198. [180] D. Xue, Q. Zheng, C. Zong, Q. Li, H. Li, S. Qian, B. Zhang, L. Yu, Z. Pan, Osteochondral repair using porous poly(lactide‐co‐glycolide)/nano‐hydroxyapatite hybrid scaffolds with undifferentiated mesenchymal stem cells in a rat model. J Biomed Mater Res, Part A, 94A(1), 259-270. https://doi.org/10.1002/jbm.a.32691.

Scaffolds for skeletal muscle tissue engineering

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Joseph Bartolacci1,2, Jenna Dziki1,3, Stephen F. Badylak1,2,3 1McGowan Institute for Regenerative Medicine, University of Pittsburgh, Pittsburgh, PA, United States; 2Department of Bioengineering, University of Pittsburgh, Pittsburgh, PA, United States; 3Department of Surgery, University of Pittsburgh, Pittsburgh, PA, United States 

12.1  Scaffolds for skeletal muscle engineering Volumetric skeletal muscle loss (VML) as a result of traumatic injury is a significant clinical problem in both military and civilian populations with an estimated annual economic burden of $4 billion [1]. Skeletal muscle has robust inherent regenerative capacity. However, defects greater than 20% of total muscle mass within a given muscle group overwhelm the ability of resident stem/progenitor cells to fully reconstitute the absent tissue and, instead, fibrotic tissue fills the void [2]. Fibrosis negatively impacts muscle function and is a significant factor in the morbidity and reduced quality of life associated with VML [3,4]. Current therapeutic approaches include rigorous physical therapy, use of orthotics, and/or autologous flap grafting. Autologous flap grafting often has limited donor site availability, can incur donor site morbidity, and has limited success in restoring appreciable function [5]. Tissue engineering and regenerative medicine strategies to address VML have been primarily cell-centric. Exogenous delivery of stem cells to repopulate the missing muscle tissue has shown moderate success in preclinical studies [6,7]. However, technical, economic, and regulatory hurdles, including low engraftment efficiency and low viability after injection, have limited the success of clinical translation [8–10]. In addition, cell isolation and maintenance requirements are costly, and allogeneic cell sourcing necessitates concomitant immunosuppression [11]. An acellular approach to VML is an attractive therapeutic alternative to cell-based strategies, and biomaterials engineered specifically to facilitate functional skeletal muscle regeneration have received considerable attention in recent years [12–15]. This chapter will focus only on biomaterial-based strategies for VML. Several approaches have been studied in preclinical animal models that have focused on novel biomaterials, with or without a cellular component, to treat nonregenerating muscle injury. An understanding of the host response to both skeletal muscle tissue injury and the biomaterials themselves is a prerequisite to strategies for functional muscle replacement therapy.

12.1.1  Response of skeletal muscle to injury The skeletal muscle response to injury consists of a series of temporally and spatially controlled cellular events dominated by the host innate immune system and local Handbook of Tissue Engineering Scaffolds: Volume One. https://doi.org/10.1016/B978-0-08-102563-5.00012-5 Copyright © 2019 Elsevier Ltd. All rights reserved.

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stem/progenitor cells. Immediately after injury, platelet activation occurs in response to contact with extravascular matrix molecules such as fibronectin and collagen. Platelet activation releases peptide fragments, growth factors, and chemokines that recruit circulating leukocytes, predominantly neutrophils, within minutes to the site of injury where they assist in clearing foreign material, necrotic debris, and bacterial contamination [16]. Fibrinogen is subsequently activated to fibrin to form the provisional blood clot [17,18]. Cytokines and chemokines released by neutrophils, as well as matricryptic peptides from the surrounding degrading extracellular matrix, promote recruitment of circulating monocytes and tissue-resident macrophages within 1–3 days of the injury [16]. Macrophages may persist at the site of injury until debris has been removed and healing is complete, a period of time that can last as long as 1 year. Infiltrating monocytes initially become activated to a proinflammatory, “M1-like” macrophage phenotype that is characterized by increased expression of inflammatory cytokines such as TNF-α and IL-1β, the generation of reactive oxygen species, and a paracrine-mediated increase in satellite cell proliferation. The inflammatory phenotype gradually transitions to a proremodeling, “M2-like” macrophage phenotype [19] that is characterized by lower production of reactive oxygen species, TGF-β and IL-10 secretion, increased arginase production, and a paracrine-mediated increase in myogenic precursor differentiation and matrix maturation. An appropriately timed macrophage transition from an M1-like to an M2-like phenotype is not just sufficient, but required, for functional repair of injured skeletal muscle [20]. Of relevance to this chapter, an increased M2:M1 ratio to a biomaterial implant during the acute host response phase is a predictor of downstream success or failure [12,21]. The sequential events of the healing response provide many potential targets for therapeutic intervention. The ideal scaffold for skeletal muscle engineering should promote an appropriately timed macrophage phenotype transition, stimulate angiogenesis to support the requirements of tissue regeneration, and serve as a provisional matrix or substrate for host tissue integration, endogenous cell migration, proliferation, and differentiation during the remodeling process. Several biomaterial-based strategies have been investigated to reconstitute injured or missing soft tissue and restore lost function. These materials can be broadly categorized into either synthetic or biologic scaffolds, with or without a cellular component. Representative commercially available materials that have been used for soft tissue repair are presented in Table 12.1. This chapter will review such materials as they have been applied to skeletal muscle reconstruction, the value and shortcomings of these approaches, and potential future directions for next-generation biomaterials.

12.2  Synthetic scaffolds 12.2.1  Nondegradable synthetic scaffolds Nondegradable synthetic scaffolds are the most commonly used surgical meshes for ventral hernia repair. The use of such materials has the intended function of mechanical reinforcement of a defect area rather than the induction of new site-appropriate

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Table 12.1  Commercially available materials [22–30]. Trade name

Scaffold material

Provider

Polypropylene ePTFE PET

CR BARD Ethicon Ethicon

PLGA-PTMC–PLA-PTMC PGA PGA-PTMC Poly-4-hydroxybutyrate

Novus Scientific Ethicon Gore CR BARD

Porcine dermal ECM Porcine small intestine submucosa Human dermal ECM

CR BARD Cook CR BARD

Polypropylene, hyaluronic acid PET, collagen I

CR BARD Cook

Nondegradable synthetics BARD Mesh DualMesh Mersilene

Biodegradable synthetics TIGR Vicryl GORE BIO-A PHASIX

Biologic scaffolds XenMatrix Surgisis Allomax

Composites Ventralight ST Parietex

tissue formation [31]. Robust mechanical properties and low cost have contributed to their widespread acceptance by the surgical community and have reduced the rate of ventral hernia recurrence dramatically [32,33]. The use of materials such as polypropylene, expanded polytetrafluoroethylene, and polyethylenepterepthalate (PP, ePTFE, and PET, respectively) has provided considerable insight regarding the host response to biomaterials [34,35]. Nondegradable synthetic materials are more susceptible to infection, a chronic inflammatory response at the site of implantation, tissue erosion, and fistula formation, among other complications than other classes of biomaterials [34,36]. The chronic inflammatory reaction to these tissues is referred to as the foreign body reaction (FBR) and is characterized by proinflammatory “M1-like” macrophages, multinucleate giant cells, and deposition of dense fibrous connective tissue [24,37]. Attempts have been made to reduce the inflammatory reaction through the use of drug-eluting polymers. Polypropylene scaffolds that elute IL-4 have been investigated for their ability to promote enhanced functional remodeling in a mouse model of muscle injury [38]. Similarly, reports have shown that coating PP mesh with ECM hydrogel prevented sustained M1 macrophage activity that is typically found with PP and facilitated host integration and repair [39]. Surface modification strategies have been developed that directly link cytokine [40] or nucleic acid–containing nanoparticles to the surface [41–43]. Genes encoding IL-4 [44], IL-10 [45], IκBα [46], or decoy TNF-α receptors [47] have been delivered using scaffolds to promote a local M2-like macrophage phenotype; however, more work is needed to determine the viability of this approach. It is important to note that surface modification techniques are not uniformly effective, the immunomodulatory effects of surface-active agents can be short lived, and pharmacokinetics of long-term drug release can be challenging [48].

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12.2.2  Biodegradable polymeric materials Many biodegradable polymers are currently in clinical use as scaffolds for ventral hernia repair, although a growing body of work has shown that these materials are also valuable tools for studying skeletal muscle regeneration in vitro and in preclinical animal testing [41,48–53]. Among the most commonly used polymers of this category are polyglycolic acid, polylactic acid, and poly-ε-caprolactone (PGA, PLA, and PCL, respectively). Importantly, the host macrophage response elicited by degradable synthetic materials consists of an M1-like reaction that is less sustained than that occurs with nondegradable synthetic scaffolds such as PP [48]. Recent preclinical data have shown that PCL microspheres can serve as a drug elution mechanism with the potential to provide long-term protection from infection [54], and steroid-eluting polymers have been developed that suppress the inflammatory reaction elicited by polymeric materials [55]. Furthermore, biodegradable materials composed of naturally occurring molecules have been developed, such as citrate-based polyesters for which the degradation products promote an M2-like macrophage phenotype [26,56]. Degradable polymers are currently an active field of investigation of materials that can be used to direct the replacement or reconstruction of host tissues. The temporary nature of degradable polymers makes them attractive materials for temporally controlled drug delivery. Resorbable microspheres with tunable degradation rates have been used to achieve long-duration growth factor release in preclinical animal models [1,57,58]. PLG microspheres, for example, have been investigated as a delivery mechanism for a range of growth factors. Results of these studies have shown that the released growth factors can be temporally controlled to sustain angiogenesis and vessel maturation [57,59]. Drug-eluting microspheres have also been used in conjunction with biologic polymers such as alginate, collagen, and fibrin, among others, to achieve functional restoration in animal models of volumetric muscle loss. One study found that the use of IGF-1 and VEGF-eluting PLG microspheres embedded in alginate hydrogels promoted early vasculogenesis and a significant increase in regenerating myotubes, resulting in near-complete functional recovery in a rodent model of ischemic muscle injury [51]. Determining the correct combinations, dosages, and dose regimens of growth factors and other bioactive molecules to obtain effective results is an active area of investigation [60]. Other studies have evaluated vascular networks that were manufactured in vitro using three-dimensional fibrin-PGA/PLLA seeded with endothelial cells, fibroblasts, and myoblasts. Results showed that the engineered constructs were capable of supporting blood flow in vivo [61], but functional testing was not performed. Cell-seeded biodegradable scaffolds, however, are plagued by poor cell survival following transplantation because of a lack of a vascular supply [62,63], costly and inefficient in vitro preparation times [9,64], and immunosuppression depending on the cell source [10].

12.2.3  Biologic scaffolds Biologic scaffolds are defined as scaffolds derived from organic sources such as plant and animal tissues, including ECM bioscaffolds and scaffolds derived from plasma

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proteins such as fibrinogen and fibronectin, or individual ECM components such as collagen I. Biologic scaffold materials have an inherent influence on cell behavior. For example, ECM bioscaffolds have been shown to promote site-appropriate stem cell differentiation and an M2-like macrophage phenotype. These properties have been observed in vitro, in preclinical animal studies, and in clinical settings where ECM bioscaffolds have been used for soft tissue repair [21]. When properly prepared (i.e., thoroughly decellularized and not chemically cross-linked), ECM bioscaffolds have been shown to promote a constructive host immune response and provide a ligand landscape that promotes functional host tissue ingrowth [65]. Importantly, ECM bioscaffolds have been shown to promote an appropriately timed macrophage phenotype transition away from the initial proinflammatory, “M1-like” phenotype, toward a proremodeling, “M2-like” macrophage phenotype [12,66]. This transition has been shown to be predictive of downstream remodeling outcomes [65]. The immunomodulatory activity associated with ECM bioscaffolds is due in part to retained cytokines and growth factors such as IL-4, as well as miRNA contained within matrix-bound nanovesicles (MBV) bound to ECM bioscaffolds [67,68]. It is also important to note that various matricryptic peptides released during ECM bioscaffold degradation have bactericidal properties [69]. A critical requirement for functional, engineered tissue is integration of newly deposited tissue with the host vascular supply [70]. ECM bioscaffolds have been shown to promote angiogenesis in vitro, in preclinical animal models, and in human patients. At least some of this angiogenesis appears to be caused by retained growth factors such as VEGF [71,72]. ECM bioscaffolds seeded with mesenchymal stem cells (MSCs) have been shown to promote greater vascularization and superior downstream functional outcomes compared with unseeded ECM bioscaffolds [73]. The seeded MSCs did not differentiate into skeletal muscle fibers, and the mechanism by which they promote increased muscle function remains an active area of investigation. ECM bioscaffolds promote stem/progenitor cell recruitment, survival, and differentiation [66]. Biologic scaffolds such as alginate, fibrin, and collagen, which lack endogenous growth factors, must rely on controlled release of added growth factors to promote a similar degree of angiogenesis [49,74–76]. Nonetheless, recent reports have shown that fibrin hydrogels were capable of supporting bundles of functional muscle tissue 1 cm thick for nearly 40 days in vitro because of the formation of a vascular network [77], but did not test whether the vascular network could integrate with host circulatory system in vivo. Other groups have shown that in vitro 3D muscle constructs had evidence of host circulatory system integration as early as 1-week postimplantation in a rat model [78]. In vitro studies suggest that retained growth factors and matricryptic peptides generated during ECM degradation are potent chemotactic agents that recruit endogenous stem/progenitor cells to the site of scaffold placement [79]. More recently, a 13-patient cohort study using ECM bioscaffolds to treat VML resulted in significantly increased in muscle volume and the associated migration of perivascular stem cells away from their microenvironmental niche [80]. When cells have been seeded on ECM bioscaffolds in vitro, spontaneous differentiation of myoblasts to myotubes has been observed [14,73], and innervation of new muscle fibers has been shown in vivo [81]. Innervation

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is a requirement for functional restoration and maintenance of muscular regeneration. The need for innervation is obvious from in the muscular atrophy that follows denervation [82]. Cell migration may be limited over large defects, and ECM bioscaffolds may benefit from seeding with exogenous cells for the repair of large defects [83]. Hybrid materials that incorporate ECM or ECM-mimetic components on a modified synthetic surface have been investigated for this ability to enhance cell migration [39,84]. Stem/progenitor cell–seeded scaffolds have been used to overcome limitations in native cell migration [2,73,83,85]. Collagen and fibrin materials seeded with myoblasts have reported moderate success in animal models, showing that cells can migrate out of the gels and integrate with host musculature [13,52,86] affording nearly 55% functional recovery in a rat model of VML. Cell-seeded constructs, however, are typically plagued by poor cell survival following transplantation because of a lack of vascular supply, costly and inefficient in vitro preparation times, and immunosuppression depending on the cell source.

12.2.4  Closing remarks No two injuries or patients are identical; therefore, the biomaterials utilized for muscle repair should be selected following evaluation of the clinical context. Chemical makeup of the scaffolds is not the only parameter to be considered in the development of tissue engineering scaffolds. Physical properties such as elasticity, mechanical strength, pore size, isotropy, and fiber size must also be considered. Stated differently, it is unlikely that a single scaffold will be applicable in clinical scenarios. Successful tissue engineering strategies will likely require a variety of materials with different biological, mechanical, and chemical properties (Table 12.2).

12.3  Cell types for skeletal muscle tissue engineering Results of preclinical work show that some cell-seeded scaffolds can provide more significant gains of function in skeletal muscle regeneration than acellular materials [14,70,83,91]. The use of different cell types, different animal models, and different analyses has yielded conflicting results, however. The cell types used in preclinical

Table 12.2  Comparison of bioactivity by material category. Nondegradable synthetic scaffolds

Degradable synthetic scaffolds

Biologics

No [39,87] No [35] No [38]

Variable [48,56] Variable [49,74–76] Yes [40,49,76,84]

Yes [88,89] Yes [72,90] Yes [12,79,90]

Properties Immunomodulation Angiogenesis Myofiber formation/ reinnervation

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animal models do not have uniform characteristics, and the ability of these cells to promote muscle repair varies widely, as can be seen in Table 12.3. It is helpful to understand these differences to better compare results and guide future strategies for skeletal muscle tissue engineering. Table 12.3  Cell types for skeletal muscle tissue engineering. Cell type

Characteristics

Advantages

Barriers to utility

Satellite cells [86,93]

Pax7+ Adult stem cells Necessary for muscle homeostasis Adult stem cells Found near muscle microvasculature Express satellite cell markers

Resident myogenic precursor

Lose myogenic potential after few passages Low engraftment efficiency Can be difficult to scale in vitro culture

Perivascular stem cells [8,95,96]

Muscle-derived stem cells [6,7,34,86,97]

Adult stem cells

Embryonic stem cells [11,62,98–100]

Pluripotent cells derived from fetal blastocyst

Induced pluripotent stem cells [62,101–103]

Genetically reprogrammed adult somatic cells with pluripotent potential

Phase I/II clinical trial using PVSCs for pediatric muscular dystrophy Can be passaged ∼20 times while retaining myogenic potential Better engraftment efficiency than satellite cells Improve muscle function and morphology Can be passaged ∼ 30 times while retaining myogenic potential Preclinical evidence suggests improved remodeling outcomes (histologically) Can generate large quantities of cells in vitro Improved engraftment efficiency in preclinical models

Can generate large quantities of autologous cells in vitro Pax7+ and muscle precursors have been established in vitro Promote skeletal muscle regeneration and functional improvements in preclinical animal models

Very low engraftment efficiency No functional improvements

Ethical concerns Difficult to recapitulate the skeletal muscle lineage in vitro Allogeneic cell source Potential cancer risk Time-consuming genetic reprogramming

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Most cell types that have been investigated for skeletal muscle repair are autologous or allogeneic stem cells that have been isolated and expanded in vitro before seeding and reimplantation. Autologous cells are advantageous because there is no immune-mediated rejection, even in the absence of immunosuppression. Allogeneic cells require prolonged immunosuppressive therapy [10] that can adversely affect healing, foster fibrosis [85], and disrupt constructive macrophage phenotype transitions [92]. Autologous, induced pluripotent stem cells and allogeneic embryonic stem cells can theoretically be passaged indefinitely [9], affording much larger populations of cells for seeding compared with satellite cells, which are very limited in their passage potential [93]. Cell survival is a concern as it has been shown that the vast majority of transplanted cells will fail to integrate with host tissue. Low engraftment efficiency is multifactorial [8], but a lack of a vascular supply and an inflammatory host environment have been implicated as key contributors to this phenomenon [94]. The number of cell layers that can be seeded, for example, is limited by the ability of nutrients and gases to diffuse from a medium to the cells. Specifically, in the absence of a vascular network, cells in excess of 1 mm distance from a nutrient- and gas-rich medium will have difficulty surviving [63]. Nutrient diffusion presents a particularly challenging obstacle for cell-seeded scaffolds that lack a vascular network. Attempts have been made to design scaffolds with provisional vascular conduits [61], but insufficient preclinical animal modeling has been conducted and is an ongoing topic of research. Furthermore, induced pluripotent stem cells (iPSCs) require complex growth factor cocktails to support appropriate differentiation, often have low viability, and there is concern surrounding the retroviruses used to induce pluripotency [62]. An appropriate level of safety must be considered, before large-scale clinical application, which in itself can be particularly challenging because no technology exists that can sterilize a scaffold without killing embedded cells. Stated differently, all steps in the manufacture of a cell-seeded scaffold must be conducted under sterile conditions, which adds time, cost, and potential risks to the development of these personalized devices. For these stated reasons, exogenously supplied autologous and allogeneic stem cells have achieved limited success.

12.4  Conclusions and future directions Skeletal muscle injury, especially volumetric muscle loss, remains an unmet medical need, making a regenerative medicine strategy to address skeletal muscle injury particularly attractive. There is no consensus to date on the most appropriate strategy to pursue. The physiology underlying muscle healing remains an active field of investigation, and new knowledge will be actively incorporated into the development of novel biomaterials and cell-based approaches. As safety and efficacy are proven for cell-based strategies, regulatory hurdles that have prevented more widespread clinical translation can be appropriately modified and the clinical translation of innovative regenerative medicine approaches can be expanded.

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Héctor Capella-­Monsonís1,2,a, Salomé Guillaumin1,2,a, Sofia Ribeiro1,3,a, Stefanie Korntner1,2, Yves Bayon3, Dimitrios I. Zeugolis1,2 1Regenerative, Modular & Developmental Engineering Laboratory (REMODEL), National University of Galway Ireland (NUI Galway), Galway, Ireland; 2Science Foundation Ireland (SFI) Centre for Research in Medical Devices (CÚRAM), National University of Galway Ireland (NUI Galway), Galway, Ireland; 3Medtronic, Sofradim Production, Trevoux, France

13.1  Introduction Tendon and ligament damage are the most frequently reported soft tissue injuries in the orthopedic field, placing an enormous burden on health-­care systems worldwide. In particular, athletes and the elderly population are predominantly affected by tendon ailments. Injury treatment either involves conservative approaches (e.g., eccentric training, pain management) or requires surgical intervention, which is often associated with long-­term pain, compromised range of motion, and an increased risk of recurrent ruptures [1–4]. Native tendon tissue exhibits a hierarchically organized structure with densely packed and highly aligned collagen fibrils and fibers, allowing for an accurate transmission of musculoskeletal forces, facilitating a wide range of joint motion [5]. On account of its high specificity, tendon represents a bradytrophic tissue with a very limited tissue turnover. A low number of active tissue-­resident cells together with poor vascularization and innervation result in a slow and incomplete healing response after injury, with the formation of a biomechanically inferior scar tissue [4,6]. Despite significant advancements in tissue engineering, no satisfying treatment is currently available to restore native tendon tissue, leaving an unmet clinical need for effective and reproducible treatment strategies. Although autografts remain the gold standard for tendon repair, allografts, xenografts, and synthetic materials are increasingly gaining interest to overcome limitations associated with autograft harvesting. However, autografts, allografts, and xenografts are associated with several complications, including donor site morbidity and immunoreaction [7]. Because tendon is of a relatively avascular nature, engineered tissue substitutes pose a promising treatment strategy to restore tissue structure and function [8,9]. One of the main challenges in scaffold-­based tissue engineering is to design and develop biocompatible and biodegradable scaffolds with optimal structural and mechanical properties. The final product should mimic the tissues’ native structure and function as close as possible. The structure of natural polymers allows for cell attachment even without surface modifications, provides mechanical support for cell a Héctor

Capella-­Monsonís, Salomé Guillaumin and Sofia Ribeiro share equally first authorship.

Handbook of Tissue Engineering Scaffolds: Volume One. https://doi.org/10.1016/B978-­0-­08-­102563-­5.00013-­7 Copyright © 2019 Elsevier Ltd. All rights reserved.

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growth, and provides a microenvironment that can regulate cell proliferation and differentiation, limiting cytocompatibility issues [10]. Because of a high modifiability, synthetic polymers became an attractive alternative to natural polymers. However, residual solvents from the fabrication process, as well as certain metabolites, can trigger inflammation or fibrous encapsulation after transplantation and other unwanted adverse effects [11,12]. Various two-­and three-­dimensional fabrication technologies have been utilized to develop nano-­and macrosized scaffolds with controllable and reproducible topographical and geometrical features to match the native tissue properties. Despite promising preclinical results, developed scaffolds still fail to fully recapitulate native tissue structure, mechanical properties, and overall composition of tendon [7,13–16]. This chapter provides an overview and critically discusses current biomaterial-­and tissue graft–based technologies for tendon tissue engineering.

13.2  Biomaterial-­based therapies Advancements in the field of biomaterial technologies have led to novel strategies for tendon tissue repair by designing three-­dimensional biomaterials that mimic native tendon structure and mechanical properties. Numerous natural and synthetic polymers have been investigated in tendon repair augmentation with variable degree of efficacy. Among them, collagen-­based devices are favored, considering the collagenous nature of tendon tissue. Functionalization strategies, based on biophysical, biochemical, and biological cues, aim to control cellular function and localization and sustained delivery of therapeutics/biologics to promote functional repair and regeneration [7,17]. Different scaffold fabrication methods are used for the generation of hierarchically structured, three-­dimensional scaffolds that closely imitate architectural features and mechanical properties of native tendon tissue, while allowing for localized and sustained delivery of therapeutic molecules. Herein, we discuss advantages and weaknesses of widely used scaffold fabrication technologies, including extrusion, lithography, and self-­assembly.

13.2.1  Electrospinning (ES) Because of its versatility and controllability, ES represents a promising technique to manufacture fibrous three-­dimensional scaffolds with controlled fiber diameter and alignment. The first record of a biomedical application using ES was published in 1978 [18]. Since then, numerous natural and synthetic polymers have been used to create electrospun scaffolds for tissue engineering applications targeting regeneration of bone [19], skin [20], cartilage [21], tendon [22], and nerves [23] (Table 13.1). Additionally, ES has been investigated toward applications in drug-­delivery systems [24,25]. In a simplistic way, ES is a technology that utilizes an electrical field to generate polymeric nonwoven fiber mats. As a bottom-­up approach, it has only minimal processing waste [25,35] and can be tailored to generate aligned nano-­and microfibers

Scaffold fabrication technique Extruded microfibers

Scaffold design

Cell type

In vitro/ in vivo

Collagen fibers

Ovine tenocytes

In vitro

Collagen fibers with incorporated DCN Collagen fibers embedded in collagen matrix

/

In vitro

Rat skin fibroblasts

/

Collagen-­ chondroitin-­6-­ sulfate scaffold reinforced with extruded type I collagen fibers Collagen fibers

/

In vitro

/

In vivo (sheep)

Outcomes

References

Higher cell proliferation and ECM production in EDC-­cross-­linked fibers compared to EGD-­ cross-­linked fibers Improved tensile properties of non-­cross-­linked fibers by DCN Similar mechanical properties of 125 μm diameter fibers and human ligaments, except viscoelastic properties. Higher tangent moduli and peak stress in cellularized scaffolds Cross-­linked scaffolds withstood a load of 60 N before failure and showed favorable pore matrix structure. Significant internal porosity and channels between fibers and matrix in non-­cross-­linked scaffolds

[8]

Extruded fibers showed inferior mechanical properties than native tendon. Better mechanical properties, better tissue ingrowth, and integration for EDC-­cross-­linked fibers compared with EGDE-­cross-­linking

[29]

Scaffolds for tendon tissue engineering

Table 13.1  In vitro and in vivo applications of extruded microfibers and electrospun fibers.

[26] [27]

[28]

261

Continued

Scaffold fabrication technique Electrospun fibers

Cell type

In vitro/ in vivo

Braided PLLA nanofibers Bilayered constructs of PCL nano-­and microfibers

Human MSCs

In vitro

Equine tendon fibroblasts

In vitro

PCL and PLA nanofiber yarn-­based woven biotextiles

Tricultures of human ADSCs, tenocytes, and umbilical vein endothelial cells /

In vitro

HA-­PELA/celecoxib–PELA bilayer biomimetic sheath Aligned and random-­ / oriented PLLA scaffolds

In vivo (chicken)

In vivo (rat)

Outcomes

References

Stimulated cyclic tensile strain leads to upregulated Scleraxis gene expression Scaffolds supported cell adhesion and proliferation. 2D aligned sheets and 3D bundles promoted parallel orientation of cells. 3D scaffolds exhibited best physical properties Triculture system promoted tenogenic differentiation by increasing collagen production, tendon-­associated protein, and gene expression

[30]

Outer celecoxib–PELA layer reduced inflammation and adhesion on tissue surroundings, whereas inner HA-­PELA layer released HA and promoted tendon gliding Tendonlike tissue formation with aligned fibrous scaffold; ectopic bone formation with randomly oriented fibrous scaffold

[33]

[31]

[32]

[34]

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Scaffold design

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Table 13.1  In vitro and in vivo applications of extruded microfibers and electrospun fibers.—cont’d

Scaffolds for tendon tissue engineering

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Figure 13.1  Scanning electron microscopy (SEM) (a) and rhodamine (red)/DAPI (blue) staining of human primary tenocytes seeded on electrospun polymeric fibers (b).

with tuneable mechanical and structural features that mimic the ECM architecture of native tissues [7,36] (Fig. 13.1). Electrospun scaffolds show high microporosity and large surface areas, which offer a suitable structure for growth and development of different types of cells. Tuning up the scaffold properties facilitates cell phenotype maintenance and allows for controlling stem cell fate in vitro [37]. These modifications are based on controlling fiber diameter by polymer solvent and concentration, voltage intensity, and distance between needle and collector [25,38]. Ultimately, fiber orientation can be controlled by optimizing architecture and rotation of the collector [15]. The technique allows for generating fibers with diameters ranging from less than 100 nm to a few μm. Because collagen fibril diameter in vivo ranges from 20 to 40 nm, this technique poses a suitable strategy for tendon tissue engineering applications [39]. Although electrospun collagen scaffolds have already been used in in vivo models, the process of ES can lead to the denaturation of natural polymers, such as collagen [40,41]. Consequently, ES using synthetic polymer fibers has been applied successfully in recent years. Several studies showed the impact of fiber organization on cellular response and matrix properties. Tenocytes cultured on aligned nanofibrous scaffolds attached parallel to the direction of the fibers, whereas cells seeded on scaffolds with random-­fiber orientation showed a disorganized morphology. Furthermore, tenocytes seeded on scaffolds with aligned electrospun fibers exhibited an organized collagen type I deposition with regard to fiber alignment, comparable with the ECM of native tendon [31,34,42,43], as well as a decreased osteogenic differentiation potential [44]. The major drawback of ES is the relatively poor mechanical properties of the obtained fibers. Generally, the mechanical properties of the scaffold are of the outmost importance, because mechanical loading is essential to maintain or regain normal tendon function [45]. However, several strategies have been reported to overcome the drawbacks and to improve the mechanical resilience of the material by knitting, twisting [22,31], and loading the scaffold with cellulose nanocrystals [46]. Moreover, bonding between electrospun sheets and a woven layer resulted in a 20-­fold stronger multilayered construct, capable of withstanding forces similar to the in vivo situation in human infraspinatus tendons [47].

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Cell-­loaded synthetic electrospun scaffolds have shown to favor tendonlike tissue formation of tenocytes from different sources, evidenced by morphological, histological, and biomechanical analysis [31,42,43,47–51]. Adipose-­derived stem cells (ADSCs) seeded in a PCL scaffold and coated with a tendon-­derived ECM resulted in improved cell attachment, increased gene expression of tendon-­related markers, and improved mechanical properties [52]. Furthermore, ADSCs loaded on a PLGA scaffold showed an upregulation of scleraxis in the presence of GDF-­5, a transcription factor critically involved in tendon development and healing, indicating enhanced tenogenic differentiation [53]. Furthermore, improved tenogenic differentiation of bone marrow–derived stem cells (BMSCs) has been described using electrospun scaffolds made of synthetic polymers, such as PLLA, PLGA, and PLGA in conjugation with silk [22,30,54,55]. More recently, it has been proven that aligned electrospun nanofibers in multilayered constructs support tenogenic differentiation by maintaining optimal cell morphology, increased tenogenic marker gene expression, and deposition of collagen and glycosaminoglycans [56,57]. ES can also be tailored toward applications for specialized tissue interfaces, such as the myotendinous junction. When a synthetic polymer, such as PLA or PCL, and a second natural polymer, such as collagen, were electrospun simultaneously, one uniform scaffold was produced. In vitro studies using this coscaffolds seeded with C2C12 myoblasts and NIH3T3 fibroblasts showed optimal cell attachment and enhanced myotube formation [58]. To investigate the impact of fiber alignment on tendon regeneration in vivo, the potential of PLLA electrospun scaffolds has been investigated using a rat Achilles tendon repair model. Immunohistochemical analysis showed increased tendonlike tissue formation compared with a random-­oriented fibrous scaffold [34]. After implantation of PLGA scaffolds loaded with autologous ADSCs in an Achilles tendon injury model in rabbits, the constructs gradually formed neotendon tissue and became more mature after 45 weeks with a histological structure similar to that of uninjured tendon [59]. In the presence of allogeneic BMCs, the scaffold showed good tissue integration and potential to promote wound repair, showing less lymphocyte infiltration and improved biomechanical properties [60]. Mice muscle-­derived cells (MDCs) were seeded on PGA fibers and sutured to the dorsal fascia muscularis in a mouse model in order that the movement of the limb provides mechanical stretching. The MDC-­engineered tendons showed abundant deposition of mature collagen, and their initial gene expression profile changed to a tenogenic profile [61]. In another study, electrospun PLGA fibers were separately cultured with porcine dermal fibroblasts and tenocytes for 7 days and subsequently implanted into a superficial digital flexor tendon defect in pigs. However, no differences were obvious between both conditions as both cell-­engineered tendon constructs showed similar tensile strength and good tissue integration. The authors concluded that fibroblasts could be a viable and more easily accessible cell source for tendon regeneration compared with tenocytes [62]. In a more complex approach, ADSCs and platelet-­derived growth factor (PDGF-­BB) were incorporated into a scaffold composed of electrospun PLGA fibers and a fibrin gel. Its potential as a delivery system was assessed in vivo using a canine flexor tendon injury model. The system proved to be a potential surgical approach for tendon healing by delivering cells and growth factors concomitantly [63].

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Even though ES has given a deeper understanding of tendon tissue engineering, all aspects of the obtained materials have yet to be fully understood. As mentioned previously, alignment of fibers is crucial for tendon regeneration; however, this paradigm has been questioned by a study where fiber diameter played a more crucial role in tenogenic differentiation of BMSCs compared with fiber orientation [64]. There is a report of reduced cell infiltration in nonwoven materials leading to limited matrix deposition, preventing these constructs to be used in broad clinical applications [56,65]. Consequently, the clinical potential of ES in tendon tissue engineering is still questionable. To date, four clinical trials with regard to “electrospinning” have been registered (ClinicalTrials.gov: NCT02255188, NCT02409628, NCT00428727, NCT00317629); however, none of these are related to tendon tissue. Companies have developed electrospun products mainly for filtration systems (e.g., HRV and United Air Specialists) and for tissue culture consumables (e.g., Nanofibre Solutions, SKE Advanced Therapies). Examples for electrospun biomedical devices are hernia meshes, such as NovaMesh and ReDura; however, these commercial products show a limited use because of their insufficient mechanical properties for functional tendon repair. Nicast Ltd has developed a vascular graft with CE Marking and clinical data published (AVflo), posing an enhanced primary graft over other products. ES is further utilized to produce medical devices, such as nonwoven polyurethane stents for acute coronary artery perforations (PK Papyrus, Biotronik International) and nerve conduits (Bioweb, NanoNerve Inc). ES is also used to transform polytetrafluoroethylene into various shapes and sizes (Zeus Inc.) [15]. Currently, there is no electrospun product for tendon repair commercially available. As ES is a relatively novel technique to be used in medical applications, electrospun medical devices are expected to be increasingly applied in the clinic in the near future.

13.2.2  Imprinting Imprinting technologies have recently attracted attention in biomedicine and tissue engineering, as they allow for the fabrication of two-­dimensional and three-­dimensional scaffolds with controllable and tuneable topographical and geometrical features [15]. Many recent developments have already given a deeper understanding of the effect of topographical cues on cellular behavior [66]. To study the effect of substrate topography in primary cells, such as osteocytes [67], fibroblasts [68], or endothelial cells [69], patterned substrates have been used and their influence on cellular adhesion, orientation, proliferation, and lineage commitment has been assessed. The effect of topography on cell differentiation has been extensively studied using polymeric patterned substrates seeded with stem cells, which have been differentiated toward the chondrogenic [70], adipogenic [71], tenogenic [72], and neural [73] lineage. Lithography technologies have surged with the demand for constructs with precise control of dimensional features down to nanoscale. To mimic the native tissue topography, studies have turned to microfabrication techniques producing structures such as grooves, pillars, pits, and wells [66]. One of the main features of imprint lithography is the precise creation of a topography on a wide range of materials without altering their intrinsic properties [74]. Optical lithography, or photolithography, makes use of a

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Figure 13.2  SEM (a) and immunocytochemistry of polymeric imprinted substrates stained for tenascin-­C (b).

light-­sensitive photoresist material to form a patterned coating on a surface. The substrate is coated with the photoresist and subsequently exposed to light using a mask with the selected features. The exposed material is either cross-­linked, polymerized, or degraded, and the substrate is then treated appropriately to obtain the selected areas [75]. Soft lithography uses elastomeric polymers to obtain patterns based on embossing, molding, and printing methods [66]. This set of techniques has many advantages because they are relatively low in cost and easy to use and allow for high-­throughput assays. Because no clean room is required, soft lithography grants a cost-­effective and easily accessible solution for many tissue engineering purposes. Initially, a master with a desired topography is produced using a rigid material. Subsequently, the master and a thermoplastic polymer film are placed under pressure and heated over the glass transition temperature. After gradually cooling down the temperature below glass transition, a patterned polymer film is obtained with feature sizes down to 5 nm [76,77]. Soft imprinted substrates have been used for approaches targeting tendon repair because lithography techniques are able to replicate the tendon topography with a similar size scale pattern (Fig. 13.2). A silicone membrane with 5 μm grooves was shown to maintain tenocyte morphology, resulting in elongated cells that were aligned in the direction of the grooves [78]. To further investigate the relationship between substrate topography and phenotype maintenance, porcine tenocytes were seeded on a microgrooved silicone membrane with features smaller than the size of the cells. This substrate promoted cell alignment and elongation and also maintained collagen type I and tenomodulin expression [79]. To investigate the effect of grooves larger than tenocyte dimensions, cells were seeded on glass substrates with a groove width of 50, 100, and 250 μm. The results showed that the topography still promoted the deposition of an aligned collagen matrix while losing its beneficial effect on both gene expression and cell phenotype maintenance [80]. Another approach to mimic tendon topography was the creation of direct tissue replicas using bovine Achilles tendons. An accurate PDMS replica of the tissue coated with collagen I and seeded with BMSCs was able to differentiate stem cells toward tenogenic lineage [72]. However, human tenocytes on microgrooved PLGA substrates

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lost their native phenotype, possibly owing to the rigidity of the substrate, and they failed to induce bidirectional neotissue formation in vivo [81]. Collectively, these results highlight the potential of imprinting technologies for in vitro cell phenotype maintenance and cell differentiation. However, future approaches should consider investigating both the effect of surface topography and substrate stiffness on cell behavior.

13.2.3  Hydrogels Hydrogels are macromolecular polymers suspended in water, composed of hydrophilic monomer chains, which form a network by cross-­linking. Hydrogels are fabricated using synthetic or natural polymers, and there are many different approaches to synthesize them [82]. The formation of a hydrogel can be triggered by physical and chemical parameters, such as temperature, pressure, electric field, pH, and ionic bonds [83]. Novel approaches aim to develop hydrogels with enhanced mechanical properties [84]. The fabrication of hydrogels for tendon repair can have different objectives. There are studies using either hydrogels alone [85], or functionalized hydrogels by the addition of molecules or cells, targeting cell differentiation toward the tenogenic lineage [86,87] (Table 13.2). Along these lines, a gelatin hydrogel sheet containing FGF-­2 was reported to stimulate endogenous tenogenic progenitor cells to differentiate into tenomodulin-­positive tenocytes after implantation in a rat rotator cuff injury model. The injury model was based on the detachment and subsequent surgical reattachment of the supraspinatus tendon via a drill hole in the humerus. Significantly improved mechanical properties, decreased vascularity, and increased collagen fiber orientation were observed in the FGF-­2 group, compared with the control group. Furthermore, the expression of tendon-­related markers Sox9, Scx, and Tnmd was increased in the FGF-­2 group in a time-­dependent manner [88]. In addition, collagen hydrogel systems have been investigated for strategies limiting fibrosis [89]. Cell alignment in hydrogels was shown to be improved after applying cyclic loading [90,91]. Using a gelatin hydrogel sheet, a study investigated the ability of cells to replace the scaffold by producing their own tissue-­specific ECM in vivo, once the hydrogel is degraded [92]. In vitro studies investigated natural hydrogels for tendon regeneration without the addition of any factors (i.e., cells or drugs). One study analyzed the alignment and electromechanical properties of rat tail tendons compared with dried isoelectrically focused type I collagen hydrogels. Despite similar piezoelectric properties, isoelectrically focused hydrogels showed threefold decreased fibril alignment compared with native rat tail tendons. The authors concluded that isoelectrically focused collagen could be used for tendon tissue engineering; however, covalent cross-­linking would need to be optimized [85]. A recent study combined an alginate hydrogel with clay nanoparticles for the controlled delivery of insulin-­like growth factor-­1 (IGF-­1) and investigated its performance in a rat Achilles tendon injury model [103]. Because inflammation is a critical factor for tendon regeneration, the downregulation of inflammatory factors such as cyclooxygenase (COX)-­1 and -­2 has been investigated. Using COX-­1-­and COX-­2-­engineered miRNA plasmid/nanoparticles trapped into a PLGA/ hyaluronic hydrogel, a decrease in tendon adhesion formation could be achieved [104].

Table 13.2  In vitro and in vivo applications of collagen hydrogens and lyophilized collagen scaffolds. Cell type

Molecule

In vitro/in vivo

Outcomes

References

Gelatin hydrogel

/

TGFβ-­1

In vivo (rat)

[86]

Alginate hydrogel

Human PDLSCs GMSCs BMSCs

TGFβ-­3

In vitro and in vivo (mouse)

Fibrin hydrogel

Rabbit BMSCs

In vitro

Gelatin hydrogel sheet

/

BMP12 BMP14 TGFβ-­3 VEGF FGF-­2

Inhibited MMP-­9 and MMP-­13 expression by TGFβ-­1 led to increased collagen accumulation. No tenogenic differentiation observed. In vitro: enhanced gene expression of Scx, Dcn, Tnmd, and Bgy (biglycan). In vivo: ectopic neotendon regeneration and greater regenerative capacity of periodontal ligament stem cells (PDLSCs) than gingival mesenchymal stem cells (GMSCs) or huBMSCs. Tenogenic differentiation of BMSCs most effective with a cocktail of BMP14, TGFβ, and VEGF.

[94]

Gelatin hydrogel sheet

/

BMP7

In vivo (rat)

Collagen hydrogel

Human tenocytes

DCN IL-­10

In vitro

Gelatin hydrogel

/

FGF-­2

In vivo (rat)

Collagen gel–sponge composite

Autologous rabbit BMSCs

/

In vivo (rabbit)

Dense tendonlike tissue formation in FGF-­2-­ impregnated gelatin hydrogel sheet group with improved biomechanical properties; no ectopic bone formation. Sustained release of BMP 7 resulted in enhanced tendon-­to-­bone healing with improved biomechanical properties. Codelivery of DCN and IL-­10 in a hydrogel decreased expression of profibrotic ECM genes via downregulation of TGFβ-­1. FGF-­2-­loaded hydrogel promoted tendon-­to-­bone healing with improved biomechanical and histological properties and upregulated expression of Sox9 of the rotator cuff. Improved biomechanical properties in cell-­loaded gel–sponge composites. Strong protein expression of type I and type V collagen, fibronectin, and decorin in both constructs.

[87]

[93]

[89]

[88]

[95]

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In vivo (rabbit)

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Scaffold type

Murine BMSCs

/

In vitro

Increased gene expression of type I collagen and increased linear stiffness after tensile stimulation compared with nonstimulated controls.

[96]

/

/

In vivo (rat)

[97]

Collagen sponge

/

GDF5 GDF6

In vivo (rat)

Knitted silk–collagen sponge

Human ESC– derived mesenchymal stem cells

/

In vitro and In vivo (mouse)

Collagen–silk sponge

Allogenous tendon stem/progenitor cells (TSPCs) /

/

In vivo (rabbit)

Accelerated Achilles tendon healing by type I collagen sponges via increased local type I collagen deposition and improved biomechanical properties compared with controls. GDF 5 and 6 encapsulated in collagen sponges enhanced tendon healing with increased tensile strength in a dose-­dependent manner compared with the sponge-­only group. In vitro: hESC–MSCs exhibited tenocyte-­like morphology and expressed tendon-­related marker genes after mechanical stimulation. Ectopic transplantation in vivo: regularly aligned cells and larger collagen fibers. Better histological scores and superior mechanical properties. Increased collagen deposition, better structural and biomechanical properties in TSPC-­seeded scaffold group compared with control group after rotator cuff injury.

SDF-­1α

In vivo (rat)

/

rhBMP-­12

Clinical trial (phase I)

Collagen/silk sponge

[99]

[100]

Improved tendon regeneration with increased tendon-­ [101] specific ECM production and decreased accumulation of inflammatory cells in bioactive scaffold group (loaded with local endogenous stromal cell-­derived factor-­1 alpha [SDF-­1α]). rhBMP-­12-­loaded collagen sponges proved to be safe [102] for use in open rotator cuff repair in a randomized controlled trial; level of evidence, 2.

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Collagen sponge (Pfizer Inc.)

[98]

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Collagen sponge (Kensey Nash Corporation, Exton, PA) Collagen sponge

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Most in vitro studies investigated the addition of cells and/or molecules to hydrogels. One of the factors responsible for fibrotic scar tissue formation is TGF-­β1. By a sustained delivery of decorin (DCN) and interleukin-­10 (IL-­10) via a type I collagen reservoir, suppressed expression of TGF-­β1 and other profibrotic ECM genes was reported in human tendon fibroblasts [89]. Furthermore, a tool for triggering the differentiation of rabbit BMSCs toward the tenogenic lineage has been developed by the association of bone morphogenetic protein (BMP) 14 with TGF-­β3 and vascular endothelial growth factor (VEGF) in a fibrin hydrogel [93]. Other studies focused on the effect of cyclic loading on cells seeded in type I collagen hydrogels, however, with controversial results [90,91]. Furthermore, investigations combining hydrogels with electrospun fibers showed promising results. A layered chitosan–collagen hydrogel/ aligned PLLA nanofiber construct was developed to mimic the ECM of native tendon [105]. A study used a photocross-­linked hydrogel fabricated by coelectrospinning of PCL and methacrylated gelatin (mGLT) combined with human ADSCs and TGF-­β3. Human ADSCs aligned in the direction of the fibers, whereas the addition of TGF-­β3 showed an upregulation of scleraxis and tenascin-­C, which are considered tenogenic markers [106]. Some in vivo studies exploited hydrogels combined with different therapeutic molecules. A gelatin hydrogel loaded with fibroblast growth factor-­2 (FGF2) has been used to induce differentiation of endogenous progenitor cells for tendon-­to-­bone healing in rat [88] and rabbit models [94]. The same authors developed a gelatin hydrogel sheet loaded with PDGF-­BB for tendon-­to-­bone healing in a rat model [107] as PDGF-­BB showed to promote healing by collagen remodeling and fibroblast proliferation [108]. Furthermore, TGF-­β1 was associated with a gelatin hydrogel for its ability to promote tenogenic differentiation of mesenchymal progenitor cells in a rat rotator cuff repair model [86]. To enhance matrix deposition in tenocytes, BMP-­7 was added to a gelatin hydrogel sheet [92]. Another experiment used TGF-­β3-­loaded RGD-­coupled alginate microspheres to which mesenchymal stem cells were added. TGF-­β3 facilitated differentiation of stem cells toward the tenogenic lineage in vitro and in vivo and resulted in the formation of ectopic neotendon-­like tissue in immunocompromised mice [87]. In the only in vivo study in a large animal model, a collagen hydrogel with ovine mesenchymal amniocytes was used to engineer a tendon substitute, which was further used for the treatment of diaphragmatic hernia in newborn lambs [109]. To date, no clinical trial has been reported using hydrogels for tendon repair nor there was any other product found on the market specifically for tendon. Commercially available hydrogels can be found for many different applications, such as wound healing; however, very frequently their composition (synthetic or natural hydrogel) is not clear.

13.2.4  Extruded microfibers Reconstituted collagen fibers are promising candidates for tendon tissue engineering, because collagen can be extruded into fibers with small diameters, which can be arranged in a parallel fashion to mimic the native tissue structure. Extruded collagen microfibers are fabricated by using a syringe filled with. The end of the needle is submerged into a neutral bath maintained at 37°C. The fibers are harvested using a

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designated tray and then washed and dried. Data to date have shown similar tensile strengths between rat tail tendon fibers and reconstituted (dry) extruded cross-­linked type I collagen fibers [110]. This result has further been substantiated by subsequent in vitro and in vivo studies (Table 13.1). In vitro, many parameters have been investigated to mimic native tendon structure, such as collagen source (bovine Achilles tendon, rat tail tendon) and concentration, the amount of PEG used in the extrusion buffer, the internal diameter of the extrusion tube [8], the extraction method (acid or pepsin solubilization) [111–113], the bath used for extrusion [114], the washing baths after extrusion of self-­assembled collagen fibers [115], and the comparison between soluble and insoluble collagen [116]. Cross-­linking is for obtaining the desired mechanical characteristics and degradation rate of extruded fibers. Without cross-­linking, fibers would degrade very quickly and would therefore not be suitable for any in vivo applications [8,117–121]. Besides, cross-­linking affects cell–fiber interactions, which determine the capacity of the scaffold to become progressively replaced by the native tissue after implantation [8]. However, especially potent cross-­linking agents have cytotoxic effects and may cause adverse tissue reactions after transplantation. Thus, different cross-­linkers and cross-­ linking approaches have been investigated over the years [122]. In an in vivo ovine patellar tendon injury model, implants of extruded bovine collagen fibers cross-­linked with either 1-­ ethyl-­ 3-­ (3-­ dimethylaminopropyl)carbodiimide (EDC) or with EDC associated with EGDE (ethyleneglycoldiglycidylether) were compared. EDC-­cross-­ linked fibers showed better mechanical properties as well as a better tissue integration after 6 months compared with EDC/EGDE-­cross-­linked fibers. However, because of a high degree of cross-­linking, both cross-­linked implants exhibited a low resorption rate after 6 months [29]. EDC also improved fibroblast migration compared with cross-­linking methods like dehydrothermal (DHT) cross-­linking and ultraviolet (UV) light [112,123]. Further studies used polyethylene glycol (PEG) [117] or an optimized DHT cross-­linking protocol [120] and also investigated the potential of the natural cross-­linker Myrica rubra [119]. Glutaraldehyde (GTA) has been disregarded as a potential cross-­linker because of its slow degradation rate, low cytocompatibility, and observed local increase in inflammatory cells after implantation in vivo [124,125]. Different strategies have been explored to improve the mechanical properties of extruded fibers. A study investigated subfibrillar orientation and DCN incorporation with regard to mechanical properties and found that DCN increased the ultimate tensile strength of non-­cross-­linked fibers [26]. Furthermore, extruded collagen fibers were submerged in a collagen chondroitin-­6-­sulfate solution and subsequently lyophilized. The porous structure obtained was favorable for cell migration, and fibers were dense and resistant to approximately 60 N [28]. By casting extruded collagen fibers in type I rat tail collagen gels and seeding with fibroblasts, a higher modulus and peak stress was reported after 25 days [27]. Extruded collagen fibers cross-­ linked with EDC lead to structural and mechanical properties (i.e., ultimate tensile strength and modulus) and degradation rate similar to autogenous tendon grafts in a rabbit Achilles tendon injury model. The animal model was based on the creation of a 3 cm gap in the gastrocnemius tendon. The collagen fibers were subsequently sutured to the gastrocnemius muscle and the calcaneus. The fibers were

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completely resorbed and replaced by neotendon tissue after 52 weeks [126]. These results have been supported by further studies using rabbit models [124]. A study performing anterior cruciate ligament (ACL) replacements in dogs showed complete remodeling of the fibers 12 weeks after implantation. Furthermore, extruded fibers have been used for ACL reconstitution in a goat model. Highly purified and cross-­linked fibers resulted in poor tissue ingrowth and resorption owing to their inertness. In summary, these studies indicate that less purified and less cross-­linked fibers result in faster connective tissue ingrowth and resorption after implantation. The authors argued that less purified collagen would lead to a greater tissue reaction after implantation, and less cross-­linked collagen would resorb faster and release earlier molecular collagen with chemotactic activity [127]. Finally, extruded collagen fibers have been used in ovine patellar tendon regeneration. The authors observed tissue ingrowth and integration, as well as excellent mechanical properties (i.e., modulus, strain to failure) [29]. In summary, the lack of clinical trials and commercially available products indicate the need for further investigation of extruded collagen fibers for biomedical applications. Scalability of the approach may be the limiting factor in the clinical translation and commercialization of this technology.

13.2.5  Lyophilized materials Approaches utilizing freeze-­drying of collagen or gelatin suspensions enable the formation of three-­dimensional scaffolds, termed sponges. Because of a low antigenicity, good mechanical properties, and the ability to promote cellular attachment and proliferation, collagen sponges are subject of investigation as biodegradable material in tendon tissue engineering (Table 13.2). Sponges can be cross-­linked before or after the freeze-­drying process. The sample is poured into a suitable vessel, which conducts the temperature uniformly. After freezing, the pressure is reduced, and the temperature is increased to allow the sublimation of the water content in a freeze dryer (i.e., lyophilization). Sponges can be generated having either a macro-­or a microporous structure with the obtained pore structure mirroring the ice crystal morphology after freezing. The potential of type I collagen sponges for tendon tissue engineering has been widely investigated in vitro. The comparison between type I collagen sponges, hydrogels, and films has been explored intensively. Superior mechanical strength accompanied by facilitated cell attachment and proliferation in sponges indicates their suitability for tendon tissue engineering applications [128] (Fig. 13.3). Furthermore, type I collagen sponges induced no inflammatory host response after implantation into an Achilles tendon defect in rats; moreover, accelerated healing and mechanical properties comparable with native tendons were observed [97]. To improve efficacy of collagen sponges, mechanical stimulation can be applied. Sponges seeded with rabbit BMSCs showed improved mechanical properties such as increased stiffness, modulus and maximum stress and force. Furthermore, enhanced cell alignment and expression of tendon-­and ECM-­related markers were observed [96,129,130]. However, the benefit of mechanical stimulation has only been shown for type I collagen sponges and not for type I collagen hydrogels [131]. Another way to improve the mechanical properties of type I collagen sponges is by simply varying the length of the scaffold [132] and by incorporating molecules such as

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Figure 13.3  SEM images of a non-­cross-­linked collagen type I sponge made of bovine Achilles tendon (BAT) at different magnifications (a); collagen type I staining (b); collagen type III staining (c); collagen type V staining (d); tenomodulin staining on a BAT type I collagen sponges seeded with tenocytes after 14 days of culture (e). DAPI is blue and the various markers are green. Scale bar: 200 μm. Density: 30,000 cells/cm2.

chondroitin-­6-­sulfate [133] or PLA [134]. The benefits of mechanical stimulation have also been confirmed in vivo [95,135,136]. The addition of growth factors such as GDF 5 and 6 has shown to improve tendon healing in a rat Achilles tendon injury model [98]. Furthermore, collagen has been combined with silk to form sponges without the addition of molecules or cells. 24 weeks after implantation in tendon defects in rabbits, tendon diameter, histological appearance, and tensile strength were similar to native tendon tissue [137]. Another study incorporated human embryonic stem cells derived from mesenchymal stem cells (hESCs–MSCs) within knitted silk/collagen sponges and reported enhanced tenogenic differentiation, shown by upregulated gene expression of tendon-­related markers, such as type I collagen, Epha4 (Ephrin type-­A receptor A4) and scleraxis [99]. When tendon stem/progenitor cells (TSPCs) were seeded on knitted silk–collagen scaffolds and implanted in a rabbit rotator cuff injury model, 12 weeks after implantation, the TSPCs-­treated group exhibited improved tendon regeneration by enhanced biomechanical properties, increased collagen deposition, and differentiation of TSPCs into tendonlike cells [100]. Bioactive silk/collagen sponges combined with SDF-­1α have been reported to enhance tendon regeneration in an Achilles tendon injury model in rats, by triggering fibroblast-­like cell migration and enhanced tendon ECM production (e.g., type I collagen, type III collagen, decorin) [101]. Because the outcomes of these studies were promising, further clinical trials need to be implemented to assess the efficiency of collagen/silk sponges for tendon tissue engineering. Collagen sponges are already used for many therapeutic applications, such as hemostatic sponges for bleeding surfaces, and are currently subject of research for tendon tissue engineering. In a clinical trial, an absorbable gelatin sponge loaded with recombinant

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human bone morphogenetic protein-­12 (rhBMP-­12) (Gelfoam, Pfizer) was investigated. Previously, rhBMP-­12 has been shown to improve ligament and tendon tissue formation and to improve tendon healing in rats [138]. However, heterotopic ossification has been observed in 10 of 16 patients. Whereas no adverse events were reported for the majority of patients, immunogenic responses have been reported in five patients. These immunogenic reactions may be explained by the presence of telopeptides. Finally, the authors concluded that complete healing of the rotator cuff occurred in 14 of 16 patients [102,139]. Furthermore, Gelfoam loaded with rhBMP-­12 was shown to reduce adhesion formation during tendon healing [140]. An in vivo study in rats tested Spongostan (Ferrosan medical devices) gelatin sponges combined with platelet-­rich plasma (PRP). Rotator cuff healing was successful with PRP, but no beneficial effect of the combination of the sponge and PRP was obvious compared with local PRP injections into the site of injury at the tendon-­to-­bone interface [141]. Even though the following products are for different applications, they have potential for tendon tissue engineering applications: collagen sponges manufactured by Integra Life , DuraMatrix-­Onlay Plus (Collagen Matrix Inc), Davol Inc. (Avitene), DePuySynthes (Biocol), SpongeCol (Advanced BioMatrix), Surgispon (Aegis Lifesciences), and Surgifoam (Ethicon, Johnson-­Johnson©).

13.3  Tissue graft–based therapies Tissue grafts are still considered the gold standard in orthopedic tissue engineering. On the one hand, they show high level of structural, compositional, and mechanical similarity to the tissue to be replaced. On the other hand, they have a higher risk for stimulating immune reactions. They are classified as autografts, being harvested and transplanted in the same individual; allografts, being transplanted between individuals of the same species; and xenografts, where the donor belongs to a different species than the recipient. Although allografts and autografts present a better compatibility in humans, they pose a high risk of disease transmission and exhibit lower availability than xenografts. Besides the species of origin, the selection of donor (e.g., age, sex) and tissue section impacts on both the properties of the tissues [142–147] and the derived grafts. The use of tissue grafts for tendon regeneration strategies is a field that is currently gaining increasing interest. Tissue grafts have already been widely used to treat tendon and ligament injuries in clinics. However, autografts have shown to cause donor site morbidity, whereas allografts and xenografts have been reported to trigger immune reactions in the host. Because of a limited risk of host rejection, the use of decellularized allografts and xenografts has arisen as an alternative and promising approach for tendon regeneration strategies [148].

13.3.1  Tissue graft fabrication The processing of tissue grafts determines their final characteristics and, hence, is crucial for their success after implantation (Table 13.3). The main steps used in tissue graft fabrication are decellularization, delipidation, cross-­linking, and sterilization. It is worth noting that delipidation is an optional step and not included in many protocols, as decellularizing

Source Species

Tissue

Processing

Porcine (xenograft)

Flexor tendon

Freeze–thaw cycles, hypotonic solution, SDS, nucleases, hypertonic solution Peracetic acid sterilization

Equine (xenograft)

Flexor digitorum superficialis tendon

Canine (xenograft)

Infraspinatus tendon slices

Equine (xenograft)

Flexor digitorum superficialis tendon

Outcomes

Effective decellularization maintained histological similarity with native tendon. Reduction of GAGs, DNA, and Gal-­epitope. No direct or indirect cytotoxicity with 3T3 cells. Low encapsulation and high host cell penetration in subcutaneous rat model. Preserved mechanical properties. Freeze–thaw cycles, SDS/ Effective decellularization with 2% SDS, reduced proTriton/TnBP, trypsin-­EDTA, tein content, preserved GAGs. Tissue structure and DNAase-­I and ethanol mechanical properties maintained. Decellularized tendons cytocompatible with equine MSCs. Freeze–thaw cycles and RNAase No inflammation after implantation in rabbit patellar treatment tendon defects. Loss of cell alignment and reduced Freezing and sectioning cell number at day 14. Increased gene expression Seeding with rabbit BMSCs of tenomodulin, type III collagen, MMP3, and MMP13 compared with implanted slices without BMSCs. Reduced expression of type I collagen. Freeze–thaw cycles, SDS, trypsin-­ MSC integrated in scaffolds and produced GAGs. EDTA, DNAase-­I, and ethanol Mechanical strain promoted tenogenic marker Seeding with equine BMSCs expression of MSCs on the grafts. Mechanical Mechanical stimulation in strain of 3% resulted in mechanical properties simibioreactor lar to native tendon.

References [153]

Scaffolds for tendon tissue engineering

Table 13.3  Tissue grafts for tendon regeneration strategies.

[154]

[155]

[156]

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Continued

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Table 13.3  Tissue grafts for tendon regeneration strategies.—cont’d Source Species

Tissue

Processing

Outcomes

References

Ovine (xenograft)

Forestomach (Endoform™)

Osmotic gradient and detergents Ethylene oxide sterilization

[157]

Canine (xenograft)

Flexor tendon

Human (allograft)

Patellar tendon

No effects on mechanical properties observed.

[159]

Human (allograft)

Dermis (GraftJacket™)

Freeze–thaw cycles and RNAase treatment Freezing, sectioning Multilayer scaffold fabrication and seeding with rat BMSCs Freeze–thaw cycles, sonication, hypotonic buffer, SDS, Benzonase, and hypertonic buffer GraftJacket™ cryogenic patented process

Supported tenocyte growth and no inflammatory response in vitro. Improved integration and histological remodeling after implantation in rat rotator cuff repair model. No improvement of mechanical properties in vivo. Improved mechanical properties of augmented tendons seeded with BMSCs in rat rotator cuff defect. More prominent, organized, and robust tissue produced in BMSC–scaffold augmented tendons.

Augmented rotator cuff repair in rats after 12 months with histological structure similar to native tendon. Poor cellularization of the graft. Poor recovery of mechanical properties.

[160]

[158]

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agents possess a certain fat removal capacity. Each of these steps can be carried out utilizing a wide range of different techniques, conditions, and agents, which differently affect ECM composition and integrity and contribute to a tissue graft with unique properties. The ideal decellularization process of a tissue has three main objectives: (1) to remove unwanted cellular tissue components efficiently; (2) to cause minimum damage to the ECM architecture and properties; and (3) to reduce the number of antigens, which may elicit a host immune response. Despite significant strides, decellularized grafts have been shown to trigger immune reactions after transplantation [149]. Remaining cellular debris is responsible for the inflammatory response of M1 macrophages [150,151], whereas effectively decellularized grafts can promote the remodeling reaction of M2 macrophages [150,152]. The techniques used to deploy decellularizing agents depend on the tissue characteristics and play a critical role in decellularization efficiency and resulting damage on the ECM. Among them, immersion and agitation are the most frequently used techniques [161]. Incubation time may vary depending on tissue size, where thicker tissues requiring extended incubation times compared with thinner tissues [162]. Perfusion is a delivery method that utilizes the tissue vasculature to deliver decellularization agents by pumping the solutions at constant or gradient pressure. It is mainly used for decellularization of either tissues with a dense vasculature or whole organs [163,164]. With regard to their nature, decellularizing agents can be classified as physical, chemical, and enzymatic [165]. Techniques using physical agents include freeze–thaw cycles, sonication, liming, hot water sanitation, or supercritical fluids. Generally, these methods are effective in disrupting the tissue immanent cells. Because these techniques are not capable of fully eliminating cellular debris, they are often used in combination with chemical or enzymatic treatments [161,166,167]. Frequently, such treatments compromise the integrity of the ECM [168,169]. Chemical agents used in decellularization strategies are classified as alkaline or acid agents, detergents, osmotic solutions, solvents, and chelators. Alkaline or acid agents (e.g., sodium hydroxide, peracetic acid, PAA) are widely used to destroy and solubilize cellular components. In addition, agents such as PAA exhibit antimicrobial properties that reduce the bioburden during processing. Their efficacy and effects on the ECM depend on the combination with other agents [170,171], as well as on other parameters, such as pH [172]. Detergents solubilize cell membranes and break interactions of proteins with other molecules, making them effective and cost-­efficient decellularizing agents. However, detergents have been reported to compromise the ECM. The nonionic detergent Triton X-­100 and the ionic detergent sodium dodecyl sulfate (SDS) are the most commonly used detergents for tissue decellularization. Nonionic detergents are less effective than ionic detergents; however, nonionic detergents cause a lower damage on tissue structure than ionic [173], whereas zwitterionic detergents such as 3-­((3-­cholamidopropyl) dimethylammonio)-­ 1-­ propanesulfonate (CHAPS) combine properties of both [174]. Osmotic solutions used as decellularizing agents, such as NaCl or Tris buffers, create an osmotic potential that lyses the cells. Because they are less effective than other chemical and enzymatic agents [175], they are generally used in combination with other agents. Solvents (e.g., isopropanol [176], ethanol [177], tributyl phosphate [TBP] [171]) and chelators (e.g., ethylenediaminetetraacetic acid [EDTA] [178,179]) are frequently used to complement the action of different decellularizing agents.

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Even though enzymatic agents have a high specificity, enzymatic decellularization of tissues remains challenging. The protease trypsin is the most commonly used enzyme for decellularization purposes. It is used in combination with chemical agents, such as detergents or chelators. However, prolonged incubation times have been reported to adversely affect tissue integrity [179]. Other enzymatic agents include dispases, which are used to remove epithelium layers [180]; nucleases, which are known to disrupt nucleic acids; or α-­galactosidases, which are used to reduce the presence of Gal-­epitopes and therefore reduce the risk of immune rejection of the tissue graft by the host [181]. In tissue grafts, the native cross-­linking of the tissue components is preserved. However, the decellularization processes often affect tissue integrity, potentially compromising its stability after implantation. Thus, exogenous cross-­ linking can be used as an optional step to enhance the durability of the graft in vivo. Nevertheless, the interactions produced may alter the properties of the graft, because many cross-­linking agents are reported to be cytotoxic and to trigger immune effects in vitro and in vivo [122,182]. GTA has been widely used because of its high cross-­linking efficiency, leading to enhanced mechanical properties and durability of tissue grafts in vivo. On the contrary, GTA has been reported to cause undesired effects, such as calcification [183] and inflammatory responses [184]. Another chemical cross-­linker is EDC, which has shown similar efficacy to GTA, while eliciting a lower cytotoxicity [185,186]. Other cross-­linking agents include hexamethylene diisocyanate (HMDI) [187] or ethylene glycol diglycidyl ether (EGDE) [188]. To overcome the drawbacks of cytotoxicity, notably observed in strongly cross-­linked grafts, natural cross-­linkers like genipin have been investigated in vitro [189] and in vivo [190], showing similar cross-­linking properties compared with GTA [189,190]. The risk of disease transmission is a hindrance in the processing of tissue grafts. Generally, allografts are not sterilized or processed in aseptic conditions, whereas xenografts undergo a final sterilization step to ensure safety for transplantation. The ideal sterilization technique must be safe, easy, and effective and must have a minimum impact on the ECM of the tissue [191]. Physical techniques like electron beam and gamma irradiation are widely used and are effective in destroying nucleic acids of pathogens. However, at effective doses, they may damage ECM proteins, such as collagen [192], and therefore compromise the mechanical properties of the tissue graft [193]. Recently, supercritical carbon dioxide gathered attention as an alternative physical sterilization method in combination with other agents [194,195]. Additionally, sterilization using chemical agents is an extended practice in tissue graft engineering. Ethylene oxide (ETO) has proven to be effective sterilizing agent by preserving the mechanical properties of the tissue [196]. However, the equipment needed is complex and cost-­intensive. A decellularizing chemical agent causing only minimal effects on the ECM is PAA, which is used because of its antiviral properties [197]. Other chemical sterilization strategies include gas plasma [198], hydrogen peroxide [199], and ethanol [200]. Overall, there are many variables during the tissue processing, which ultimately define the final properties of the tissue graft, which, in return, determine its suitability for specific clinical applications.

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13.3.2  Tissue grafts from decellularized tendons Tissue graft decellularization for tendon tissue engineering is a vivid area of research, particularly for decellularization of xenografts. Until now, only a few investigations have assessed the decellularization of tendon allografts. Numerous studies focus on in vitro assessment of decellularization, investigating the efficacy of different techniques, as well as their effects on tissue integrity. Histological and topographical assessment and/or mechanical testing, together with other biochemical assays, are used to detect residual DNA, detergents, or signs of ECM degradation [148]. Recently, several protocols for tendon graft decellularization have been developed, which result in native tissue preservation, including treatments with detergents such as TBP + SDS [201], hypotonic solution + SDS + PAA [202], TBP + PAA [203], or freeze-­thawing + Triton X-­100 [204]. Although these procedures are optimized to minimize adverse effects on the tendon structure, changes, such as partial loss of structure and disorganized collagen fibers, are frequently observed. These changes also affect the biomechanical behavior of the tissue graft. However, decellularized tendon was reported to maintain its native mechanical properties after optimized processing [205]. Decellularization of tendon tissue usually results in cytocompatible scaffolds, as proven by numerous studies [148]. Tendon tissue grafts have been successfully reseeded with fibroblasts [206], tenocytes [207], tendon-­derived stem cells [208], ADSCs [209], and BMSCs [210]. However, homogenous repopulation and graft penetration of decellularized tendon scaffolds still remains challenging. Because of a very compact and dense tissue structure, cell attachment and proliferation is often restricted to matrix surfaces [211]. To overcome these drawbacks, different strategies are currently under investigation. In particular, recellularization protocols [205], applying cyclic loading via a bioreactor, pose promising strategies to facilitate cell infiltration and to enhance subsequent tenogenic differentiation [209,210,212]. Several in vivo studies have investigated decellularized tendon tissue to prove its suitability as a tendon graft. After xenograft transplantation in rats, no acute immune reaction was observed, which was attributed to the decellularization process [213]. Similar results were observed using subcutaneous implantation models [214]. After implantation of decellularized tendon slices from dogs into a rabbit rotator cuff repair model, complete tissue resorption and improved mechanical properties were observed after 12 weeks, compared with the control defects [215]. Similarly, positive integration of decellularized autografts was observed in a bovine superficial digital tendon injury model, with an improved overall outcome compared with GTA-­preserved tendon autografts [216]. Despite the good tissue integration, grafts still fail to fully restore the native properties of the tendon. Thus, an alternative approach aimed to implant decellularized tendons, reseeded with autologous cells, to improve tissue remodeling and to reduce immunogenicity. Studies have been conducted transplanting human flexor tendons populated with human ADSCs in a subcutaneous rat model [217]. In a long-­ term study, tenocyte-­seeded Achilles tendon allografts were used for partial tendon replacement in a rat model, and no adverse immune reactions were observed after 6 months [218]. Another study successfully used xenogeneic tendon slices seeded with BMSCs for patellar tendon defect in rabbits [155]. Although new approaches seem to

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improve tissue remodeling and inflammatory reactions, the resulting biomechanical properties still do not match those observed in native tendon [148] or tendon–bone interfaces [219]. In the clinic, the use of tendon autografts and allografts without further processing is currently considered the gold standard to treat tendon injuries. However, their side effects and immune reactions still pose a challenge that needs to be overcome for successful tissue replacement. In summary, the use of decellularized tissue grafts poses a very promising approach in tendon repair.

13.3.3  Tissue grafts from other tissues Besides tendon grafts, a wide range of allograft and xenograft products from different sources is already in clinical use. Augmentation systems are used to reinforce tendons during the healing process and provide mechanical support and a suitable milieu for cell repopulation [220,221]. Decellularized tissue grafts from other sources than tendon have already shown acceptable cytocompatibility in vitro (Fig. 13.4), where higher cytocompatibility was observed in nonchemically cross-­linked materials [222]. In vivo models have been used to assess the immune response to these materials, with regard to effects caused through tissue processing [223] and Gal-­epitope [224]. Although various systems (e.g. biologically active scaffolds) have shown promise in vitro, none of them was able to induce regeneration and the repaired tissue had inferior mechanical properties to the tissue prior to injury (in the case of enthesis, for example) [225]. Commercially available tissue graft products used as augmentation systems include human dermis (GraftJacket, Wright Medical), porcine dermis (Permacol, Covidien; Conexa, Tornier), porcine small intestinal submucosa (Restore, DePuy Orthopedics; Cuffpatch, Organogenesis), equine pericardium (OrthADAPT, Pegasus Biologic Inc.), and bovine fetal dermis (TissueMend, Stryker Orthopedics). Generally, tissue grafts with higher mechanical properties (i.e., cross-­linked dermis–derived materials) pose valuable augmentation systems for irreparable tendon tears, as frequently observed in rotator cuff injuries. However, they have been shown to trigger adverse effects in moderate injuries, as observed with Restore, which is no longer used in clinics [226].

Figure 13.4  SEM of decellularized porcine peritoneum (a). Immunocytochemistry of human primary tenocytes seeded on decellularized porcine peritoneum for tenomodulin (b) and tenascin-­C (c). DAPI is blue and the various markers are green. Scale bars 100 μm.

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13.4  Scaffold-­free tissue engineering by self-­assembly To engineer a complex tissue such as tendon, cells and ECM need to be organized three-­ dimensionally, as in the native tissue. The development of stratified or multiphasic tissue substitutes allows for combining cells and/or growth factors with scaffolds to support the tissue architecture and organization and to mimic the mechanical properties of the native tissue as close as possible [227,228]. Ultimately, biological integration of the scaffold at the implantation site must be considered to augment tissue regeneration. However, most cell delivery techniques offer little control over location and differentiation of implanted cells. Furthermore, scaffold materials often fail to fully recapitulate structure and biomechanical properties of the native tissue, leading to impaired tissue remodeling [9,229]. Therefore, functional regeneration by using the innate capacity of cells to create their own tissue-­specific ECM is highly desirable [230,231]. In contrast to limitations of tissue reconstruction using scaffolds or cell injections, an alternative approach for tissue engineering enables the production of scaffold-­free cell constructs using surfaces coated with temperature-­responsive polymers. This technique allows for a noninvasive harvest of contiguous cell sheets without the usage of proteolytic enzymes, which are stabilized by preserved cell–cell contacts and deposited ECM. Thus, viable cell sheets can be transplanted, bypassing the need for biodegradable scaffolds and their shortfalls [232–235]. Cell sheets adhere to each other rapidly, making it possible to generate more complex multilayer tissue sheets that are able to communicate and become synchronized as functional tissues [236]. Because of these advantages, cell sheet technology has been extensively studied for various clinical targets and has already been successfully applied in clinical trials, targeting the regeneration of cornea, esophageal epithelia, heart, periodontal ligament, bladder epithelia, liver, and cartilage [237–242]. The suitability of bioengineered tendon-­derived stem cell (TDSC) sheets has been investigated for enhancing ACL (anterior cruciate ligament) recovery and Achilles tendon healing as well as for recreation of the complex tendon-­to-­bone interface, the enthesis [243–245]. These studies demonstrated the feasibility of implanting TDSC sheets in tendons in vivo, representing a novel and promising therapeutic strategy for tendon tissue engineering. However, manufacturing times of several months pose a considerable challenge for future clinical applications. To reach functional regeneration of tendon tissue, replacements need to incorporate as many of the native tissue properties as possible. Another technology allows for the generation of tendonlike constructs in vitro, without the use of artificial scaffolding. Cells isolated from tendons were grown to confluence in culture and allowed to self-­ assemble into a cylinder between two anchor points. The resulting scaffold-­free tissue exhibited many characteristics of embryonic tendon. It was composed of aligned collagen fibrils, a large number of cells, and an excess of noncollagenous extracellular matrix [246]. Mature human tendon fibroblasts have been shown to hold an intrinsic capability to perform embryonic-­like collagen fibrillogenesis when cultured under tension using a cell-­embedded matrix model [247,248]. These technologies, however, require prolonged culture times, which often results in phenotypic drift and cellular senescence [249]. To overcome these drawbacks, attempts have been made combining cell sheets with biodegradable meshes or tissue

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grafts [250,251]. The resulting implantable devices exhibited high mechanical stability and promising results after implantation in in vivo studies. Macromolecular crowding (MMC) is an approach where the naturally dense and crowded in vivo environment of stem cells is imitated by the addition of inert macromolecules to the culture media [252–255]. MMC has been shown to dramatically increase tissue-­specific ECM deposition and to maintain permanently differentiated cell phenotype [256–260]. Using MMC, ECM-­rich tendon cell sheets with intact cell–cell junctions were produced [230]. MMC poses a promising technique to be combined with scaffold-­free approaches to enhance ECM deposition, tissue integrity, and mechanical stability for successful regeneration after transplantation. Despite all the advancements that have become available in recent years, a functional three-­dimensional tendon equivalent has yet to be developed.

13.5  Conclusion and future perspectives Despite extensive ongoing research and recent advancements in tissue engineering, not many devices have been successfully translated into the clinic. The lack of effective devices used for clinical applications is primarily due to the difficulty in mimicking the exact architecture and composition of the native tissue and simultaneously providing a suitable microenvironment for cell attachment, migration, and proliferation. Furthermore, the complex biological environment and the spatiotemporal signaling events driving tendon repair and regeneration are still poorly understood. The lack of successful clinical outcomes underlines the importance of engineering reproducible, scalable, nontoxic, and nonimmunogenic, with adequate mechanical integrity and ability to deliver spatiotemporal cues implantable devices for functional tendon regeneration.

List of abbreviations ACL  Anterior cruciate ligament ADSC  Adipose-­derived stem cells BMP  Bone morphogenetic protein BMSC  Bone marrow stem cells CE Mark  (Conformité Européenne)—certification mark indicating conformity with health, safety, and environmental protection standards for products sold within European Economic Area (EEA) CHAPS  3-­((3-­Cholamidopropyl)dimethylammonio)-­1-­propanesulfonate DCN  Decorin DHT  Dehydrothermal treatment ECM  Extracellular matrix EDC  1-­Ethyl-­3-­(3-­dimethylaminopropyl)carbodiimide EDTA  Ethylenediaminetetraacetic acid ES  Electrospinning ETO  Ethylene oxide FGF2  Fibroblast growth factor-­2 GDF  Growth/differentiation factor

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GTA  Glutaraldehyde HDMI  Hexamethylene diisocyanate hESC–MSCs  Human embryonic stem cell–derived mesenchymal stem cells IL  Interleukin MDC  Muscle-­derived cells mGLT  Methacrylated gelatin PAA  Peracetic acid PCL  Poly-­Ɛ-­caprolactone PDGF  Platelet-­derived growth factor PEG  Polyethylene glycol PGA  Poly(glycolic acid) PLA  Poly(lactic acid) PLLA  Poly(l-­lactic acid) PLGA  Poly(lactic-­co-­glycolic acid) PRP  Platelet-­rich plasma RGD  Arginine–glycine–aspartic acid tripeptide rhBMP-­12  Recombinant human bone morphogenetic protein-­12 SDF-­1 α  Stromal-­derived factor 1 α SDS  Sodium dodecyl sulfate TBP  Tributyl phosphate TGF  Transforming growth factor VEGF  Vascular endothelial growth factor UV  Ultraviolet

Acknowledgments This work has been supported from the Science Foundation Ireland, Career Development Award Programme (grant agreement number: 15/CDA/3629); Science Foundation Ireland and the European Regional Development Fund (grant agreement number: 13/RC/2073); EU H2020, ITN award, Tendon Therapy Train Project (grant agreement number: 676338). HC-­M, SG, SK, and DIZ have no competing interests. SR and YB are employees of Sofradim Production—a Medtronic company.

References [1] P.B. Voleti, M.R. Buckley, L.J. Soslowsky, Tendon healing: repair and regeneration, Annu Rev Biomed Eng 14 (2012) 47–71. [2] H. Tempfer, C. Lehner, M. Grütz, R. Gehwolf, A. Traweger, Biological augmentation for tendon repair: lessons to be learned from development, disease, and tendon stem cell research, in: J.M. Gimble, D. Marolt, R. Oreffo, H. Redl, S. Wolbank (Eds.), Cell Engineering and Regeneration, Springer International Publishing, Cham, 2017, pp. 1–31. [3] G. Walden, X. Liao, S. Donell, M.J. Raxworthy, G.P. Riley, A. Saeed, A clinical, biological, and biomaterials perspective into tendon injuries and regeneration, Tissue Eng B 23 (1) (2016) 44–58. [4] H. Tempfer, A. Traweger, Tendon vasculature in health and disease, Front Physiol 6 (2015) 330, https://doi.org/10.3389/fphys.2015.00330. 2015.

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[208] L.-­J. Ning, Y.-­J. Zhang, Y. Zhang, Q. Qing, Y.-­L. Jiang, J.-­L. Yang, J.-­C. Luo, T.-­W. Qin, The utilization of decellularized tendon slices to provide an inductive microenvironment for the proliferation and tenogenic differentiation of stem cells, Biomaterials 52 (2015) 539–550. [209] D.W. Youngstrom, J.E. LaDow, J.G. Barrett, Tenogenesis of bone marrow-­, adipose-­, and tendon-­derived stem cells in a dynamic bioreactor, Connect Tissue Res 57 (6) (2016) 454–465. [210] J.H. Wu, A.R. Thoreson, A. Gingery, K.N. An, S.L. Moran, P.C. Amadio, C. Zhao, The revitalisation of flexor tendon allografts with bone marrow stromal cells and mechanical stimulation: an ex vivo model revitalising flexor tendon allografts, Bone Joint Res 6 (3) (2017) 179–185. [211] G. Schulze-­Tanzil, O. Al-­Sadi, W. Ertel, A. Lohan, Decellularized tendon extracellular matrix—a valuable approach for tendon reconstruction? Cells 1 (4) (2012) 1010–1028. [212] D.W. Youngstrom, J.G. Barrett, Tendon differentiation on decellularized extracellular matrix under cyclic loading, Methods Mol Biol 1502 (2016) 195–202. [213] S.S. Raghavan, C.Y. Woon, A. Kraus, K. Megerle, M.S. Choi, B.C. Pridgen, H. Pham, J. Chang, Human flexor tendon tissue engineering: decellularization of human flexor tendons reduces immunogenicity in vivo, Tissue Eng 18 (7–8) (2012) 796–805. [214] K.A. Alberti, Q. Xu, Biocompatibility and degradation of tendon-­derived scaffolds, Regen Biomater 3 (1) (2016) 1–11. [215] J. Pan, G.-­M. Liu, L.-­J. Ning, Y. Zhang, J.-­C. Luo, F.-­G. Huang, T.-­W. Qin, Rotator cuff repair using a decellularized tendon slices graft: an in vivo study in a rabbit model, Knee Surg Sports Traumatol Arthrosc 23 (5) (2015) 1524–1535. [216] R. Ramesh, N. Kumar, A.K. Sharma, S.K. Maiti, S. Kumar, K. Charan, Acellular and glutaraldehyde‐preserved tendon allografts for reconstruction of superficial digital flexor tendon in bovines: Part II – gross, microscopic and scanning electron microscopic observations, J Vet Med Ser A 50 (10) (2003) 520–526. [217] T. Schmitt, P.M. Fox, C.Y. Woon, S.J. Farnebo, J.A. Bronstein, A. Behn, H. Pham, J. Chang, Human flexor tendon tissue engineering: in vivo effects of stem cell reseeding, Plast Reconstr Surg 132 (4) (2013) 567e–576e. [218] C. Güngörmüş, D. Kolankaya, E. Aydin, Histopathological and biomechanical evaluation of tenocyte seeded allografts on rat Achilles tendon regeneration, Biomaterials 51 (2015) 108–118. [219] J.M. Chen, C. Willers, J. Xu, A. Wang, M.H. Zheng, Autologous tenocyte therapy using porcine-­derived bioscaffolds for massive rotator cuff defect in rabbits, Tissue Eng 13 (7) (2007) 1479–1491. [220] U.G. Longo, A. Lamberti, N. Maffulli, V. Denaro, Tendon augmentation grafts: a systematic review, Br Med Bull 94 (2010) 165–188. [221] S. Chaudhury, C. Holland, M.S. Thompson, F. Vollrath, A.J. Carr, Tensile and shear mechanical properties of rotator cuff repair patches, J Shoulder Elbow Surg 21 (9) (2012) 1168–1176. [222] K.P. Shea, M.B. McCarthy, F. Ledgard, C. Arciero, D. Chowaniec, A.D. Mazzocca, Human tendon cell response to 7 commercially available extracellular matrix materials: an in vitro study, Arthrosc J Arthrosc Relat Surg 26 (9) (2010) 1181–1188. [223] J.E. Valentin, J.S. Badylak, G.P. McCabe, S.F. Badylak, Extracellular matrix bioscaffolds for orthopaedic applications: a comparative histologic study, J Bone Joint Surg 88 (12) (2006) 2673–2686. [224] H. Xu, H. Wan, W. Zuo, W. Sun, R.T. Owens, J.R. Harper, D.L. Ayares, D.J. McQuillan, A porcine-­derived acellular dermal scaffold that supports soft tissue regeneration: removal of terminal galactose-­alpha-­(1,3)-­galactose and retention of matrix structure, Tissue Eng 15 (7) (2009) 1807–1819.

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[225] T. Thangarajah, C.J. Pendegrass, S. Shahbazi, S. Lambert, S. Alexander, G.W. Blunn, Augmentation of rotator cuff repair with soft tissue scaffolds, Orthop J Sports Med 3 (6) (2015). [226] Y. Ono, D.A. Dávalos Herrera, J.M. Woodmass, R.S. Boorman, G.M. Thornton, I.K.Y. Lo, Can grafts provide superior tendon healing and clinical outcomes after rotator cuff repairs? A meta-­analysis, Orthop J Sports Med 4 (12) (2016) 2325967116674191. [227] G. Criscenti, A. Longoni, A. Di Luca, C. De Maria, C. van Blitterswijk, G. Vozzi, L. Moroni, Triphasic scaffolds for the regeneration of the bone–ligament interface, Biofabrication 8 (1) (2016) 015009. [228] S. Nossov, J.S. Dines, G.A. Murrell, S.A. Rodeo, A. Bedi, Biologic augmentation of tendon-­ to-­ bone healing: scaffolds, mechanical load, vitamin D, and diabetes, Instr Course Lect 63 (2014) 451–462. [229] X. Zhang, D. Bogdanowicz, C. Erisken, N.M. Lee, H.H. Lu, Biomimetic scaffold design for functional and integrative tendon repair, J Shoulder Elbow Surg 21 (2) (2012) 266–277. [230] A. Satyam, P. Kumar, X. Fan, A. Gorelov, Y. Rochev, L. Joshi, H. Peinado, D. Lyden, B. Thomas, B. Rodriguez, Macromolecular crowding meets tissue engineering by self‐assembly: a paradigm shift in regenerative medicine, Adv Mater 26 (19) (2014) 3024–3034. [231] J. Apostolakos, T.J. Durant, C.R. Dwyer, R.P. Russell, J.H. Weinreb, F. Alaee, K. Beitzel, M.B.R. McCarthy, M.P. Cote, A.D. Mazzocca, The enthesis: a review of the tendon-­to-­ bone insertion, Muscles, Ligaments Tendons J 4 (3) (2014) 333–342. [232] M. Yamato, T. Okano, Cell sheet engineering, Mater Today 7 (5) (2004) 42–47. [233] T. Iwata, M. Yamato, H. Tsuchioka, R. Takagi, S. Mukobata, K. Washio, T. Okano, I. Ishikawa, Periodontal regeneration with multi-­layered periodontal ligament-­derived cell sheets in a canine model, Biomaterials 30 (14) (2009) 2716–2723. [234] T. Iwata, K. Washio, T. Yoshida, I. Ishikawa, T. Ando, M. Yamato, T. Okano, Cell sheet engineering and its application for periodontal regeneration, J Tissue Eng Regenerat Med 9 (4) (2013) 343–356. [235] D. Thomas, D. Gaspar, A. Sorushanova, G. Milcovich, K. Spanoudes, A.M. Mullen, T. O’Brien, A. Pandit, D.I. Zeugolis, Scaffold and scaffold-­free self-­assembled systems in regenerative medicine, Biotechnol Bioeng 113 (6) (2016) 1155–1163. [236] H. Takahashi, T. Okano, Cell sheet‐based tissue engineering for organizing anisotropic tissue constructs produced using microfabricated thermoresponsive substrates, Adv Healthc Mater 4 (16) (2015) 2388–2407. [237] N. Matsuda, T. Shimizu, M. Yamato, T. Okano, Tissue engineering based on cell sheet technology, Adv Mater 19 (20) (2007) 3089–3099. [238] M. Sato, M. Yamato, K. Hamahashi, T. Okano, J. Mochida, Articular cartilage regeneration using cell sheet technology, Anat Rec 297 (1) (2014) 36–43. [239] K. Nishida, M. Yamato, Y. Hayashida, K. Watanabe, K. Yamamoto, E. Adachi, S. Nagai, A. Kikuchi, N. Maeda, H. Watanabe, Corneal reconstruction with tissue-­engineered cell sheets composed of autologous oral mucosal epithelium, N Engl J Med 351 (12) (2004) 1187–1196. [240] T. Ohki, M. Yamato, M. Ota, R. Takagi, M. Kondo, N. Kanai, T. Okano, M. Yamamoto, Application of regenerative medical technology using tissue‐engineered cell sheets for endoscopic submucosal dissection of esophageal neoplasms, Dig Endosc 27 (2) (2015) 182–188. [241] I. Ishikawa, T. Iwata, K. Washio, T. Okano, T. Nagasawa, K. Iwasaki, T. Ando, Cell sheet engineering and other novel cell‐based approaches to periodontal regeneration, Periodontol 2000 51 (1) (2009) 220–238. [242] K. Matsuura, Y. Haraguchi, T. Shimizu, T. Okano, Cell sheet transplantation for heart tissue repair, J Contr Release 169 (3) (2013) 336–340.

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[243] G. Chen, Y. Qi, L.I.E. Niu, T. Di, J. Zhong, T. Fang, W. Yan, Application of the cell sheet technique in tissue engineering, Biomed Rep 3 (6) (2015) 749–757. [244] P.P. Lui, O.T. Wong, Y.W. Lee, Application of tendon-­derived stem cell sheet for the promotion of graft healing in anterior cruciate ligament reconstruction, Am J Sports Med 42 (3) (2014) 681–689. [245] I. Komatsu, J.H. Wang, K. Iwasaki, T. Shimizu, T. Okano, The effect of tendon stem/ progenitor cell (TSC) sheet on the early tendon healing in a rat Achilles tendon injury model, Acta Biomater 42 (2016) 136–146. [246] S. Calve, R.G. Dennis, P.E.n. Kosnik, K. Baar, K. Grosh, E.M. Arruda, Engineering of functional tendon, Tissue Eng 10 (5–6) (2004) 755–761. [247] M.L. Bayer, C.-­Y.C. Yeung, K.E. Kadler, K. Qvortrup, K. Baar, R.B. Svensson, S.P. Magnusson, M. Krogsgaard, M. Koch, M. Kjaer, The initiation of embryonic-­like collagen fibrillogenesis by adult human tendon fibroblasts when cultured under tension, Biomaterials 31 (18) (2010) 4889–4897. [248] Z. Kapacee, S.H. Richardson, Y. Lu, T. Starborg, D.F. Holmes, K. Baar, K.E. Kadler, Tension is required for fibripositor formation, Matrix Biol 27 (4) (2008) 371–375. [249] D. Cigognini, A. Lomas, P. Kumar, A. Satyam, A. English, A. Azeem, A. Pandit, D. Zeugolis, Engineering in vitro microenvironments for cell based therapies and drug discovery, Drug Discov Today 18 (21) (2013) 1099–1108. [250] G. Mitani, M. Sato, M. Yamato, M. Kokubo, T. Takagaki, G. Ebihara, T. Okano, J. Mochida, Potential utility of cell sheets derived from the anterior cruciate ligament and synovium fabricated in temperature‐responsive culture dishes, J Biomed Mater Res 102 (9) (2014) 2927–2933. [251] Y. Inagaki, K. Uematsu, M. Akahane, Y. Morita, M. Ogawa, T. Ueha, T. Shimizu, T. Kura, K. Kawate, Y. Tanaka, Osteogenic matrix cell sheet transplantation enhances early tendon graft to bone tunnel healing in rabbits, BioMed Res Int 2013 (2013). [252] S.B. Zimmerman, B. Harrison, Macromolecular crowding increases binding of DNA polymerase to DNA: an adaptive effect, Proc Natl Acad Sci USA 84 (7) (1987) 1871–1875. [253] A.P. Minton, G.C. Colclasure, J.C. Parker, Model for the role of macromolecular crowding in regulation of cellular volume, Proc Natl Acad Sci USA 89 (21) (1992) 10504–10506. [254] R.J. Ellis, A.P. Minton, Cell biology: join the crowd, Nature 425 (6953) (2003) 27–28. [255] C. Chen, F. Loe, A. Blocki, Y. Peng, M. Raghunath, Applying macromolecular crowding to enhance extracellular matrix deposition and its remodeling in vitro for tissue engineering and cell-­based therapies, Adv Drug Deliv Rev 63 (4–5) (2011) 277–290. [256] D. Cigognini, D. Gaspar, P. Kumar, A. Satyam, S. Alagesan, C. Sanz-­Nogués, M. Griffin, T. O’Brien, A. Pandit, D.I. Zeugolis, Macromolecular crowding meets oxygen tension in human mesenchymal stem cell culture -­a step closer to physiologically relevant in vitro organogenesis, Sci Rep 6 (30764) (2016). [257] P. Kumar, A. Satyam, D. Cigognini, A. Pandit, D.I. Zeugolis, Low oxygen tension and macromolecular crowding accelerate extracellular matrix deposition in human corneal fibroblast culture, J Tissue Eng Regenerat Med 12 (1) (2018) 6–18. [258] P. Kumar, A. Satyam, D. Gaspar, D. Cigognini, C. Sanz-­Nogués, T. O’Brien, A. Pandit, D. Zeugolis, Macromolecular crowding: the next frontier in tissue engineering, in: Advances in Science and Technology, Trans Tech Publ, 2014. [259] A. Satyam, P. Kumar, D. Cigognini, A. Pandit, D.I. Zeugolis, Low, but not too low, oxygen tension and macromolecular crowding accelerate extracellular matrix deposition in human dermal fibroblast culture, Acta Biomater 44 (2016) 221–231. [260] K. Spanoudes, D. Gaspar, A. Pandit, D.I. Zeugolis, The biophysical, biochemical, and biological toolbox for tenogenic phenotype maintenance in vitro, Trends Biotechnol 32 (9) (2014) 474–482.

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Agbabiaka Oluwadamilola1, Safiyya Yousaf1, Mahbubeh Zare1, Masoud Mozafari2,3,5, Mansour Youseffi1, Peter Twigg1, Farshid Sefat1,4,6,7 1Department of Biomedical and Electronics Engineering, School of Engineering, University of Bradford, Bradford, United Kingdom; 2Bioengineering Research Group, Nanotechnology and Advanced Materials Department, Materials and Energy Research Center (MERC), Tehran, Iran; 3Department of Tissue Engineering and Regenerative Medicine, Faculty of Advanced Technologies in Medicine, Iran University of Medical Sciences, Tehran, Iran; 4Interdisciplinary Research Centre in Polymer Science and Technology (IRC Polymer), University of Bradford, Bradford, United Kingdom; 5Cellular and Molecular Research Centre, Iran University of Medical Sciences, Tehran, Iran; 6Department of Materials and Polymer Engineering, Hakim Sabzevari University, Sabzevar, Iran; 7Department of Biology, Faculty of Sciences, Hakim Sabzevari University, Sabzevar, Iran 

14.1  Introduction Ligament is a dense, soft, connective fibrous, musculoskeletal tissue that connects bones together, stabilizing the movement of joints. The main ligaments of the knee include anterior cruciate ligament (ACL), posterior cruciate ligament (PCL), medial collateral ligament, and lateral collateral ligament. There is a need for surgical restoration of impaired ligament tissue due to an increasing aging population, who desire better standards of living and the associated quality-adjusted life years. This need has encouraged medical research to establish, expand, and evolve approaches, which have confronted obstacles to tissue restoration [1]. Another demographics who are prevalently associated with ligament damaged are athletes, where the majority of ligament-based injuries reside in the knee, the ACL specifically [2]. ACL sport-related injuries are usually observed in “cutting sports” such as soccer, basketball, American football, hockey, skiing, netball, rugby, and tennis. These injuries (ruptures or tears) are normally noncontact injuries, when an athlete is required to change direction at substantial speeds with cutting and turning movements; the forces exerted by this movement overcome the resistive force of the ACL, leading to ruptures or tears. If the tensile strength of the ACL is exceeded, there is a rupture of the ligament due to dislocation of the femur from the tibia [3,4,57]. Iobst and Stanitski documented over 200,000 ACL injures, which are sustained every year within the United States of America. Surgical restoration of damaged ligament tissue may be required due to congenital deformation of tissue, accidental traumas, or pathological degradation [5–8]. The ACL and PCL have a weak intrinsic healing property, which is restricted by the absence of vasculature and the presence of synovial fluid [9,10]. Current ligament Handbook of Tissue Engineering Scaffolds: Volume One. https://doi.org/10.1016/B978-0-08-102563-5.00014-9 Copyright © 2019 Elsevier Ltd. All rights reserved.

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tissue replacement techniques include the use of autograft and allograft implants along with stem cell therapies, tissue-engineered reconstruction, and rehabilitation [11–14]. There is a need to retrieve successful implantation, biocompatibility, biodegradability, and high immune response of ligament tissue restoration for orthopedic and reconstructive surgery, for the advancement of tissue engineering principles, which will ultimately be aimed at bypassing current challenges. These challenges are currently being tackled through revised strategies such as implementing suitable biomaterials, chemical growth factors, isolated cells, stem cell therapy, and gene therapy to improve overall quality of life through aiding the natural healing process and functional tissue restoration [6,7,15,16].

14.2  Anatomy, physiology, and function of ligament This section presents the anatomy, physiology, functionality, cellular organization, structure, and composition of ACL tissue. The subsections depict the fiber bundle anatomy, bone–ligament interface, and the mechanical properties of ligament tissue. The ACL attaches the anterior-medial section of the tibia to the posterolateral portion of the femur, while being surrounded by the synovial membrane within the knee joint (Fig. 14.1). This structure of the tissue is made up of collagen molecules, which are hierarchically grouped into microfibrils, macrofibrils, collagen fibers, and finally into fascicles. The ACL and PCL are the main intraarticular ligaments and are essential for the support and movement of the knee [13,17–20]. The synovial fluid inhibits the diffusion of fluids across into the ligament as well as limiting fluid flow, which

Patella (kneecap)

Femur (thighbone) Lateral collateral ligament Anterior cruciate ligament

Posterior cruciate ligament Medial collateral ligament

Lateral meniscus

Fibula Tibia (shinbone)

Figure 14.1  Anatomy of knee—anterior cruciate ligament (ACL) and posterior cruciate ligament (PCL).

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ultimately results in the poor healing capabilities of damaged ligaments [21]. The PCL works in association with the ACL to maintain primary and secondary support and stability of the knee during internal rotation, anterior displacement, and tibial adduction [22,23]. The femoral insertion site of the ACL is at the posteromedial section of the lateral condylar region (semicircular in shape), whereas the tibial attachment of the ACL is wide and nonuniform in front of the intercondylar area, which is also the attachment location of the meniscus and PCL [17–20]. Cruciate ligaments comprise a combination of collagen, fibronectin, elastin, glycosaminoglycan, and laminin networks, all contributing to the structural properties of the cruciate ligament. The type I collagen helical fiber network defines the capacity of tensile strength of the ligament. Collagen fibers are arranged into intermedial, posterolateral, and anteromedial bundles, which create low tension and friction with regular movement. During motion, posterolateral bundles tighten at extension, whereas anteromedial bundles tighten at flexion, and at rotation of the knee, there is a twist of 90 degrees at the peripheral fiber bundles and ACL. ACL fibers are isometric in shape, enabling an equal distribution of load to the fiber bundles during flexion [17–20,24–27]. The research of Madey et al. on the sensory role of the ACL, suggests ligament nerve fibers relay sensory signals to the central nervous system regarding movement and positioning of the joint [28–31]. The size of the ACL in humans differs between genders; women often have a smaller ACL in comparison with males. The average length of the ACL is 29–37 mm with an average width of 9–12 mm [6,13,18,24,27,32,33]. The ACL fiber structure, orientation, interaction, and architecture comprises of 5–100 nm collagen fibers (diameter), which are arranged as bundles (1–20 μm diameter), which are further arranged into greater bundles named fascicles (100–400 μm diameter). Fascicles are arranged in parallel and helical patterns along the ACL, which is dependent on the distance from the bone interface. The variation of the nonparallel fibers enables different loading degrees, ultimately contributing to the sophistication, complexity, and unique qualities of the ACL [6,13,27,34,35].

14.2.1  Fiber bundle anatomy Fiber bundle anatomy has been defined differently by researchers. Odensten and Gillquist and Dorlot et al. [36,37] regard fascicles as not being able to be distinguished by anatomical groups, but Norwood and Cross, Amis and Dawkins, and Girgis et al. [38–40] have explained that two to three fascicle bundles/groups anatomically define cruciate ligaments. Cruciate ligaments are divided into fiber bundle, to better understand the complex nature of ligaments [41]. Changes in the distances between distinct fiber bundles form the basis of “length patterns” of ligaments [42–44]. What has been questioned is how the function of cruciate ligaments can be understood by varying the length of only a few fiber bundles. Blankevoort et al. showed that a ligament represented by two fiber bundles is not capable of stabilizing the knee joint in all positions of the knee, and from the cruciate ligament appearance, fiber bundles in multiple directions are present. More precise and detailed research needs to be performed to understand the insertion sites of the ligaments. Bundles are necessary to represent the major fascicle directions of the ligament; fiber bundle number and connection vary

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between all ligaments. Cruciate ligaments, which are composed of fascicles, have the capability of stabilizing motions of the knee in several directions. Understanding the complex functionalities of fascicle behavior in relation to tibia and femur sites of insertion and spatial orientation will potentially lead to a more accurate explanation of cruciate ligaments [1].

14.2.2  Ligament and bone interface Ligament attachment to bone in the human body is based on the transition of ligament to bone and vice versa. One attachment involves the ligament incidentally attaching to the bone through the periosteum; this method of attachment is called “periosteal insertion.” The tissue transition of ligament or tendon to bone is called “enthesis,” the first transition explained is the fibrous enthesis, made up of collagen fibers and fibroblasts (Sharpey’s fibers) [45–48]. The other tissue transition attachment is a gradual change of tissue from ligament to bone and is known as fibrocartilaginous enthesis. This enthesis is further subclassified into four specific regions. The first region within the midsection of the ligament has fibers parallel to the direction of load bearing, with dense packed fibrous connective tissue which results in high tensile strengths. This region is composed of fibroblasts and collagen. The second region is located within the transition of ligament to bone, consisting of fibrocartilage. The populace of chondrocytes increases, whereas there is a decrease in fibroblastic cells within the fibrocartilage. These cells are not aligned and are spindle-like in shape in contrast to the middle of the ligament. The cartilaginous transition to calcified fibrocartilage characterizes the third region, which is populated by a high density of chondrocytes with alignments that are not parallel to loading. The third region is known as “tide mark” and can be identified through histology. This region is at the border of transition from soft uncalcified tissue to calcified hard tissue [49]. The fourth and final region consists largely of osteocytes and is mainly bone. This region depicts the end of the fibrocartilaginous enthesis. Most replacement techniques cannot imitate the ligament-to-bone transition regions, opening a challenge in creating an ideal scaffold replacement for ligament tissue replacement [4,45,48].

14.2.3  Mechanical properties of the ligament The mechanical properties of the ligament provide the tissue with functional performance necessary for supporting the joint. The ligament constantly has loads and stresses applied on it, leading to deformation. There is a “crimping” factor to the fascicle microstructure that has a significant effect on the stress–strain curve for ligament at low loading and extension. This crimp mechanism enables the straightening of the ligament tissue to a small degree while being stressed at low loads before a significant load is applied. This phenomenon enables ligaments within their joint environment to act passively while undertaking daily activities, while specifically tightening up during substantial and excessive movement, all for the protection of the joint [1]. After an applied tensile load has been placed on the ligament, the crimping mechanism is stretched out before fibrils being loaded. After tensile load is removed, the

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ligament relaxes and then ligament elastin fibers act in restoring the original crimp structure. Because of the viscoelastic properties, tissue behavior is time-dependent, and these properties allow elastic deformation during tension at a physiological level. A characteristic called “hysteresis” occurs when a high frequency tensile load is applied to a ligament and it loses the ability to restore its original behavior, between loading and unloading cycles. Hysteresis can often lead to a change in mechanical property. A phenomenon known as “lag” occurs when there is a difference between tissue deformation and the signal of deformation over time. Recreating these properties of the ligament proves to be another challenge to be overcome in creating an ideal synthetic tissue substitute for ligament tissue replacement. These characteristics are crucial in the restoration and reconstruction of damaged ligament tissue [50]. Chen and Black measured standard physiological loading of human ACL, which ranged between 700 and 67 N, in relation to the applied stress during strolling, jumping, descending, and ascending a staircase. Fisher and Ferkel measured the cyclic loading that can be withstood by the ACL as approximately 300 N, 1–2 million times annually. The human ACL ultimate tensile strength varies from 2200 N to around 1730 N [6,24,27,51,52].

14.3  Conditions and injuries, diseases, and disorders of ligament tissue This section presents the conditions of damaged ligaments, possible injuries associated with ligaments, treatments, diseases, and disorders of ligament tissue. Ligament injuries occur when these tissues are placed in traumatic situations, which damage the structure of the tissue. When placed in an unfamiliar and uncomfortable position, due to an excessive force during motion, the result could lead to damage, tearing, and ultimately severance of the ligament (Fig. 14.2). Twisting of the knee is a frequent cause leading to overstretching or tearing of the ACL. When the ACL is injured, a popping sound may be heard and the respective limb may suddenly give out. Swelling and pain at the joint are symptoms of a damaged ligament; walking may induce pain leading the knee to feel unstable. Ligament injury could also be caused by certain factors such as the rate of impact of applied load, as well as the magnitude of the load [6,24,27]. Normal

Partial

Complete

Figure 14.2  Normal, partial, and complete injury to cruciate ligament.

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Partial tear, laceration, or damage to the ligament can easily be treated by elevating the damaged limb to permit fluid to flow from the joint position, compression of the impaired joint with an ice pack, and immobilization of the localized area. In the case of a fully torn ligament at the insertion site to the bone or the midsection of the tissue, surgical intervention is the recommended method of treatment. After surgery, recovery is prolonged and low impact exercises are recommended to strengthen the joint. Ligament injuries create disruptions in the balance between joint mobility and joint stability, which can lead to abnormal transmission of forces throughout the joint. This results in damage to other structures in and around the joint. Knees, hips, shoulders, ankles, elbows, and wrists are among some of the joints most commonly affected by ligament injuries. The ACL is the most commonly injured ligament of the knee. ACL damage is known to be caused by age-specific factors such as degenerative musculoskeletal disorders (found in individuals living in their 40s) and physical activities (in individuals in their 20s) [53,54]. The epidemiology shows gender predisposition, with males having a lower injury rate in comparison with females [55–57]. The majority of ACL and PCL injuries are known to occur in sports, such as football, rugby, skiing, basketball, soccer, and netball. Sports that fall into the category of “cutting sports” are known to be affiliated with ACL injuries. Sporting activities that involve athletes having to equip themselves with cleated footwear are more likely to experience ACL or PCL injuries. These cruciate ligament injuries often result in permanent joint disabilities and early osteoarthritis [19,27,58–60]. Ligament injuries are among the most common causes of musculoskeletal joint pain and disability encountered in primary practice today. Although there is a vast body of knowledge available regarding the structure and function of normal ligaments, understanding the structure and function of injured ligaments becomes more complicated due to the variability and unpredictable nature of ligament healing. If the well-being of the ligament is ignored, conditions such as tears in the meniscus, osteoarthritis, degenerative joint diseases, stress on articular cartilage, and joint instability would affect the joint [1,59,61]. Although ligament suffers from several diseases and disorders, it also contributes to several knee conditions. Some of these conditions are degenerative joint disorder, inflammation, osteoarthritis, and ligamentous laxity. Osteoarthritis is a common knee injury, known to be caused by wear and tear of joints. Researchers have realized that ligament damage can start the decline into osteoarthritis [62]. Degenerative joint disorder is an affliction leading to excess bone formation, degradation of cartilage, and grinding of bones at joints, which results in mechanical and joint instability. Inflammation, usually caused by trauma infection, leads to joint instability and hinders movement. Inflammation is not an abnormal reaction, but if ligament tissue remains in this phase for prolonged periods, the ligament tissue could lose functionality (elasticity) or hinder other tissue, such as articular cartilage and subchondral bone, from functioning, which ultimately leads to joint pain [63]. The laxity of damaged ligaments is a major cause of chronic joint pain during degradation of tissue. Once the ligaments cannot function, the muscles and bone compensate for the lack in ligament elasticity. This leads to pain, and later the whole joint attempts to compensate for the damaged faulty ligament.

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The compensation reactions aid in stabilizing the joint, but, unfortunately, the next stage is osteoarthritis [64]. Without proper care, ligament laxity promotes subchondral bone degradation and then osteoarthritis. More research needs to go into the understanding of how ligament-joint disorders are initiated, which will fortunately guide studies toward a strategy to improve ligament healing or delay the effects of degenerative joint disorder [65].

14.4  Ligament healing This section outlines the wound healing ability of ligament tissue, explaining the four phases involved. The healing capability of ligament is influenced by the mobilization of the ligament and the age of the victim at injury. In addition, being enveloped in synovial fluid and having poor vascularity, the ligament healing capacity is hindered. Mobilization permits ligaments to respond to mechanical stress during the remodeling stage of the tissue through mechanotransduction. Ligaments that are immobilized after injury fail to receive mechanotransduction stimuli, making remodeling weaker and less healthy. A reduction in the metabolism of cells and an alteration in collagen structure are caused by aging, which ultimately results in a fall of mechanical properties [9,10]. The healing process of ligament is divided into four phases: inflammation phase, matrix and cellular proliferation phase, remodeling phase, and maturation phase [23]. Immediately after injury, the inflammation phase begins and lasts for about 72 h, with synthesis of the matrix and high cellular activity. Early in this stage, blood cells and inflammatory cells such as monocytes, macrophages, polymorphonuclear leukocytes, lymphocytes, and phagocytose necrotic cells migrate to the site of injury. The next phase, matrix and cellular proliferation, begins at the point where the majority of the cell population switches from inflammatory to fibroblastic cells. In this phase, fibroblasts dominate the injury site by actively synthesizing extracellular matrix (ECM), while mast cells and macrophages are still present. The proliferation period lasts for approximately 6 weeks as fibrous capsule and highly cellularized scar tissue (greater surface area) forms throughout the wound, aiding vascular development. At the transition from proliferation to remodeling phase, there is a large amount of collagen type III within the healing tissue in relation to collagen type I. Collagen type I has a greater resistance to tensile loads along the ligament than collagen type III, as type III is usually correlated to disorganized collagen fibrils. There is a decline in vascularity, fibrous capsule, and cell density during the remodeling phase transition, which occurs over a few months. The ratio of collagen type I to type III becomes greater in comparison with collagen fibril alignment organized along the ligament’s primary axis. The maturation phase of the ligament tissue occurs from months to years, with slow maturation of ligament scars, and abnormalities and deformation persist due to mechanical and environmental elements [13,18,22,23,35]. The unpredictability of ligament healing can be due to the dramatic physiological and structural changes that ligaments sustain because of injury, as well as the complex and dynamic cellular processes that occur during healing. These processes create alterations in the biology and biomechanics of the injured ligament, leading to inadequate healing and tissue formation that is inferior to the tissue it replaces.

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14.5  Scaffold design and fabrication techniques This section outlines design fabrication methods in tissue reconstruction and some of the strategies that are used in the fabrication of ligament tissue. An ideal tissue-engineered scaffold provides a foundation that directs cell regeneration, tissue formation, mass transportation, mechanical support, and cell activities such as differentiation, proliferation, viability, and migration. The transfer of load and facilitation of tissue reconstruction are also mediated by the scaffold [66]. Ligament tissue scaffolds should be designed to promote a complex multicellular environment, which ultimately supports tissue maturation and development [67]. Ligament tissue constructs would require cell-containing scaffolds to maintain integrity. An ideal cell type would be fibroblasts, with the function of recreating ECM. This can be performed either by seeding cells onto scaffolds or nurturing the cell–scaffold complex within a bioreactor. Polymer biomaterials and organic biomaterials are used in common joint-related tissue engineering procedures [68]. Synthetic polymer materials are commonly used in the synthesis of ligament scaffolds due to their reproducibility and relative ease of production [69,70]. There are a few techniques/processes used in the fabrication of ligament scaffolds. Some of these methods include the following: • Melt spinning • Solvent casting • Gas foaming • Freeze-drying • Three-dimensional printing • Polymerization • Hydrogel solution mixing • Cross-linking • Electrospinning [71,72]

Hot pressed melt spun fiber technique involves the use of hot-pressing methods to fabricate a scaffold with optimal stress and water absorption, porosity, diameter, and physical properties [72]. Hydrogel synthesis can be carried out using different methods; a common procedure is solution mixing/acid dilution, whereby a potential biomaterial is dissolved into an acid [73]. Electrospinning is a common technique used in synthesizing ligament tissue constructs, a process by which a potential difference is applied to a polymer solution as it is extruded from a syringe and onto a collector plate [74–76]. This creates fine aligned nano/microscopic fibers with appropriate pore sizes for cell seeding [71,77]. Researchers have also tried to incorporate different fabrication techniques and technologies as well as strategies to optimize the design, characteristics, and structure of ligament tissue constructs. In current research, electrospinning techniques are often modified with different strategies, and some of these combined techniques include cell sheet technology, stem cell technology, hydrogel synthesis, growth factors, vapor phase polymerization, freeze-drying, and electrospraying [73,78–84].

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Some important scaffold characteristics include the following: • Mechanical competence and stability • Scaffold composition • Microstructure • External geometry • Chemical properties • Surface to volume ratio • Macrostructure • Degradation characteristics • Porosity and interconnectivity

Tissue scaffold constructs can be differentiated by chemical composition (organic or nonorganic materials) or state (e.g., hydrogels (gel-like) or solid state). Macrostructure describes the interior structure and exterior geometry of a scaffold. A well-fabricated scaffold mimics the native ECM environment, with precise macrostructure, and promotes tissue formation. The geometry of scaffolds is classified into pore and bulk geometry. A porous scaffold promotes interconnectivity, which nurtures the supply of nutrition, and a hydrodynamic microenvironment is mediated by high interconnectivity. The pore size of a scaffold controls the scaffold fluid flow and vascular ingrowth, ultimately improving tissue replacement strategies. Optimal scaffold porosity depends on what tissue is being engineered: bone tissue formation relies on porosity, whereas cartilage has no benefit from porosity [85]. Pore sizes need to be small enough to allow the cells to sit on scaffold material. Cellular attachment and migration into porous scaffolds are supported by high surface area to volume ratios. A compromise between mechanical stability and surface area needs to be found because internal surface area and pore diameter are linearly related. Chemical properties should promote the transportation of proteins within native tissue and be released in an ordered manner. Mechanical properties, such as mechanical strength, contribute to resistance to the physiological mechanical environment during load-bearing tissue regeneration within implantation site [85]. Scaffold degradation must retain integrity as and before the new native tissue fully regenerates. Scaffold degradation needs to happen in parallel with the formation of new tissue, as well as being manipulated appropriately for easy replacement of the scaffold material by the new native tissue. Degradation characteristics should be reviewed before implantation, to prevent a need for extraction for any needed correction [86].

14.6  Biomaterials available for ligament tissue engineering This section details the purpose of incorporating biomaterials in tissue reconstruction, requirements for an ideal biomaterial, and natural and synthetic biomaterials used in fabricating ligament tissue constructs with some respective properties.

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Biomaterials are an essential component of tissue reconstruction regarding applications of medical technology, while being one of the building blocks of tissue repair. Biomaterial design requires a scientific understanding of challenges such as cytotoxicity, tissue/host implant reactions, sterility, storage, and biocompatibility, to fabricate ideal tissue-engineered constructs for replacement. Focusing on biomaterials improves on the quality of the engineered construct. Biomaterials affect these characteristics in ways that will determine the outcome of the fabricated construct [51,87–89]. Biomaterials for ligament tissue are tailored toward the requirements of the application (based on functionality). Some of the general requirements of an ideal biomaterial include (1) functional ability: preferred biomaterials provide the desired function of the fabricated tissue construct through economical fabrication and synthesis techniques. (2) Biocompatibility: biomaterials should not stimulate a strong tissue rejection response from the host. An immediate inflammatory response is anticipated and considered necessary to the wound healing process, although a constant immune response may signify an incompatibility with scaffold or presence of tissue necrosis. (3) Sterilizability: biomaterials must be able to undergo the required sterilization protocols and not produce hazardous reactions in response to the sterilization techniques. Possible sterilization techniques include steam autoclaving, ethylene oxide, and gamma radiation. (4) Manufacturability: scaffold materials will have to be manufactured to a functional level and be able to be stored before commercial use [90,170,171]. Biomaterials are generally classified into natural and synthetic biomaterials. In some cases, both natural and synthetic biomaterials are used to create tissue constructs; these are called semisynthetic materials. Synthetic biomaterials are further divided into polymeric, composite, ceramic, and metallic biomaterials, whereas natural biomaterials are further divided into soft and hard tissue [51,87]. Biomaterial mechanical properties are simply defined by the material’s ultimate tensile strength (its ability to bear load before failure), elastic modulus (the stiffness of the material), fracture toughness (resistance to crack propagation), and the elongation until failure (the amount of strain withstood before failure) [90]. The normal biomaterials used in repairing ligaments are natural, synthetic, and composite materials. Some of these materials include the following: • Collagen, • Silk, • Fibrin, • Poly (lactic acid), • Polyglycolide acid (PGA), • Poly (lactic-co-glycolic acid) (PLGA), • Polymethyl methacrylate (PMMA), • Ultrahigh molecular weight polyethylene, • Polypropylene (PP), • Polyethylene terephthalate (PET), • Polytetrafluoroethylene (PTFE), • Polyurethane (PU), • Poly (2-hydroxyethyl methacrylate) (PHEMA),

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• Poly (ethyl acrylate) (PEA), • Kevlar-49, • Dacron, • Polyether ether ketone (PEEK), and • Gore-Tex [26,90,91].

Researchers have combined synthetic materials and autogenous tissue to create ligament tissue construct, for example, Ambrosio et al. [69] established a ligament prosthesis by fortifying a PHEMA hydrogel with wound helical PET fibers and revealed that ligament mechanical behavior (dynamic and static) can be duplicated [69]. By joining PEA with Kevlar-49 fibers, natural ligament stress–strain behavior can be mimicked to a certain degree; this was reported by Pradas and Calleja [90–92,174].

14.6.1  Natural materials This subsection details various natural materials used for the reconstruction of ligament tissue along with their characteristics. These materials are derived from natural resources, and some common natural biomaterials are collagen, silk, fibrin, chitosan, and gelatin. Silk is a natural biomaterial used in tissue fabrication through bio 3D printing as bioink for hydrogels. Silk as a biomaterial has the properties for supporting cell attachment, multilineage differentiation, tissue formation, and cell proliferation [78,93]. Hyaluronic acid (HA) is a biomaterial acquired from joint synovial fluid with water solubility, high molecular weight, and lubricious characteristics. HA solution is used in intraocular surgery (lens) for securing the iris and cornea during operation. HA is compatible with the eye because it can be found within the aqueous humor. Commercial HA solutions used in ophthalmic surgery have an average molecular weight ranging from 8.0 to 0.5 × 106 g/mol and average viscosity ranging from 2.2 × 104 to 7 × 106 mPas [94]. HA can be cross-linked with different reagents, due to its functional group flexibility, and this brings up a potential for applications in hydrogel drug delivery [95]. Starch is a polysaccharide that is the basis of carbohydrates in a balanced human diet. Amylopectin is a branched polymer with a linear backbone (molecular weight ranging from 105 to 5 × 105 g/mol), with 4% branching. Amylose on the other hand is a linear polymer with molecular weight ranging from 1 to 5 × 105 g/ mol. These monosaccharides are the constituents of starch. Starch is insoluble in water by nature, although research has shown starch can be soluble when dissolved in dilute hydrochloric acid. Soluble starch is used in the application of blood plasma extenders. The globular protein albumin has 17 disulfide bridges and molecular weight 66,200, making up to half of the human plasma [96]. The presence of several amino and carboxylate groups provides cross-liking reactions bringing about a wide variety of mechanical behavior. Albumin structural properties have been implemented into optimizing drug delivery and tissue substitute applications [91]. Gelatin is a biodegradable protein, acquired by hydrolyzing collagen type I with the use of bases and acids. Collagen loses molecular weight, and its tertiary triple helix structure is disassembled to yield soluble gelatin. Gelatin has the property of undergoing gelation with a drop-in temperature. Gelatin is applied in capsule biomaterials and coating tablets

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for the property of controlled drug release. Fibrin is another biodegradable monomer found within the blood, with a role in forming blood clots during wound healing. This monomer is a protein formed by fibrinogen, used in creating ECMs and bioadhesives for tissue engineering purposes. The gel formation is orchestrated by the enzyme thrombin, a calcium solution and fibrinogen, and varying the concentrations controls the speed of gelation. Fibrin is used in tissue scaffolds and soft tissue applications for its fast biodegradability due to natural tissue enzymatic activity through fibrinolysis arising from tissue plasmin [91]. Chitin and chitosan are polysaccharides derived from the shells of shellfish and insects. Deacetylation of chitin produces chitosan, using high temperature basic solutions. Chitosan is 70%, with apertures to 350 μm to support cell migration (differentiated cells’ size ranging from 20 to 120 μm) and, more specifically, ossification [44]. Additionally, as in many studies outlined here, vascularization is not conventionally evaluated in in vivo works, limiting the understanding on the remodeling process. Blood supply is paramount to the damaged tissues to deliver nutrients and growth factors and remove waste and supports continuous healing, development of nerve tissue, and ultimately remodeling [45]. The incorporation of microchannels deep into the scaffold (up to 100 μm) may support the development of harversian canals for microvascularization and angiogenesis to occur. Scaffolds materials choice is a significant concern and is still dependent on the clinical case and the size of the construct needed. Metals have been shown to have the required mechanical strength yet have been found to contribute to future degeneration of local tissues, or lead to fragmental displacements around the body. Some materials are not able to support osteointegration, whereas other materials such as bioglass or ceramics, containing calcium or other mineral components, are able to support ossification and more rapid new bone formation [46]. Scaffolds that incorporate hydroxyapatite appear to support mineralization of the scaffolds at an increased rate compared with those without HAP [43,47]. The use of hydrogels provides the need for the high water content found in bone matrix, which carries various growth factors and cells [39]. However, hydrogels are intrinsically soft, notoriously known to lack the mechanical strength required for the mechanical impacts exerted on bone. Therefore, this type of hydrogel requires rapid support of new bone formation, increased mineralization, and increased bone density, as the gel resorbs. Development of a novel hydrogel may help support better the dynamic niches required for cell behavior and to ensure that regeneration is clinically successful not only in small defects. Use of growth factors to date in scaffolds has shown support in regeneration process, yet at this time they do not have a significant impact on regeneration, which therefore implies it is not a practical or finically sensible option. Nanoparticles, used as carriers for drug delivery or used to control scaffold resorption, require the right biomedical size, not too large to cause granulomas, requiring splenic breakdown, small enough to be renally excreted, and not too small that they can cause cytotoxicity [36]. Generally, nanoparticles are well characterized with some work already in clinical settings, such as silver nanoparticles in medical dressings and wound healing. Nanoparticles combined with various scaffold materials such as HAP can control the resorption of different scaffold components at various rates [44]. For regenerative purpose of mandibular smaller defect sizes 2 cm2, within zones 1 and 2 The defect of the entire orbital floor and the medial wall, extending into the posterior third (zone 3)

Bony ledge preserved at the medial margin of the infraorbital fissure Missing bony ledge medial to the infraorbital fissure Missing bony ledge medial to the infraorbital fissure

With permission from L. Dubois, S.A. Steenen, P.J.J. Gooris, R.R.M. Bos, A.G. Becking, Controversies in orbital reconstruction-III. Biomaterials for orbital reconstruction: a review with clinical recommendations, Int J Oral Maxillofac Surg 45 (1) (2016) 41–50.

Figure 19.1  Tomographical image (coronal plane) of a patient suffering from a right orbital floor fracture, vertical elongation of the right orbit, and a reduction in the size of the right maxillary sinus. (1) Orbital cavities; (2) maxillary sinuses; (3) fracture area. With permission from F. Baino, Biomaterials and implants for orbital floor repair, Acta Biomater 7 (9) (2011) 3248–3266.

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Table 19.2  The characteristics required for an ideal orbital reconstruction material. Stability and fixation

Contouring and handling

Biological behavior

Drainage Donor site morbidity Radiopacity Availability and cost-effectiveness

• Strong

enough to support the orbital content and related forces to be stable and retain its shape once manipulated • No deformation (sagging of material into maxillary sinus) under pressure load • Stable over time • The possibility of being fixed to surrounding structures • Restores adequate volume to treat enophthalmos, diplopia, and motility disorders • Easy to shape to fit the orbital defect and regional anatomy/ malleability • Sufficient in three-wall fractures • No sharp edges and smooth surface • Biocompatibility: No infection/extrusion/migration/foreign body reaction • Chemically inert, nonallergenic, noncarcinogenic • Durable with minimal resorption • Osteoinductive/osteoconductive • High tissue incorporation but readily dissected in implant removal during secondary reconstruction • Spaces within the implant to allow drainage of orbital fluids • Does not increase the surgical complication rate/donor site morbidity (pain, swelling, etc.) • Radiopaque to enable radiographic evaluation without artifacts • Readily available in sufficient quantities Acceptable costs • Ability

With permission from L. Dubois, S.A. Steenen, P.J.J. Gooris, R.R.M. Bos, A.G. Becking, Controversies in orbital reconstruction-III. Biomaterials for orbital reconstruction: a review with clinical recommendations, Int J Oral Maxillofac Surg 45 (1) (2016) 41–50.

use of xenografts (e.g., porcine collagen) has been tested for the treatment of this type of defects [14]. In recent years, considerable efforts have been carried out to propose alternatives to transplant materials, the properties of which can be finely designed and tailored according to the specific application. This chapter provides a picture of the current state-of-the-art related to orbital wall repair implants and highlights the potential of recent strategies based on tissue engineering approaches.

19.2  Transplant materials Up to now, several materials of biological origin have been evaluated and applied to the restoration of orbital floor defects (Fig. 19.2). These materials include a wide range of substances harvested from human donor or animal species, as described in the following sections where the pros and cons of each option are shortly discussed.

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(b)

(f)

(c)

(g)

(d)

(h)

Figure 19.2  Some examples of biomaterials/implants used for repair and regeneration of orbital wall defects: (a) iliac crest autograft; (b) bioactive glass plates with their corresponding “kidney-shaped” and “drop-shaped” stainless steel templates; (c) titanium mesh on a solid orbital model; (d) porous polyethylene sheet (Medpor); (e) poly(l-lactide) implant; (f) poly(L-lactic) acid/polyglycolic acid composite implant (Lactosorb panel); (g) polyglactin 910/polydioxanone patch (Ethisorb); (h) upper and lower side of titanium/polyethylene composite implant (Medpor Titan). Reproduced with permission from F. Baino, Biomaterials and implants for orbital floor repair, Acta Biomater 7 (9) (2011) 3248–3266. (d and h) Courtesy of Porex Surgical.

19.2.1  Autologous bone The use of autologous grafts has some superiority for the treatment of orbital floor defects compared with the other transplant materials such as high biocompatibility as well as lack of immunogenicity and disease transmission [15]. Autologous bone also retains proteins and growth factors, which are crucial to stimulate tissue regeneration. Moreover, the inherent strength, rigidity, osteoconductivity, and osteoinductivity are counted as merits of this type of grafts [16]. The cranium, iliac crest, and ribs are considered the commonplaces of harvesting the bone grafts [17]. However, other sites such as mandibular symphysis and more recently the olecranon are also examined as donor sites for harvesting the bone required for the treatment of orbital floor defect [18,19]. Among various bone grafts, the anterior wall of the maxilla is considered a simple graft, which is used for many years for the orbital floor restoration. This graft takes advantages of easy accessibility, characteristics resembling those of the orbital floor, and rapid harvesting procedure [20]. Over the last four decades, the approach based on using autologous bone has been conducted on a large number of patients after being first described in a case series of three patients requiring orbital floor repair in 1966 [21]. Although a large number of clinical studies have confirmed the efficacy of bone autografts as the preferable substitutes of choice in orbital floor restoration [22–24], some surgeons have a negative attitude against the idea of using such autografts because of some specific drawbacks such as donor-site morbidity [25].

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19.2.2  Cartilage autografts This type of grafts is currently obtained from some specific places in the human body, including ear [26] and nasal septum [27–29]. Easy harvesting procedure and shaping of cartilage autografts, as well as the capability of providing long-term support to the surrounding tissues with minimum morbidity at the donor site, are counted as advantages of this grafts as compared with autologous bone [16]. As an illustration, Ozyazgan et al. treated 10 patients with traumatic orbital wall defects (size: 100– 400 mm2) by using conchal cartilage [30]. The grafts have been taken from the posterior and the anterior regions of the external ear without any hypertrophic scar or keloid development. Postimplantation results showed that 9 of 10 patients have no limitation in eye movement and no signs of diplopia and enophthalmos. The authors concluded that conchal cartilage could be successfully used for the repair of defective orbital wall fractures that are not oversized ( PLLA; cell proliferation of PLLA/Ts 10%), CanFite Biopharmaceuticals and lesser (